Next Article in Journal
Lactic Acid-Based Natural Deep Eutectic Solvents to Extract Bioactives from Marine By-Products
Next Article in Special Issue
Fumigant Activity of Bacterial Volatile Organic Compounds against the Nematodes Caenorhabditis elegans and Meloidogyne incognita
Previous Article in Journal
The Expression Profiles of the Salvia miltiorrhiza 3-Hydroxy-3-methylglutaryl-coenzyme A Reductase 4 Gene and Its Influence on the Biosynthesis of Tanshinones
Previous Article in Special Issue
Seasonal Variability of a Caryophyllane Chemotype Essential Oil of Eugenia patrisii Vahl Occurring in the Brazilian Amazon
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:

Application Potential of Bacterial Volatile Organic Compounds in the Control of Root-Knot Nematodes

Graduate Institute of Biotechnology, National Chung Hsing University, Taichung 402, Taiwan
Department of Life Science, National Chung Hsing University, Taichung 402, Taiwan
Author to whom correspondence should be addressed.
Molecules 2022, 27(14), 4355;
Received: 15 June 2022 / Revised: 4 July 2022 / Accepted: 5 July 2022 / Published: 7 July 2022
(This article belongs to the Special Issue Progress in Volatile Organic Compounds Research II)


Plant-parasitic nematodes (PPNs) constitute the most damaging group of plant pathogens. Plant infections by root-knot nematodes (RKNs) alone could cause approximately 5% of global crop loss. Conventionally, chemical-based methods are used to control PPNs at the expense of the environment and human health. Accordingly, the development of eco-friendly and safer methods has been urged to supplement or replace chemical-based methods for the control of RKNs. Using microorganisms or their metabolites as biological control agents (BCAs) is a promising approach to controlling RKNs. Among the metabolites, volatile organic compounds (VOCs) have gained increasing attention because of their potential in the control of not only RKNs but also other plant pathogens, such as insects, fungi, and bacteria. This review discusses the biology of RKNs as well as the status of various control strategies. The discovery of VOCs emitted by bacteria from various environmental sources and their application potential as BCAs in controlling RKNs are specifically addressed.

1. Introduction

Plant-parasitic nematodes (PPNs) are known to be one of the greatest threats to agricultural production, causing an annual crop loss of more than USD 150 billion worldwide [1,2,3,4]. To date, over 4100 species of PPNs have been reported [5,6], and they can be classified into three groups according to their lifestyles: sedentary endoparasites, e.g., root-knot nematodes (Meloidogyne spp.) and cyst nematodes (Heterodera and Globodera spp.); migratory endoparasites, e.g., lesion nematodes (Pratylenchus spp.) and burrowing nematodes (Radopholus spp.); and migratory ectoparasites, e.g., Belonolaimus spp., Xiphenema spp., and Trichodorus spp. [7]. Among them, root-knot nematodes (RKNs) are the most important agricultural pests, infecting the roots of over 3000 plant species [8,9,10].
Although chemical nematicides are still the most effective means for the management of RKNs, withdrawal of such chemical agents from the market has been continuously urged due to safety and environmental concerns [11,12,13,14]. To respond to the increasing demand for eco-friendly and sustainable management to control RKNs, methods using live microorganisms or their metabolites have been intensively explored recently.
Nematodes in soil are exposed to a diversity of microorganisms [15], of which nematophagous bacteria and fungi represent the most promising candidates to control RKNs. Bacterial species of a range of genera, such as Bacillus, Pseudomonas, and Pasteuria, were observed to exhibit antagonistic activity against RKNs, while the fungi that were detrimental to RKNs were commonly isolated from the phylum Ascomycota, Basidiomycota, Zygomycota, and Chytridiomycota [7,15,16,17,18]. With regard to microbial metabolites, volatile organic compounds (VOCs) have attracted research attention in recent years due to their efficacy in killing RKNs [7,19,20]. Additionally, the application of VOCs in agricultural practice could be both economically affordable and less toxic to humans than conventional nematicides [21].
This review paper summarizes (i) the general knowledge of the life cycle and genome of RKNs, (ii) the current status of the management strategies used in the control of RKNs, and (iii) recent progress in the identification of bacterial VOCs and their application potential in the control of RKNs.

