Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1 -- 3884 2022-11-24 09:04:23 |
2 format correct -3 word(s) 3881 2022-11-24 09:31:05 |

Video Upload Options

Do you have a full video?

Confirm

Are you sure to Delete?
Cite
If you have any further questions, please contact Encyclopedia Editorial Office.
Abd-Elgawad, M.M.M. Xenorhabdus spp.. Encyclopedia. Available online: https://encyclopedia.pub/entry/36300 (accessed on 15 June 2024).
Abd-Elgawad MMM. Xenorhabdus spp.. Encyclopedia. Available at: https://encyclopedia.pub/entry/36300. Accessed June 15, 2024.
Abd-Elgawad, Mahfouz M. M.. "Xenorhabdus spp." Encyclopedia, https://encyclopedia.pub/entry/36300 (accessed June 15, 2024).
Abd-Elgawad, M.M.M. (2022, November 24). Xenorhabdus spp.. In Encyclopedia. https://encyclopedia.pub/entry/36300
Abd-Elgawad, Mahfouz M. M.. "Xenorhabdus spp.." Encyclopedia. Web. 24 November, 2022.
Xenorhabdus spp.
Edit

Xenorhabdus bacteria, as pesticidal symbionts of the entomopathogenic nematodes Steinernema species, can contribute to this solution with a treasure trove of insecticidal compounds and an ability to suppress a variety of plant pathogens. As many challenges face sound exploitation of plant–phytonematode interactions, a full useful spectrum of such interactions should address nematicidal activity of XenorhabdusSteinernemaXenorhabdus complex or Xenorhabdus individually should be involved in mechanisms underlying the favorable side of plant–nematode interactions in emerging cropping systems. Using Xenorhabdus bacteria should earnestly be harnessed to control not only phytonematodes, but also other plant pests and pathogens within integrated pest management plans. 

biocontrol Steinernema pest and pathogen management

1. Identification, Taxonomy, Lifestyle, and Diversity of Xenorhabdus spp.

1.1. Their Identification/Taxonomy

As Xenorhabdus and Photorhabdus bacteria (Enterobacterales: Morganellaceae) are phylogenetically close, it is not surprising that at first they were of the same genus [1]. Thus, two bacterial species in this genus (that is, Xenorhabdus nematophila (type species) and Xluminesces) including symbionts of the nematode genera Steinernema and Heterorhabditis, respectively, were exclusively present until 1993 [1]. Yet, the important variations in the phenotypic and molecular traits could distinguish Xnematophila from Xluminesces, leading to the transfer of all symbionts related to Heterorhabditis into Photorhabdus as a new genus with the type species Photorhabdus luminescens [2]. Before splitting into the two genera, some phenotypic features and symbiotic properties were utilized to characterize Xenorhabdus and Photorhabdus bacteria as two recognized groups; Pluminescens distinctly had a DNA relatedness group unlike all Xenorhabdus strains with significant variations in phenotypic traits [3]Xenorhabdus bacteria are obviously separated from Photorhabdus species/strains by the 16S rDNA signature sequences [4]. Yet, both Photorhabdus luminescens and Xnematophila received much research work, due to being type species of their two genera, with high insecticidal activities and a global distribution.
Interestingly, the genus Xenorhabdus still has more homogenous species than Photorhabdus, but the former genus possesses a higher number of species than the latter. For example, Sajnaga and Kazimierczak [5] reported 26 Xenorhabdus spp. versus 19 Photorhabdus spp. This is possibly due to higher number of the current Steinernema species (the mutualistic partner of Xenorhabdus spp.) than Heterorhabditis spp. (the mutualistic partner of Photorhabdus spp.), i.e., >100 Steinernema spp. but >20 Heterorhabditis spp. [6]. Admittedly, there are other undescribed species related to both Xenorhabdus and their Steinernema partner which are recognized or are likely to become recognized soon. In this respect, the number of their close relatives, Photorhabdus species, has recently doubled, from four to twenty in the last few years [7].

1.2. The Lifestyle and Diversity of Xenorhabdus spp.

Basically, the bacterial species in the genus Xenorhabdus have a mutualistic relationship with the entomopathogenic nematodes (EPNs) of the genus Steinernema. The bacteria live symbiotically in the specialized intestinal vesicles of Steinernema. The two partners naturally form an antagonist sharing mainly against their insect hosts. The EPN-third-stage infective juveniles (IJs) conserve the bacteria in their body from the outer environmental stresses until these IJs release them within the insect body. In addition, after EPN infection and depleting the host resources, the IJs vector the bacteria from one susceptible host to another. In turn, Xenorhabdus spp. generate antimicrobial compounds and secondary metabolites into the insect. These materials can not only kill the insect host and prepare the contents of its body to feed the nematodes for their development and reproduction, but also protect the insect cadaver from soil scavengers and saprobes [8]. During their feeding, the nematodes also swallow the bacteria in order to grow and reproduce.
Steinernema–bacterial symbiont specificity and their coevolution have been thoroughly studied for many involved axenic (free of bacteria) and monoxenic (having a Xenorhabdus species) Steinernema species [5]. While a Steinernema species can presumably set up symbiosis with only one species of Xenorhabdus, any of numerous Xenorhabdus species are able to associate with several Steinernema. On the contrary, the symbiotic HeterorhabditisPhotorhabdus associations are more adaptable as many species of each partner can engage in symbiotic relationships with multiple species of the other partner [9]. These facts have recently been reviewed and backed by plenty of data [10]. However, the mechanisms underlying these relationships remain to be clarified [5]. These associations do not negate the fact that the robust specificity that favors symbionts with the most useful attributes facilitates effective transfer of such a nematode–bacterium pair from a susceptible insect pest to another. Generally, Sajnaga and Kazimierczak [5] concluded that there is a possibility of horizontal transfer of Xenorhabdus bacteria between different Steinernema species, relying on the species of XenorhabdusSteinernema pair used. However, such switching in the bacterium–nematode pair may have its pros and cons. On the positive side, associations of Steinernema species with new Xenorhabdus partner may validate colonization of novel niches or expand one by offering considerable fitness benefits [11][12]. These favorable results may occur when the introduced bacteria/symbiont is closely related to its native Steinernema species. On the negative side, the Xenorhabdus partner switching frequently has a harmful effect on the Steinernema host in terms of a reduction in their fitness, reproduction, and symbiont carriage as well as virulence. For example, Xenorhabdus bovienii is the native symbiotic bacteria of Sfeltiae. However, using Xnematophila strain HGB315, not the native symbiont of Sfeltiae, this nematode species developed and turned into gravid much faster at approximately 4 days on Xbovienii (versus 5–6 days on Xnematophila HGB315) post-seeding [13]. These detrimental outputs are usually associated with non-cognate and phylogenetically distant symbionts [5]. Eventually, researchers and stakeholders should be aware of the fact that the Steinernema host diversity substantially impacts coadaptation between various XenorhabdusSteinernema partners [14][15] for their further wise application. In this respect, Tailliez et al. [16] could identify two main groups of Xenorhabdus strains based on phenotypic analysis. A group included bacterial strains that can commonly grow at 35–42 °C, while the other group included Xenorhabdus strains that grow below 35 °C. Hence, Xenorhabdus bacteria may be adapted to temperate, subtropical, or tropical regions. They are also differently impacted by the growth temperature of their Steinernema host [16]. Moreover, the wide host range of their nematode host along with their major attributes can prove their diversity and global distribution [17] as well as give opportunities to familiarize stakeholders with the potential usage of these symbiotic bacteria [6][8]. With the global spread of the SteinernemaXenorhabdus complex, recent references, e.g., [7], still indicate that EPNs are not discovered in Antarctica. The long-established realization regarding the species-specific characterization for the dyad SteinernemaXenorhabdus complex as partners for the mutualism is still effective. Thus, it can bode well for more investigations concerning their distribution in diversity and space [18]. Moreover, current research efforts have been focusing on optimizing methods and techniques for EPN surveys and extraction [4][5] to detect novel species/strains that bode well for effective and safe biocontrol of pests and pathogens and adaptation to local conditions. Therefore, it can be mostly presumed that the distribution and diversity of the EPN species is only an artefact of the linked sampling efforts [7]. However, growing interest is mainly dedicated to these bacteria when applied to suppress pests and pathogens independently, i.e., without the EPN partners [8][19][20][21][22][23].
Similar to their near relatives, Photorhabdus species, all the species of Xenorhabdus are exclusively linked symbionts to the Steinernema spp.–IJ stage [2]. The exception of Photorhabdus, materialized in P. asymbiotica as a human pathogen in addition to infecting insects [24], is not found in Xenorhabdus. None of the Xenorhabdus bacteria were found in free-living order in nature; therefore, they had formerly boosted doubts concerning their ability to survive and infect pathogens/pests without the EPN partner.

2. Pathogenicity of Xenorhabdus spp.

2.1. Magnitude and Profile of Pathogenicity

Traditionally, various SteinernemaXenorhabdus partnerships, to attack and kill numerous arthropod pests, have been marketed and utilized as biocontrol agents [6], with increasing ambition for their expansion to prepare them for reliable alternatives in pest management and plant protection [24][25][26][27][28]. In the original status of the natural SteinernemaXenorhabdus complex, Xenorhabdus host range is surely limited to the ability of the IJs to locate and infect the host. This is a prerequisite for the development and multiplication of Xenorhabdus spp. to achieve high levels of cells within the host. Factually, both mutualists, Steinernema and Xenorhabdus, can generate bioactive compounds to kill the invaded host [13][20]. Thereafter, the bacterial cells can modify the insect host tissues to become a nutrient diet needed for the IJ development and multiplication. Hence, the pathogenicity depends on the bacterial activity and growth. Thus, the Xenorhabdus rate of growth is tightly related to the time needed to kill the insect host. Clearly, Xenorhabdus spp. are quite virulent pathogens of a broad range of pests/pathogens including insects, fungi, bacteria, protozoa, and nematodes [20][21].
Discovering the competency of Xenorhabdus bacteria to live in fresh water and in soil for 6 days has surely opened a new avenue with fixed timeframe for their further biocontrol usages, apart from their mutualistic Steinernema [27]. Consequently, various formulations and techniques, fundamentally comprising just the bacteria or/and bacterial metabolites, have been used [6][8][20][21][28][29][30][31][32][33][34]. In this regard, boosted pathogenicity islands of the Xenorhabdus chromosome, having numerous genes that encode various antibiotics, insecticidal protein toxins, enzymes, and bacteriocins, were investigated [8][20][35], and more are still to be further characterized, e.g., [21][28][36][37]. For instance, only one Xenorhabdus strain may generate a variety of antifungal and antibacterial compounds. Some of its compounds are active against insects, protozoa, nematodes, and cancer cells, too [20]. All tested Xnematophila strains showed insecticidal activity against representative pests of three insect orders; the cabbage white caterpillar Pieris brassicae (Lepidoptera: Pieridae), the mosquito larva Aedes aegypti (Diptera: Culicidae), and the mustard beetle Phaedon cochleariae (Coleoptera: Chrysomelidae) [34]. In this study and others [19][23][38][39], an important note is the variation in the abilities of different Xenorhabdus species/strains to kill/inhibit the growth of the intended pest or pathogen. These variations are based either on the ability of each Xenorhabdus species/strain to generate effective metabolites or the relative susceptibility/tolerance of the targeted pest/pathogen. These differences are found not only between Xenorhabdus species/strains, but surely exist to varying degrees between bacterial species/strains belonging to different genera. 
An obvious technique to circumvent the lack of appropriate efficacy of Xenorhabdus species/strain and/or to increase its potency is to introduce other antagonists in combination with Xenorhabdus bacteria and/or their bioactive compounds. This approach can establish and boost the efficacy of the introduced organism, too. Clearly, synergistic activity to kill the beet army worm Spodoptera exigua could be obtained by mixing growth media supernatants of Xenorhabdus bacteria with B. cereus or B. thuringiensis spores. In this case, while the supernatant of Xenorhabdus bacteria could exert its impact on the insect hemocoel, the Bacillus cells were able to perforate the insect midgut epithelium [40][41]. Later, Eom et al. [41] could develop a “dual Bt-plus” product by mixing B. thuringiensis (Bt) spores and culture broth of X. nematophila (Xn). Although this product demonstrated high toxicity, it has also some modification to widen its efficacy against a diverse insect pest spectrum. Their tests centered on increasing “Bt-Plus” toxicity against a semi-susceptible insect, Sexigua, via adding Xn metabolites. Given the fact that Xn metabolites, benzylideneacetone (BZA) and oxindole (OI), can boost the Bt insecticidal activities, adding each of them (OI or BZA) could significantly enhance Bt-Plus pathogenicity. Moreover, when the freeze-dried Xn culture broth was included into Bt-Plus, a much smaller amount could suffice to raise the toxicity relative to the amount of BZA or OI. High-performance liquid chromatography analysis revealed that there were more than 12 unidentified Xnematophila metabolites in Xn culture broth. Therefore, they [41] proposed that there are other potent biological response modifiers in Xnematophila metabolites, not solely OI and BZA. Likewise, a Xenorhabdus species could induce high mortality of S. exigua third-instar larvae but its pathogenicity was much less for the fifth-instar larvae. Seongchae and Yonggyun [42] speculated that the high mortalities in the third-instar larvae were due to antibiotic activity against B. cereus, a gut symbiont needed to optimize S. exigua development. To enhance the Xenorhabdus species pathogenicity in the fifth instar, the bacteria should be delivered into the hemocoel. Thus, the authors utilized B. thuringiensis aizawai (Bt) as a synergist to back entry of the bacteria from the insect gut lumen into its hemocoel by disrupting the S. exigua gut epithelium. As a result, the applied bacterial mixture was highly synergistic against the S. exigua fifth-instar larvae. This synergism was proved via the successful infection of X. sp. or Bt in the insect hemocoel. Therefore, Xenorhabdus bacteria can be used to kill S. exigua by oral treatment in a mixture with Bt [42].
Usually, re-extraction of the Xenorhabdus bacteria from the insect cadavers and comparison with the standard (original) culture can assure Koch’s postulates [20][43]. Although the obtained data confirmed the direct toxicity of the bacteria to definite insect species in nature, e.g., [8][34][43], particular Xenorhabdus bacteria may have wide host range of insect pests. For example, 122 strains of symbiotic bacteria associated with 23 EPNs were gathered from various Chinese localities [44]. These extracted strains displayed oral growth inhibition and/or insecticidal activity against the Ostrinia furnacalis larvae. One of the strains, however, Xenorhabdus sp. SY5, with determined partial toxin gene sequence, exhibited strong insecticidal activity to a variety of economically significant agricultural pests. Their species comprised Plutella xylostellaOstrinia furnacalisTenebrio molitorS. exigua, and Mythimna separata. The strain isolated from Steinernema sp. SY5 appeared to have seven purified toxins based on DEAE-52 column chromatography. These toxins exhibited, to a certain extent, growth inhibition and/or insecticidal activity to these insect species. The authors [44] stressed the high virulence of this strain as a potential asset for biological pest control.