2. Root-Knot Nematodes (RKNs)

Many economically important crops are hosts of RKNs, including tomato, potato, corn, soybean, maize, oats, wheat, and cotton [22,23,24]. The economic loss caused by RKNs has been estimated at USD 78 billion annually worldwide, accounting for half of the total loss due to PPNs [25]. Although the genus Meloidogyne consists of about 100 species [26], M. incognita, M. arenaria, M. javanica, and M. hapla are the four major species that infect more than 2000 plant species, particularly underground plant organs [22,27,28,29].

2.1. Life Cycle

The life span of RKNs is about three to six weeks with a cycle comprising embryo, juvenile (J1, J2, J3, and J4), and adult stages [22]. RKNs reproduce via diverse mechanisms but mostly by parthenogenesis. The eggs of RKNs are laid in gelatinous masses in the soil or plant residues. The worms hatch as second-stage juveniles (J2), and they immediately move toward the roots of plant hosts, attack the elongation zone, and migrate to the root tip [5,30]. When they reach the apical meristem region, they transmigrate to the developing vascular cylinder, triggering the formation of giant cells, which serve as nutrient sinks to support the growth of the nematode. The juveniles then become sedentary and undergo three more molts before they turn into adults [31,32]. In the adult stage, the worm-shaped males move out of the plant root, but the sedentary females continuously develop into pear-shaped females. Afterward, the female adults begin laying eggs (more than 1000 eggs per female) on the external surface of the root [22,33,34].

2.2. Genome

The whole genome of mitotic obligate parthenogenetic M. incognita was determined to be approximately 86 Mb, which contains 19,212 protein-coding genes, while that of meiotic facultative parthenogenetic M. hapla was about 54 Mb, containing 14,700 protein-coding genes [1,35]. Lately, the gene numbers of M. arenaria, M. javanica, and M. incognita were predicted to be 30,308, 26,917, and 24,714, respectively [36]. These genomes share some common features but with their own characteristics. One of the features shared by M. incognita and M. hapla is the possession of genes encoding distinct plant-cell-wall-degrading enzymes. A phylogenetic analysis suggested that these genes, which are absent in animals, were probably obtained via horizontal gene transfer from fungi or bacteria [37]. Since these enzymes are also present in some other PPNs of the order Tylenchida, the acquisition of these genes might occur earlier in an ancestor of Tylenchida during evolution, which supported the progress of their capability to parasitize plants [32,38,39].
The most notable differences between M. incognita and M. hapla are their genome structure and reproduction mode. M. hapla has an ordinary genome structure of diploid sexual species, while M. incognita is a hypotriploid with a proportion of one genome present in a second copy. Furthermore, M. hapla reproduces with meiosis, whereas M. incognita reproduces without meiosis and fusion of gametes.

3. Control Strategies for RKNs

Given the great damage to crop production due to the infestation of PPNs, a variety of methods have been used to control nematodes. These methods can be categorized into physical, chemical, and biological control strategies.

3.1. Physical Control Strategies

Nematodes are highly vulnerable before they penetrate the host plant’s roots. Therefore, targeting PPNs at their vulnerable stages could be effective. For instance, increasing soil temperatures above 40 °C by solarization is an effective way to reduce the number of nematodes in soil [24]. Moisture is another critical factor for the survival of nematodes. It has been highlighted that an insufficient amount of water in the soil would affect nematodes’ ability to move toward their host roots [40,41]. Flooding represents an opposite strategy to control nematodes in soils. Many PPNs are intolerant to oxygen starvation; therefore, flooding can kill nematodes by limiting their supply of oxygen. Similar effects were observed when nematodes were stored in deep water in a laboratory. To be effective in the field, the duration of anaerobiosis must be long enough to kill the nematodes. However, flooding may not be practicable for every agricultural practice. Taking into consideration the threat of global climate change, flooding would not be a good option to control PPNs. In brief, physical control strategies are less effective than conventional chemical control strategies, although the cost of physical control strategies is relatively lower [42].

3.2. Chemical Control Strategies

Using synthetic chemicals with the features of fumigants or nematicides to control PPNs was a common method applied in agriculture in the previous half-century [43,44]. For example, methyl bromide and dibromochloropropane were intensively used as soil fumigants due to their effectiveness. However, they are highly toxic chemicals causing acute respiratory toxicity and neurotoxicity via inhalation [45,46,47,48]. Exposure to the dibromochloropropane that had accumulated in the soil was found to influence men’s fertility and was linked to certain human cancers [49,50,51]. Therefore, its use in agriculture was banned in 1979. In addition, methyl bromide is a strong ozone-depleting substance. The use of methyl bromide in fumigation was banned globally after 2015 under the directive of the Montreal Protocol, except for quarantine and pre-shipment treatments [52,53]. Recently, a couple of less environmentally toxic chemicals have been suggested as alternatives to methyl bromide [42,54,55,56]. However, they have not yet been registered for use in agriculture [14,16]. Nevertheless, farmers need more reliable, eco-friendly, and low-cost approaches for sustainable agriculture.

3.3. Biological Control Strategies

Biological control refers to the suppression of a pest population, or the pest’s harmful impact, by using living organisms (natural enemies) or their metabolites [57,58]. Because biological control imitates the competition among species in nature, it is generally thought to be more environmentally friendly than chemical control. The strategies of biological control can be classified into conservation, importation, and augmentation according to the source of the deployed organisms [59]. The conservation strategy is carried out to maintain the existing natural enemies in an environment; the importation strategy is carried out to introduce exotic enemies of the pests where they do not occur naturally; and the augmentation strategy is carried out to release reared natural enemies periodically into the habitat where the pests occur [60,61].
An organism (or its metabolites) that reduces the density of the pest population is defined as a biological control agent (BCA). An ideal BCA should exert its effects by multiple mechanisms without producing harmful substances to humans and the environment [62]. Bacteria from a wide range of genera have demonstrated the capability to control RKNs [63,64]. The common genera include Achromobacter, Arthrobacter, Bacillus, Burkholderia, Pasteuria, Pseudomonas, Rhizobium, and Serratia. The beneficial effects come from mechanisms such as parasitism, niche competition, the induction of plant systemic resistance, and the production of antagonistic substances (antibiotics, toxins, enzymes, VOCs, etc.) [15,63,65].
There is a growing interest in using Bacillus spp. to control PPNs. For example, Bacillus subtilis conferred induced systemic resistance to M. incognita on tomato plants under greenhouse conditions [66]. The treatment of tomato seeds with several strains of B. subtilis as well as the cell-free supernatant reduced the number of galls and egg masses of M. incognita. 9H-purine, uracil, and dihydrouracil, produced by Bacillus cereus and B. subtilis, showed nematicidal activity against Meloidogyne exigua [67]. The activity of dihydrouracil was even stronger than that of the commercial nematicide carbofuran. Bacillus firmus DS-1 had nematicidal activity against M. incognita. The serine protease produced by this strain, known as Sep1, was toxic to both M. incognita and C. elegans. In vitro experiments on C. elegans demonstrated that Sep1 has a destructive effect on multiple intestinal and cuticle-associated proteins, resulting in impaired physical barriers of the worm [17]. Bacillus amyloliquefaciens D1 efficiently influenced the mortality of M. incognita J2s and suppressed its egg hatching rate; it also had a plant growth-promoting effect. B. amyloliquefaciens Y1 produced cyclo(d-Pro-l-Leu) that functions as a nematicide against M. incognita [65]. Treatments of potato plants with a recombinant B. subtilis strain, which secreted plant-defense elicitor peptide StPep1, effectively reduced root galling caused by Meloidogyne chitwoodi [68]. The treatment of cucumber and tomato plants with Bacillus velezensis BZR 86 significantly reduced the development of root-knot disease caused by M. incognita, and, as a result, the growth of the plants was enhanced [69].
Bacillus thuringiensis (Bt) is a spore-forming bacterium that produces parasporal crystals (Cry) during the sporulation phase. Indeed, Cry proteins have been used as biological insecticides around the world for decades. Ingestion of Cry proteins is a prerequisite for the proteins to damage the guts of insects [70]. Although most Cry proteins are toxic to insects, experiments on different nematode species have confirmed that several families of Cry proteins, including Cry5, Cry6, Cry12, Cry13, Cry14, Cry21, and Cry55, target nematodes and exhibit nematicidal activity [71,72,73]. Feeding M. incognita with transgenic tomato roots that expressed Cry6A decreased the reproduction rate of the worm by a factor of 4 [74]. Cry6Aa2 not only showed toxicity to J2s but also suppressed the egg-hatching rate of M. hapla. In addition, a pot experiment indicated that soil drenching with a mixture of spores and Cry6Aa2 could reduce the number of galls and egg masses on plant roots as well as enhancing the growth of the plant [75]. Cry5 produced by Bt strain Sbt003 adversely affected the life span and reproduction of C. elegans; in addition, it had a detrimental impact on the worm’s intestine [76].
Pasteuria penetrans is a Gram-positive nematode-parasitic bacterium. The capability of P. penetrans to control RKNs has been investigated in several studies. The bacterial parasitism starts when endospores of P. penetrans attach to the cuticle of J2 nematodes; consequently, the infected J2s show a reduction in mobility and the ability to enter the roots of plant hosts [77]. The treatment of cucumber with P. penetrans in greenhouse trials reduced M. incognita populations in the roots of the plant [78]. The RNAi-mediated silencing of the selenium-binding protein Mi-SeBP-1 of M. incognita increased the attachment of P. penetrans endospore onto the J2s’ cuticles, revealing the involvement of Mi-SeBP-1 in the adhesion of the bacterial endospore on the nematode cuticle [79].
Pseudomonas simiae sMB751 and its secreted cyclic dipeptide, cyclo(l-Pro-l-Leu), displayed significant nematicidal activity against M. incognita J2s. In fact, it was observed in a pot experiment that the fermentation broth of P. simiae MB751 could suppress M. incognita infection and confer induced systemic resistance against nematodes on tomato plants [80]. The E. coli-expressed and purified Nif3-family protein YqfO03, originally from Pseudomonas syringae MB03, had nematicidal activity against both C. elegans and M. incognita [64]. The treatment of M. incognita-infected bell pepper plants with Burkholderia cepacia Bc-2 and Bc-F strains showed a reduction in the numbers of eggs and J2s of the worm [81]. Prodigiosin, the red pigment produced by Serratia marcescens, had toxicity against juveniles of M. javanica and Radopholus similis [82].