It is likely that the arsenal of Xenorhabdus spp. still possesses much that has not been discovered yet, for controlling wide categories of many pathogens. In this respect, Hajihassani et al. [45] assessed the efficacy of application timing, that is, 5 days before planting (DBP) and at planting (AP) of X. bovienii and X. szentirmaii metabolites for the root-knot nematode (RKN) Meloidogyne incognita control on cabbage roots in two environmental conditions. At-plant applications of Paecilomyces lilacinus strain 251 (MeloCon WG) and secondary metabolites of Burkholderia rinojensis strain A396 (Majestene) and oxamyl (Vydate) were used for comparison. In the greenhouse, X. szentirmaii and Vydate at 5 DBP had a lower (p < 0.05) root gall rating than the untreated control. Vydate and all metabolite treatments showed significantly lower root galling relative to Majestene, MeloCon, and the control. In addition, the metabolites and Vydate decreased (p < 0.05) RKN egg counts per gram of root compared to the other treatments in the greenhouse. No differences were observed in the egg count between Vydate and the metabolites. At-plant and 5 DBP applications of X. bovienii and X. szentirmaii at decreased the total egg count relative to Majestene and the control in the greenhouse. Thus, the natural metabolites generated by the two Xenorhabdus species can control Mincognita regardless of application timing and are suggested as a potential alternative to nematicides in organic production systems [45]. In addition, direct effect of X. lircayensis, identified using the whole genome, was evaluated on a population of the plant-parasitic nematode Xiphinema index [46]. Supernatants of bacteria were discarded via centrifugation, then Xlircayensis were resuspended in phosphate-buffered saline (PBS) and set to 1 × 106 and 1 × 107 CFU mL−1 for laboratory and semi-field assays, respectively. Cell bacteria (1 × 107 CFU mL−1) were applied in the semi-field assay by 30 min dipping grapevine roots in the bacterial suspension. Afterward, these plants were established in 5 L pots filled with naturally Xindex-infested soil and immediately inoculated with 350 mL of the same Xlircayensis suspension. The nematicidal effects of Xlircayensis suspension appeared at 24 h post-inoculation but attained full (100%) X. index mortality after 72 h exposition (p < 0.001) in laboratory assays. In addition, under semi-field conditions, Xlircayensis suspension significantly (p ≤ 0.05) reduced X. index populations. While the study recommended Xlircayensis as a good candidate for further assesses in field conditions, additional analyses must be performed to set the metabolites, enzymes, and mode of action for its nematicidal aptitude [46]. Vicente-Díez et al. [47] tested the antibiotic impact of cell-free supernatants (CFSs) and unfiltered ferments (UFs) of X. nematophila and P. laumondii on another plant pathogenic category represented by the fungus Botrytis cinerea growth and compared the activity of bacteria isolated from a bio-fermenter with the commercial B. amyloliquefaciens (Serenade®ASO, Bayern CropScience). The UF and CFS of X. nematophila suppressed about 95% and 80% of Bcinerea growth, respectively, while both UF and CFS of P. laumondii inhibited only about 40%. These data showed the potential of CFS and UF of X. nematophila for B. cinerea control.
In another study [48]X. bovienii metabolite treatment was comparable to fenbuconazole (a commercial fungicide) in decreasing Fusicladium effusum sporulation on pecan (Carya illinoinensis) terminals. X. bovienii metabolite and broth treatments suppressed development of lesions brought about by Phytophthora cactorum (using pecan tree leaves maintained on agar). The bacterial metabolite treatment was also toxic to Armillaria tabescens, another important pathogen but of peach (Prunus persica) trees, especially in the southeastern United States [48]. These results offer a basis for further investigations on utilizing the bacterial metabolites or broth for suppression of economically significant diseases of pecan and peach. Likewise, Xnematophila generates many metabolites during growth and multiplication. Only one of these secondary metabolites (xenocoumacin 1) proved to have a robust antifungal activity for controlling Rhizoctonia solani [49]Botrytis cinerea [50]Alternaria alternate [51], several Phytophthora species, etc. [50][51][52][53]. These effects suggest, a priori, that other Xenorhabdus species, which are available or are likely to befit broadly soon, are able to control other pests and diseases. Recently, Xenorhabdus budapestensis strain C72 showed remarkable suppressing effect on spore germination and mycelial growth of the fungus Bipolaris maydis which causes the Southern corn leaf blight [54]. The relative control effect of the bacterial cell-free culture media reached 59.15% and 77.96% in greenhouse and field experiments, respectively, which was as efficacious as a commercial fungicide. The in vitro tests also indicated that C72 cell-free culture media with thermostability proved wide-spectrum antifungal efficacy towards other economically significant fungi and pathogens of plants [54]. Chacón-Orozco et al. [55] reported that among 16 strains of EPN-symbiotic bacteria, cell-free supernatants of Xszentrimaii had the highest fungicidal effect on mycelium growth of Sclerotinia sclerotiorum. They reported that Xszentrimaii produces volatile organic compounds that inhibit Ssclerotiorum growth and/or its consequent generation of sclerotia.
The discovery and cloning of additional useful compounds from Xenorhabdus are still in progress [22][36][37][56][57]. Factually, these bacteria can demonstrate metabolites with the major characteristics of safe pesticides. In other words, their effect is boosted with an enhanced dose, but a negative correlation is found between the number or density of pathogen/pest eggs, adult survival of the pest, percentage of hatching, and the Xenorhabdus bacterial dosage [8][20][21].

2.2. Xenorhabdus Bacterial Mechanism via Their Secreted Materials

The Xenorhabdus bacteria are typically famous for killing their hosts via toxemia/septicemia, within the form of the normal XenorhabdusSteinernema complex [8]. However, as different Steinernema species carrying specific Xenorhabdus strains can invade a single insect, Xenorhabdus spp. are also engaged in competition with both related strains and nonrelated gut microbes of the insect host [58]. This competition, in addition to Xenorhabdus having the capability to kill the insect host, can explain why Xenorhabdus spp. produce a treasure trove of diverse insecticidal and antimicrobial compounds. Moreover, Ciezki [58] found that Xbovienii and Xnematophila can generate R-type bacteriocins (xenorhabdicins) that are specifically active towards different Xenorhabdus and Photorhabdus species. The latter author stressed that xenorhabdicin activity could be predictive of competitive results between two Xenorhabdus strains, while other determinants, besides xenorhabdicins, were mainly included in the competitive success between the other Xenorhabdus strains. Thus, Ciezki [58] demonstrated that various Xenorhabdus antibiotics could define the output of interspecies competition in a natural host environment.
The mounting ambition to harness Xenorhabdus-derived compounds in industrial products stems from not only their abundance, but also their qualities that enhance their functions. Initially, standalone pathogenicity trials of Xenorhabdus bacteria and/or their released materials usually start with their direct injection into the haemocoel of insects via artificial means [8][43]Xenorhabdus protein toxins ordinarily have oral or/and injectable toxicity to insects. Xenorhabdus-derived compounds have a variety of modes of action that have been reviewed [20][46][58]. The suggested mode of action of Xenorhabdus–dithiopyrrolone derivatives (comprising the two metabolites xenorhabdins and xenorxides) is inhibition of RNA synthesis [59]. However, Xenorhabdus–indole-containing compounds could show additional mechanism via weak phospholipase A2 inhibitory effects. This latter, phospholipase A2, is necessary for producing eicosanoids. Eicosanoids have substantial role for activating the insect-immune response via mediating and modulating hemocyte behavior [60]. Thus, Dreyer et al. [20] assumed that indole-containing materials produced by Xnematophila can suppress the immune response of the insect host to be more vulnerable to microbial infection. Xenorhabdus budapestensis has two antimicrobial peptides, GP-19 and EP-20, with wide-spectrum antimicrobial activity against bacteria and fungi [61]. The first peptide, with a neutral charge, is suggested to cause a disruptive impact to the host membrane by moving to the cell surface and penetrating the membrane. The second peptide likely has a different mode of action. It is suggested to have an intracellular influence, by inhibiting protein synthesis, cell wall, and nucleic acid [61].
Complete genome sequencing of various Xenorhabdus species/strains has been uncovering the ability of these bacteria to produce numerous secondary metabolites. Thus, it can contribute to comprehensive examination of the molecular basis underlying the biological control activity of this Xenorhabdus strain [62]. Various types of biological molecules have been detected and characterized for Xenorhabdus bacteria. The main antimicrobial materials comprise ribosomal-encoded benzylideneace-tone [63] xenocin and bicornutin [64][65], and non-ribosomally generated xenematides [66], fabclavines [67], xenocoumacin [68], nematophin [69], rhabdopeptides [70], and peptide–antimicrobial–Xenorhabdus lipopeptides [71]. Knowing the attributes of these compounds, e.g., the range of pH and heating needed for their stability, should enable their successful use as alternatives to chemical pesticides in agriculture [19][20]. For example, depsipeptides are peptides that generally have alternating peptide and ester bonds, and five classes of depsipeptides have been characterized. The first class, produced by Xenorhabdus doucetiae and X. mauleonii and known as xenoamicin, are tridecadepsipeptides with hydrophobic amino acids [72]. The genome sequence of X. doucetiae DSM 17909 revealed that xenoamicins are encoded by five non-ribosomal peptide synthetases (NRPSs), XabABCD, and an aspartic acid decarboxylase (XabE). Due to its hydrophobic characteristics, xenoamicin can interact with the host–cytoplasmic membrane. Nevertheless, no antifungal or antibacterial activity has been listed for xenoamicin A, which displays a different mechanism. Xenoamicin A has weak cytotoxic and anti-protozoal activities [72]. The second class of depsipeptides, the lipodepsipeptides produced by X. indica, has supplemental fatty acid chain linked to one of the amino acids [73]. The peptides are named after their amino acid sequence and are known as taxlllaids (A–G). Natural taxlllaid A and synthetic taxlllaids B–G can manifest antiprotozoal activity. Taxlllaid A is optimistically cytotoxic to human carcinoma cells [73]. The third depsipeptides class are grouped as indole-containing xenematides. Xenematide A, secreted by Xnematophila [74], is antibacterial and insecticidal. The other two depsipeptide classes contain szentiamide and xenobactin isolated from Xszentirmaii and Xenorhabdus sp., strain PB30.3 [75][76]. Both compounds are active against Plasmodium falciparum (protozoan parasite of humans) and have some activity against Trypanosoma brucei rhodesiense and Trypanosoma cruzi (parasites of many vertebrates). Szentiamide possesses a weak cytotoxic activity against Galleria mellonella hemocytes. Xenobactin has no cytotoxic activity; yet, it is active against Micrococcus luteus. This antibacterial activity is mostly due to its hydrophobic status where it probably targets the bacterial cell membrane [19]. Eventually, each of the aforementioned groups of toxins has a conceivable role as a biocontrol material, via a particular mode of action against pathogens and arthropod pests such as vector insects. The differential virulence of the candidate toxins can be correlated not only with their interspecies/strain gene sequence diversity of the same EPN-symbiotic bacterial genus but also between the two EPN-symbiotic bacterial genera, Xenorhabdus and Photorhabdus [33][55]. Fabclavine is broadly generated in Xenorhabdus species but Photorhabdus species do not produce fabclavines, except for P. asymbiotica [77]. This can elucidate partially why the tested Photorhabdus species (PkayaiiPnamnaoensisPlaumondiiPakhurstiiPthracensis) did not show antiprotozoal activity [33]. On the contrary, fabclavines 1a and 1b demonstrate diverse bioactivities against various bacterial, fungal, and protozoal organisms [68]. Other antiprotozoal bioactive materials produced by 22 Xenorhabdus species are xenorhabdins, xenocoumacins, and PAX peptides. Thus, the tested Xenorhabdus species were more effective against the serious human protozoal parasites Entamoeba histolyticaAcanthamoeba castellaniiTrichomonas vaginalisTrypanosoma cruzi, and Leishmania tropica [33]. Furthermore, it is quite possible that more Xenorhabdus-derived toxins will uncover certain variations among bacterial strains regarding toxicity to these pests. Various features and details concerning the mode of action, structure, and putative function of the Xenorhabdus-bioactive compounds in the process of infection have been clarified [20][21][58].
Xenorhabdus bacteria can control economically significant endoparasitic species of nematodes inside plant roots via their antibiotic compounds and toxins [78]. Moreover, the bacterium-derived protease inhibitor protein could be genetically transformed into tobacco plants in order to offer protection from the aphids Myzus persicae [31]. Therefore, such genetically engineered techniques are suggested as a promising replacement to the Bt toxin [7], to preclude development of insect resistance [56]. The numerous instances of pathogen and arthropod pest killing induced by Xenorhabdus spp. [8][21][25][36][37][61][78] do not deny the variations in the immune response among their pathogen/pest hosts [33][34][36][79]. In addition, the difference in immune reaction among host species/strains may be based on biologic/genetic and evolutionary/ecological factors set for each pathogen–host system. The various system constituents, including specificity, induction, and memory of the immunity, can determine the cognate resistance mechanism of the intended insect population/species [80]. Generally, physical parameters, especially pH, temperature, and sodium chloride, could variably affect the mortality percentage induced by these metabolites to the Gmellonella larvae [81].