4. Volatile Organic Compounds (VOCs)

VOCs are carbon-based, low-molecular-weight compounds that have high vapor pressure and easily evaporate at room temperature [83,84,85]. VOCs emitted by microorganisms are capable of controlling plant-parasitic fungi, insects, bacteria, and nematodes [86]. Therefore, microbial VOCs are suitable to apply to different agricultural systems with relatively low concentrations compared to agrochemicals, and supplemental spray or drench is not essential for the application of VOCs [62,87,88,89]. Microbial VOCs are diverse in terms of their chemical structures. They can be alcohols, ketones, hydrocarbons, terpenes, fatty acids, or heteroatom-containing compounds [90]. A vast number of microbial VOCs are archived in the mVOC 2.0 database, in which more than 2000 VOCs from approximately 1000 different microorganisms are categorized based on chemical structures, mass spectra, and microbial emitters [91,92].
Solid-phase micro-extraction (SPME) is widely used for the collection of VOCs. In this method, VOCs are adsorbed by the SPME fiber from the headspace of a culture medium. The adsorbed compounds are then separated with gas chromatography and further identified with mass spectrometry. The culture conditions (medium composition, oxygen level, temperature, etc.) and physiological stages of microorganisms may influence the production of VOCs in terms of chemical types and amounts [93]. For instance, Lysobacter strains grown on potato dextrose agar (PDA) and nutrient agar (NA) produced different VOCs. Pyrazines, decanal, pyrrole, δ-hexalactone, and ethanol were emitted as VOCs when Lysobacter strains were cultivated on NA; however, indole and acetoin were the major VOCs when the bacteria were cultivated on PDA [94]. A recent study showed that B. gladioli BBB-01 emitted dimethyl disulfide as the primary VOC when the bacterium was cultivated on LB agar, whereas 2,5-dimethylfuran was emitted when the bacterium was cultivated on PDA [92]. Although there is insufficient information on the mechanism of VOC emission, it has been reported that the production of certain bacterial VOCs is regulated by the GacS/GacA two-component regulatory system [95].