References

  1. Thomas, G.M.; Poinar, G.O. Xenorhabdus gen. nov., a genus of entomopathogenic nematophilic bacteria of the family Enterobacteriaceae. Int. J. Syst. Bacteriol. 1979, 29, 352–360.
  2. Boemare, N.E.; Akhurst, R.J.; Mourant, R.G. DNA relatedness between Xenorhabdus spp. (Enterobacteriaceae), symbiotic bacteria of entomopathogenic nematodes, and a proposal to transfer Xenorhabdus luminescens to a new genus, Photorhabdus gen. nov. Int. J. Syst. Bacteriol. 1993, 43, 249–255.
  3. Fischer-Le Saux, M.; Viallard, V.; Brunel, B.; Normand, P.; Boemare, N.E. Polyphasic classification of the genus Photorhabdus and proposal of new taxa: P. luminescens subsp. luminescens subsp. nov., P. luminescens subsp. akhurstii subsp. nov., P. luminescens subsp. laumondii subsp. nov., P. temperata sp. nov., P. temperate subsp. temperatas ubsp. nov. and P. asymbiotica sp. nov. Int. J. Syst. Bacteriol. 1999, 49, 1645–1656.
  4. Boemare, N. Biology, taxonomy and systematics of Photorhabdus and Xenorhabdus. In Entomopathogenic Nematology; Gaugler, R., Ed.; CAB International: Wallingford, UK, 2002; pp. 35–56.
  5. Sajnaga, E.; Kazimierczak, W. Evolution and taxonomy of nematode-associated entomopathogenic bacteria of the genera Xenorhabdus and Photorhabdus: An overview. Symbiosis 2020, 80, 1–13.
  6. Shapiro-Ilan, D.; Hazir, S.; Glazer, I. Advances in use of entomopathogenic nematodes. In Integrated Management of Insect Pests: Current and Future Developments; Kogan, M., Heinrichs, E.A., Eds.; Burleigh Dodds Science Publication: Cambridge, UK, 2020; pp. 1–30.
  7. Abd-Elgawad, M.M.M. Photorhabdus spp.: An overview of the beneficial aspects of mutualistic bacteria of insecticidal nematodes. Plants 2021, 10, 1660.
  8. Javed, N.; Kamran, M.; Abbas, H. Toxic secretions of Xenorhabdus and their efficacy against crop insect pests. In Biocontrol Agents: Entomopathogenic and Slug Parasitic Nematodes; Abd-Elgawad, M.M.M., Askary, T.H., Coupland, J., Eds.; CAB International: Wallingford, UK, 2017; pp. 223–230.
  9. Koppenhöfer, H.S.; Gaugler, R. Entomopathogenic nematode and bacteria mutualism. In Defensive Mutualism in Microbial Symbiosis; White, J., Torres, M., Eds.; CRC Press: Boca Raton, FL, USA, 2009; pp. 99–116.
  10. Hillman, K.; Goodrich-Blair, H. Are you my symbiont? Microbial polymorphic toxins and antimicrobial compounds as honest signals of beneficial symbiotic defensive traits. Curr. Opin. Microbiol. 2016, 31, 184–190.
  11. Henry, L.M.; Peccoud, J.; Simon, J.C.; Hadfield, J.D.; Maiden, M.J.; Ferrari, J.; Godfray, H.C. Horizontally transmitted symbionts and host colonization of ecological niches. Curr. Biol. 2013, 9, 1713–1717.
  12. Maher, A.M.D.; Asaiyah, M.A.M.; Brophy, C.; Griffin, C.T. An entomopathogenic nematode extends its niche by associating with different symbionts. Microb. Ecol. 2017, 73, 211–223.
  13. Chang, D.Z.; Serra, L.; Lu, D.; Mortazavi, A.; Dillman, A.R.A. Core set of venom proteins is released by entomopathogenic nematodes in the genus Steinernema. PLoS Pathog. 2019, 15, e1007626.
  14. Murfin, K.E.; Whooley, A.C.; Klassen, J.L.; Goodrich-Blair, H. Comparison of Xenorhabdus bovienii bacterial strain genomes reveals diversity in symbiotic functions. BMC Genom. 2015, 16, 889.
  15. McMullen, J.G.; Peterson, B.F.; Forst, S.; Goodrich-Blair, H.; Stock, S.P. Fitness costs of symbiont switching using entomopathogenic nematodes as a model. BMC Evol. Biol. 2017, 17, 100.
  16. Tailliez, P.; Laroui, C.; Ginibre, N.; Paule, A.; Pages, S.; Boemare, N. Phylogeny of Photorhabdus and Xenorhabdus based on universally conserved protein-coding sequences and implications for the taxonomy of these two genera. Proposal of new taxa. X. vietnamensis sp. nov., P. luminescens subsp. caribbeanensis subsp. nov., P. luminescens subsp. hainanensis subsp. nov., P. temperata subsp. khanii subsp. nov., P. temperata subsp. tasmaniensis subsp. nov., and the reclassification of P. luminescens subsp. thracensis as P. temperata subsp. thracensis comb. nov. Int. J. Syst. Evol. Microbiol. 2010, 60, 1921–1937.
  17. Askary, T.H.; Abd-Elgawad, M.M.M. Opportunities and challenges of entomopathogenic nematodes as biocontrol agents in their tripartite interactions. Egypt. J. Biol. Pest Cont. 2021, 31, 42.
  18. Baiocchi, T.; Abd-Elgawad, M.M.M.; Dillman, A.R. Genetic improvement of entomopathogenic nematodes for enhanced biological control. In Biocontrol Agents: Entomopathogenic and Slug Parasitic Nematodes; Abd-Elgawad, M.M.M., Askary, T.H., Coupland, J., Eds.; CAB International: Wallingford, UK, 2017; pp. 505–517.
  19. Hazir, S.; Shapiro-Ilan, D.I.; Bock, C.H.; Hazir, C.; Leite, L.G.; Hotchkiss, M.W. Relative potency of culture supernatants of Xenorhabdus and Photorhabdus spp. on growth of some fungal phytopathogens. Eur. J. Plant Pathol. 2016, 146, 369–381.
  20. Dreyer, J.; Malan, A.P.; Dicks, L.M.T. Bacteria of the genus Xenorhabdus, a novel source of bioactive compounds. Front. Microbiol. 2018, 9, 3177.
  21. Cimen, H.; Touray, M.; Gulsen, S.H.; Hazir, S. Natural products from Photorhabdus and Xenorhabdus: Mechanisms and impacts. Appl. Microbiol. Biotechnol. 2022, 106, 4387–4399.
  22. Booysen, E.; Rautenbach, M.; Stander, M.A.; Dicks, L.M. Profiling the production of antimicrobial secondary metabolites by Xenorhabdus khoisanae J194 under different culturing conditions. Front. Chem. 2021, 9, 626653.
  23. Nunez-Valdez, M.E.; Lanois, A.; Pages, S.; Duvic, B.; Gaudriault, S. Inhibition of Spodoptera frugiperda phenoloxidase activity by the products of the Xenorhabdus rhabduscin gene cluster. PLoS ONE 2019, 22, e0212809.
  24. Plichta, K.L.; Joyce, S.A.; Clarke, D.; Waterfield, N.; Stock, S.P. Heterorhabditis gerrardi n. sp. (Nematoda: Heterorhabditidae): The hidden host of Photorhabdus asymbiotica (Enterobacteriaceae:g-Proteobacteria). J. Helminthol. 2009, 83, 309–320.
  25. Abd-Elgawad, M.M.M. Towards optimization of entomopathogenic nematodes for more service in the biological control of insect pests. Egypt. J. Biol. Pest Cont. 2019, 29, 77.
  26. Koppenhöfer, A.M.; Shapiro-Ilan, D.I.; Hiltpold, I. Entomopathogenic nematodes in sustainable food production. Front. Sustain. Food Syst. 2020, 4, 125.
  27. Morgan, J.A.W.; Kuntzelmann, V.; Tavernor, S.; Ousley, M.A.; Winstanley, C. Survival of Xenorhabdus nematophilus and Photorhabdus luminescens in water and soil. J. Appl. Microbiol. 1997, 83, 665–670.
  28. Antonello, A.M.; Sartori, T.; Silva, M.B.; Prophiro, J.S.; Pinge-Filho, P.; Heermann, R. Anti-Trypanosoma activity of bioactive metabolites from Photorhabdus luminescens and Xenorhabdus nematophila. Exp. Parasitol. 2019, 204, 107724.
  29. da Silva, W.J.; Pilz-Júnior, H.L.; Heermann, R.; da Silva, O.S. The great potential of entomopathogenic bacteria Xenorhabdus and Photorhabdus for mosquito control: A review. Parasites Vectors 2020, 13, 376.
  30. Migunova, V.D.; Sasanelli, N. Bacteria as biocontrol tool against phytoparasitic nematodes. Plants 2021, 10, 389.
  31. Zhang, H.; Mao, J.; Liu, F.; Zeng, F. Expression of a nematode symbiotic bacterium-derived protease inhibitor protein in tobacco enhanced tolerance against Myzus persicae. Plant Cell Rep. 2012, 31, 1981–1989.
  32. Zhang, Y.; Nangong, Z.; Kong, F.; Song, P.; Wang, Q. Biological activity of Xenorhabdus nematophila HB310 against Locusta migratoria manilensis. Chin. J. Pest Sci. 2013, 15, 516–522.
  33. Gulsen, S.H.; Tileklioglu, E.; Bode, E.; Cimen, H.; Ertabaklar, H.; Ulug, D.; Ertug, S.; Wenski, S.L.; Touray, M.; Hazir, C.; et al. Antiprotozoal activity of different Xenorhabdus and Photorhabdus bacterial secondary metabolites and identification of bioactive compounds using the easyPACId approach. Sci. Rep. 2022, 12, 10779.
  34. Sergeant, M.; Baxter, L.; Jarrett, P.; Shaw, E.; Ousley, M.; Winstanley, C.; Alun, J.; Morgan, W. Identification, typing and insecticidal activity of Xenorhabdus isolates from entomopathogenic nematodes in United Kingdom soil and characterization of the xpt toxin loci. Appl. Environ. Microbiol. 2006, 72, 5895–5907.
  35. Pidot, S.J.; Coyne, S.; Kloss, F.; Hertweck, C. Antibiotics from neglected bacterial sources. Int. J. Med. Microbiol. 2014, 304, 14–22.
  36. Tomar, P.; Thakur, N.; Yadav, A.N. Endosymbiotic microbes from entomopathogenic nematode (EPNs) and their applications as biocontrol agents for agro-environmental sustainability. Egypt. J. Biol. Pest. Control 2022, 32, 80.
  37. Abebew, D.; Sayedain, F.S.; Bode, E.; Bode, H.B. Uncovering nematicidal natural products from Xenorhabdus bacteria. J. Agric. Food Chem. 2022, 70, 498–506.
  38. Dreyer, J.; Rautenbach, M.; Booysen, E.; Van Staden, A.; Deane, S.; Dicks, L. Xenorhabdus khoisanae SB10 produces Lys-rich PAX lipopeptides and a Xenocoumacin in its antimicrobial complex. BMC Microbiol. 2019, 19, 1–11.
  39. Abdel-Razek, A.S. Pathogenic effects of Xenorhabdus nematophilus and Photorhabdus luminescens (Enterobacteriaceae) against pupae of the Diamondback Moth, Plutella xylostella (L.). J. Pest Sci. 2003, 76, 108–111.
  40. Askary, T.H.; Abd-Elgawad, M.M.M. Beneficial nematodes in agroecosystems: A global perspective. In Biocontrol Agents: Entomopathogenic and Slug Parasitic Nematodes; Abd-Elgawad, M.M.M., Askary, T.H., Coupland, J., Eds.; CAB International: Wallingford, UK, 2017; pp. 3–25.
  41. Eom, S.; Park, Y.; Kim, H.; Kim, Y. Development of a high efficient“dual Bt-plus” insecticide using a primary form of an entomopathogenic bacterium, Xenorhabdus nematophila. J. Microbiol. Biotechnol. 2014, 24, 507–521.