4.1. Biocontrol of RKNs with Bacterial VOCs

The toxicity of microbial VOCs to RKNs has been shown in numerous reports. A VOC could affect nematodes by acting as a contact nematicide, fumigant, repellent, or attractant. It could also suppress the hatching of eggs. Some of these reports are briefly described in the following text. The frequently discovered VOCs and their reported functions are summarized in Table 1.
The nematicidal activity of Bacillus spp. has been shown in many reports. VOCs emitted by Bacillus megaterium YFM3.25 inhibited the hatching of eggs and reduced the infection of M. incognita in a pot experiment. Among the 17 VOCs, 2-nonanone, 2-undecanone, decanal, dimethyl disulfide, and benzeneacetaldehyde accounted for the fumigant toxicity against juveniles and eggs of the worm [96]. Bacillus atrophaeus GBSC56 emitted methyl isovalerate, 2-undecanone, and dimethyl disulfide, which exhibited strong nematicidal activity against M. incognita [21]. B. cereus Bc-cm103 exhibited repellent activity to J2s of M. incognita. In addition, VOCs from Bc-cm103, mainly consisting of dimethyl disulfide and S-methyl ester butanethioic acid, displayed fumigant toxicity to M. incognita J2s and reduced the number of root galls on a cucumber plant in a double-layered pot test [97]. Bacillus aryabhattai MCCC 1K02966 emitted dimethyl disulfide, methyl thioacetate, 1-butanol, and pentane. Among the four VOCs, methyl thioacetate displayed the strongest contact and fumigant toxicity as well as repellent activity against M. incognita [16]. Bacillus altitudinis AMCC 1040 emitted eight VOCs. Of these, acetic acid, octanoic acid, 2-methyl-butanoic acid, 3-methyl-butanoic acid, 2,3-butanedione, and 2-isopropoxy ethylamine had nematicidal activity against M. incognita [98].
Table 1. In vitro activity of bacterial VOCs on Meloidogyne incognita.
Table 1. In vitro activity of bacterial VOCs on Meloidogyne incognita.
VOCEmitterEffects on J2sEgg Hatching Suppression
Contact ToxicityFumigant Activity
AcetaldehydeVirgibacillus dokdonensis MCCC 1A00493 [99][99][99][99] [99]
Acetic acidBacillus altitudinis AMCC 1040 [98][98]
AcetonePaenibacillus polymyxa KM2501-1 [2] [2]
AcetophenonePseudochrobactrum saccharolyticum [100]
Arthrobacter nicotianae [100]
Achromobacter xylosoxidans [100]
4-acetylbenzoicPaenibacillus polymyxa KM2501-1 [2][2]
BenzaldehydeOchrobactrum pseudogrignonense NC1 [101][101]
BenzeneacetaldehydeBacillus megaterium YMF3.25 [96] [96] [96]
2,3-ButanedioneBacillus altitudinis AMCC 1040 [98][98]
2-butanoneVirgibacillus dokdonensis MCCC 1A00493 [99] [99]
Butyl isovalerateWautersiella falsenii [100] [100]
DecanalBacillus megaterium YMF3.25 [96] [96] [96]
2-decanolPaenibacillus polymyxa KM2501-1 [2][2][2][2]
2-decanonePaenibacillus polymyxa KM2501-1 [2][2][2]
Dimethyl disulfidePseudochrobactrum saccharolyticum [100]
Wautersiella falsenii [100]
Proteus hauseri [100]
Arthrobacter nicotianae [100]
Achromobacter xylosoxidans [100]
Bacillus megaterium YMF3.25 [96]
Bacillus atrophaeus GBSC56 [21]
Ochrobactrum pseudogrignonense NC1 [101]
Virgibacillus dokdonensis MCCC 1A00493 [99]
Pseudomonas putida 1A00316 [6]
Bacillus cereus Bc-cm103 [97]
Bacillus aryabhattai MCCC 1K02966 [16]
1-(ethenyloxy)-octadecanePseudomonas putida 1A00316 [6] [6][6]
EthylbenzeneVirgibacillus dokdonensis MCCC 1A00493 [99] [99]
Ethyl 3,3-dimethylacrylatePseudochrobactrum saccharolyticum [100] [100]
Furfural acetonePaenibacillus polymyxa KM2501-1 [2][2][2][2]
(Z)-hexen-1-ol acetatePseudomonas putida 1A00316 [6][6] [6][6]
2-Isopropoxy ethylamineBacillus altitudinis AMCC 1040 [98][98]
1-methoxy-4-methylbenzeneWautersiella falsenii [100]
Proteus hauseri [100]
Achromobacter xylosoxidans [100]
2-Methyl-butanoic acidBacillus altitudinis AMCC 1040
3-Methyl-butanoic acidBacillus altitudinis AMCC 1040
Methyl isovalerateBacillus atrophaeus GBSC56 [21][21]