  42. Seongchae, J.; Yonggyun, K. Synergistic effect of entomopathogenic bacteria (Xenorhabdus sp. and Photorhabdus temperata ssp. temperata) on the pathogenicity of Bacillus thuringiensis ssp. Aizawai against Spodoptera exigua (Lepidoptera: Noctuidae). Environ. Entomol. 2006, 35, 1584–1589.
  43. Mahar, A.N.; Munir, M.; Gowen, S.R.; Hague, N.G.M. Role of entomopathogenic bacteria, Photorhabdus luminescens and its toxic secretions against Galleria mellonella larvae. J. Entomol. 2005, 2, 69–76.
  44. Wang, H.; Dong, H.; Qian, H.; Xia, R.; Cong, B. Isolation, bioassay and characterisation of Xenorhabdus sp. SY5, a highly virulent symbiotic bacterium of an entomopathogenic nematode isolated from China. Nematology 2013, 15, 153–163.
  45. Hajihassani, A.; Gitonga, D.; Timper, P.; Shapiro-Ilan, D. Effects of application timing on the efficacy of Xenorhabdus and Photorhabdus metabolites for control of Meloidogyne incognita. In Proceedings of the 7th International Congress of Nematology, Antibes Juan-les-Pins, France, 1–5 May 2022; p. 598.
  46. Castaneda-Alvarez, C.; Prodan, S.; San-Blas, E.; Aballay, E. Symbiotic bacteria of entomopathogenic nematodes for the biocontrol of dagger nematode Xiphinema index. In Proceedings of the 7th International Congress of Nematology, Antibes Juan-les-Pins, France, 1–5 May 2022; p. 587.
  47. Vicente-Díez, I.; Blanco-Pérez, R.; Chelkha, M.; Pou1, A.; Campos-Herrera, R. Plasticity in the use of Xenorhabdus nematophila and Photorhabdus laumondii against Botrytis cinerea. In Proceedings of the 7th International Congress of Nematology, Antibes Juan-les-Pins, France, 1–5 May 2022; p. 660.
  48. Shapiro-Ilan, D.I.; Bock, C.H.; Hotchkiss, M.W. Suppression of pecan and peach pathogens on different substrates using Xenorhabdus bovienii and Photorhabdus luminescens. Biol. Cont. 2014, 77, 1–6.
  49. Zhang, S.; Liu, Q.; Han, Y.; Han, J.; Yan, Z.; Wang, Y.; Zhang, X. Nematophin, an antimicrobial dipeptide compound from Xenorhabdus nematophila YL001 as a potent biopesticide for Rhizoctonia solani control. Front. Microbiol. 2019, 10, 1765.
  50. Fang, X.; Zhang, M.; Tang, Q.; Wang, Y.; Zhang, X. Inhibitory effect of Xenorhabdus nematophila TB on plant pathogens Phytophthora capsici and Botrytis cinerea in vitro and in planta. Sci. Rep. 2014, 4, 4300.
  51. Qin, Y.; Jia, F.; Li, X.; Li, B.; Ren, J.; Yang, X.; Li, G. Improving the yield of xenocoumacin 1 by PBAD promoter replacement in Xenorhabdus nematophila CB6. Agriculture 2021, 11, 1251.
  52. Yang, X.; Qiu, D.; Yang, H.; Liu, Z.; Zeng, H.; Yuan, J. Antifungal activity of xenocoumacin 1 from Xenorhabdus nematophilus var. pekingensis against Phytophthora infestans. World J. Microb. Biotechnol. 2011, 27, 523–528.
  53. Zhou, T.; Yang, X.; Qiu, D.; Zeng, H. Inhibitory effects of xenocoumacin 1 on the different stages of Phytophthora capsici and its control effect on Phytophthora blight of pepper. BioControl 2017, 62, 151–160.
  54. Li, B.; Kong, L.; Qiu, D.; Francis, F.; Wang, S. Biocontrol potential and mode of action of entomopathogenic bacteria Xenorhabdus budapestensis C72 against Bipolaris maydis. Biol. Cont. 2021, 158, 104605.
  55. Chacón-Orozco, J.G.; Bueno, C., Jr.; Shapiro Ilan, D.I.; Hazir, S.; Leite, L.G.; Harakava, R. Antifungal activity of Xenorhabdus spp. and Photorhabdus spp. against the soybean pathogenic Sclerotinia sclerotiorum. Sci. Rep. 2020, 10, 20649.
  56. Xiao, Y.; Wu, K. Recent progress on the interaction between insects and Bacillus thuringiensis crops. Philos. Trans. R. Soc. 2019, 374, 20180316.
  57. Alotaibi, S.S.; Darwish, H.; Alharthi, S.; Alghamdi, A.; Noureldeen, A.; Fallatah, A.M.; Fodor, A.; Al-Barty, A.; Albogami, B.; Baazeem, A. Control potentials of three entomopathogenic bacterial isolates for the carob moth, Ectomyelois ceratoniae (Lepidoptera: Pyralidae) in pomegranates. Agriculture 2021, 11, 1256.
  58. Ciezki, K.J. New Insights into the Role of Antimicrobials of Xenorhabdus in Interspecies Competition. Ph.D. Dissertation, University of Wisconsin-Milwaukee, Milwaukee, WI, USA, 2017; 103p.
  59. Oliva, B.; O’Neill, A.; Wilson, J.M.; O’Hanlon, P.J.; Chopra, I. Antimicrobial properties and mode of action of the pyrrothine holomycin. Antimicrob. Agents Chemother. 2001, 45, 532–539.
  60. Shrestha, S.; Kim, Y. An entomopathogenic bacterium, Xenorhabdus nematophila, inhibits hemocyte phagocytosis of Spodoptera exigua by inhibiting phospholipase A2. J. Invertebr. Pathol. 2007, 96, 64–70.
  61. Xiao, Y.; Meng, F.; Qiu, D.; Yang, X. Two novel antimicrobial peptides purified from the symbiotic bacteria Xenorhabdus budapestensis NMC-10. Peptides 2012, 35, 253–260.
  62. Li, B.; Qiu, D.; Wang, S. Complete genome sequence data of Xenorhabdus budapestensis strain C72, a candidate biological control agent from China. Plant Dis. 2021, 105, 3276–3278.
  63. Ji, D.; Yi, Y.; Kang, G.-H.; Choi, Y.-H.; Kim, P.; Baek, N.-I.; Kim, Y. Identification of an antibacterial compound, benzylideneacetone, from Xenorhabdus nematophila against major plant-pathogenic bacteria. FEMS Microbiol. Lett. 2004, 239, 241–248.
  64. Böszörményi, E.; Ersek, T.; Fodor, A.; Fodor, A.M.; Földes, L.S.; Hevesi, M.; Hogan, J.S.; Katona, Z.; Klein, M.G.; Kormany, A.; et al. Isolation and activity of Xenorhabdus antimicrobial compounds against the plant pathogens Erwinia amylovora and Phytophthora nicotianae. J. Appl. Microbiol. 2009, 107, 746–759.
  65. Rathore, J.S. Expression, purification, and functional characterization of atypical xenocin, its immunity protein, and their domains from Xenorhabdus nematophila. Int. J. Bacteriol. 2013, 2013, 746862.
  66. Xi, X.; Lu, X.; Zhang, X.; Bi, Y.; Li, X.; Yu, Z. Two novel cyclic depsipeptides xenematides F and G from the entomopathogenic bacterium Xenorhabdus budapestensis. J. Antibiot. 2019, 72, 736–743.
  67. Fuchs, S.W.; Grundmann, F.; Kurz, M.; Kaiser, M.; Bode, H.B. Fabclavines: Bioactive peptide-polyketide-polyamino hybrids from Xenorhabdus. ChemBioChem 2014, 15, 512–516.
  68. Park, D.; Ciezki, K.; van der Hoeven, R.; Singh, S.; Reimer, D.; Bode, H.B.; Forst, S. Genetic analysis of xenocoumacin antibiotic production in the mutualistic bacterium Xenorhabdus nematophila. Mol. Microbiol. 2009, 73, 938–949.
  69. Li, J.; Chen, G.; Webster, J.M. Nematophin, a novel antimicrobial substance produced by Xenorhabdus nematophilus (Enterobactereaceae). Can. J. Microbiol. 1997, 43, 770–773.
  70. Zhao, L.; Kaiser, M.; Bode, H.B. Rhabdopeptide/xenortide-like peptides from Xenorhabdus innexi with terminal amines showing potent antiprotozoal activity. Org. Lett. 2018, 20, 5116–5120.
  71. Fuchs, S.W.; Proschak, A.; Jaskolla, T.W.; Karas, M.; Bode, H.B. Structure elucidation and biosynthesis of lysine-rich cyclic peptides in Xenorhabdus nematophila. Org. Biomol. Chem. 2011, 9, 3130–3132.
  72. Zhou, Q.; Grundmann, F.; Kaiser, M.; Schiell, M.; Gaudriault, S.; Batzer, A. Structure and biosynthesis of xenoamicins from entomopathogenic Xenorhabdus. Chem. Eur. J. 2013, 19, 16772–16779.
  73. Kronenwerth, M.; Brachmann, A.O.; Kaiser, M.; Bode, H.B. Bioactive derivatives of isopropylstilbene from mutasynthesis and chemical synthesis. ChemBioChem 2014, 15, 2689–2691.
  74. Lang, G.; Kalvelage, T.; Peters, A.; Wiese, J.; Imhoff, J.F. Linear and cyclic peptides from the entomopathogenic bacterium Xenorhabdus nematophilus. J. Nat. Prod. 2008, 71, 1074–1077.
  75. Nollmann, F.I.; Dowling, A.; Kaiser, M.; Deckmann, K.; Grösch, S.; French-Constant, R.; Bode, H.B. Synthesis of szentiamide, a depsipeptide from entomopathogenic Xenorhabdus szentirmaii with activity against Plasmodium falciparum. Beilstein J. Org. Chem. 2012, 8, 528–533.
  76. Grundmann, F.; Kaiser, M.; Kurz, M.; Schiell, M.; Batzer, A.; Bode, H.B. Structure determination of the bioactive depsipeptide xenobactin from Xenorhabdus sp. PB30. 3. RSC Adv. 2013, 3, 22072–22077.
  77. Tobias, N.J.; Wolff, H.; Djahanschiri, B.; Grundmann, F.; Kronenwerth, M.; Shi, Y.M. Natural product diversity associated with the nematode symbionts Photorhabdus and Xenorhabdus. Nat. Microbiol. 2017, 2, 1676–1685.
  78. Kepenekci, I.; Hazir, S.; Oksal, E.; Lewis, E.E. Application methods of Steinernema feltiae, Xenorhabdus bovienii and Purpureocillium lilacinum to control root-knot nematodes in greenhouse tomato systems. Crop Prot. 2018, 108, 31–38.
  79. Eroglu, C.; Cimenb, H.; Ulug, D.; Karagoz, M.; Hazir, S.; Cakmaka, I. Acaricidal effect of cell-free supernatants from Xenorhabdus and Photorhabdus bacteria against Tetranychus urticae (Acari: Tetranychidae). J. Invertebr. Pathol. 2019, 106, 61–66.
  80. Erler, S.; Popp, M.; Lattorff, M. Dynamics of immune system gene expression upon bacterial challenge and wounding in a social insect (Bombus terrestris). PLoS ONE 2011, 6, e18126.
  81. Abd El-Zaher, F.H.; Abd-Elgawad, M.M.M.; Abd El-Maksoud, H.K. Use of the entomopathogenic nematode symbiont Photorhabdus luminescens as a biocontrol agent. B- Factors affecting the cell-free filtrates from the bacterium. J. Appl. Sci. Res. 2012, 8, 4600–4614.
More
Information
Subjects: Microbiology
Contributor MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to https://encyclopedia.pub/register :
View Times: 510
Revisions: 2 times (View History)
Update Date: 24 Nov 2022
1000/1000
Video Production Service