Methyl thioacetateBacillus aryabhattai MCCC 1K02966 [16][16][16] [16][16]
S-methyl thiobutyratePseudochrobactrum saccharolyticum [100]
Wautersiella falsenii [100]
Proteus hauseri [100]
Arthrobacter nicotianae [100]
Achromobacter xylosoxidans [100]
2-nonanolPaenibacillus polymyxa KM2501-1 [2][2][2]
2-nonanonePseudochrobactrum saccharolyticum [100]
Wautersiella falsenii [100]
Proteus hauseri [100]
Achromobacter xylosoxidans [100]
Bacillus megaterium YMF3.25 [96]
Paenibacillus polymyxa KM2501-1 [2]
Pseudomonas putida 1A00316 [6]
[2,6][96,100] [6][6,96]
Octanoic acidBacillus altitudinis AMCC 1040 [98][98]
2-octanonePseudomonas putida 1A00316 [6][6] [6][6]
2-undecanolPaenibacillus polymyxa KM2501-1 [2][2][2]
2-undecanoneBacillus megaterium YMF3.25 [96]
Bacillus atrophaeus GBSC56 [21]
Pseudomonas putida 1A00316 [6]
Paenibacillus polymyxa KM2501-1 [2]
[2,6,21][2,6,96] [2,6][6,96]
1-undecenePseudomonas putida 1A00316 [6] [6][6]
Paenibacillus polymyxa KM2501-1 caused 87.6% and 82.6% mortality of M. incognita under both in vitro and in planta conditions, respectively. Eleven VOCs were emitted by P. polymyxa KM2501-1. Among them, furfural acetone and 2-decanol could attract M. incognita and then kill the worm by acting as fumigants or contact nematicides [2]. VOCs produced by Virgibacillus dokdonensis MCCC 1A00493 displayed several activities against M. incognita. Acetaldehyde acted as an attractant, contact nematicide, and fumigant, whereas ethylbenzene acted as an attractant and 2-butanone as a repellent [99].
Pseudomonas putida strain 1A00316, isolated from Antarctic soil, emitted 2-nonanone, 2-octanone, 2-undecanone, dimethyl disulfide, (Z)-hexen-1-ol acetate, 1-undecene, and 1-(ethenyloxy)-octadecane. Of these, 2-nonanone, 2-octanone, 2-undecanone, dimethyl disulfide, and (Z)-hexen-1-ol acetate showed contact nematicidal activity against M. incognita; however, only 2-undecanone exhibited fumigant activity. In addition, all seven VOCs suppressed egg hatching and showed repellent activity to M. incognita J2s in Petri plate experiments [6].
In total, 53 VOCs were identified from five bacteria, namely, Pseudochrobactrum saccharolyticum, Wautersiella falsenii, Proteus hauseri, Arthrobacter nicotianae, and Achromobacter xylosoxidans. Among the VOCs, S-methyl thiobutyrate, dimethyl disulfide, acetophenone, 2-nonanone, butyl isovalerate, ethyl 3,3-dimethylacrylate, and 1-methoxy-4-methylbenzene, exhibited significant nematicidal activity against both C. elegans and M. incognita in Petri plate experiments. Moreover, S-methyl thiobutyrate was the most active VOC [100]. Ochrobactrum pseudogrignonense NC1 significantly inhibited M. incognita in Petri plate and greenhouse trials. The main VOCs emitted by NC1, namely, dimethyl disulfide and benzaldehyde, also had nematicidal activity against M. incognita [101].
Besides M. incognita, some reports addressed the microbial fumigant toxicity to other Meloidogyne species. Three bacterial strains (Bacillus sp., Paenibacillus sp., and Xanthomonas sp.) emitted VOCs that were toxic to rice RKN Meloidogyne graminicola in both in vitro and in planta studies [102]. In vitro treatment with P. putida, Microbacterium sp., Bacillus methylotrophicus, and Bacillus pumilus caused significant mortality of M. exigua via the release of VOCs [103]. Variovorax paradoxus, Comamonas sediminis, Pseudomonas soli, Pseudomonas koreensis, and two strains of Pseudomonas monteilii were reported to exhibit nematicidal activity. They showed strong virulent effects on M. javanica through the production of VOCs [104].
Among the microbial VOCs identified thus far, dimethyl disulfide is the most commonly identified. In light of its toxicity to a broad spectrum of pests, dimethyl disulfide was registered by Arkema as a pesticide by the name of Paladin in 2012 [105].

4.2. Mechanism of Action of Bacterial VOCs

It is thought that VOCs may destroy nematodes by targeting the intestine, nervous system, surface coat, pharynx, or other tissues [2,17,106]. A recent study has claimed that VOCs cause rapid death by inducing severe oxidative stress in nematodes [21]. However, the detailed molecular mechanisms underlying the nematicidal activity of VOCs are poorly understood, with a few exceptions. A well-studied VOC, dimethyl disulfide, exerts its toxicity by blocking the activity of the enzyme cytochrome oxidase, consequently stopping the mitochondrial respiration of the pests [105].
Bacterial VOCs have also been reported to regulate the key genes involved in different signaling pathways by which plant growth is stimulated and induced systemic resistance against phytopathogens is triggered. For example, methyl isovalerate and 2-undecanone promoted plant growth and stimulated induced systemic resistance by enhancing the antioxidant enzyme activity in plant roots infested with M. incognita [21]. The effects of bacterial VOCs on plant morphology and physiology are discussed in a recent review paper [107].

5. Concluding Remarks and Future Perspectives

Driven by the concerns about the negative impacts of chemical nematicides on human health and the environment, there has been a surge of interest in the development of sustainable methods to replace the chemical strategy of controlling RKNs. A large number of reports have demonstrated that microorganisms constitute a rich source for the discovery of potentially useful VOCs in the control of RKNs. Although most of the data came from in vitro tests, some were from in planta experiments performed in greenhouse conditions. However, extensive investigations are needed to confirm whether VOCs are also effective against RKNs in open fields.
Dimethyl disulfide represents a successfully commercialized VOC, which not only is emitted by a broad spectrum of bacteria but is also effective for the control of a variety of pests. Some other VOCs, particularly the sulfur-containing ones, such as S-methyl thiobutyrate and S-methyl thioacetate, are also promising candidates because of their strong toxicity to nematodes. Further assessments of their potential in agricultural practice should be encouraged.
Investigations into how nematodes are affected by VOCs at the molecular level are still rare. Since the chemical nature of VOCs is diverse, each type of VOC might have its own mode of action. The answer to this query is not only of interest for academic research purposes but is also crucial for the development of VOCs for nematode control in the future.

Author Contributions

Conceptualization, A.D. and M.M.; data curation, A.D.; writing—original draft preparation, A.D. and M.O.; writing—review and editing, M.M.; project administration, A.D.; funding acquisition, M.M. All authors have read and agreed to the published version of the manuscript.


This research was funded by the Ministry of Science and Technology, Taiwan, under the grant number MOST 110-2823-8-005-001.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Abad, P.; Gouzy, J.; Aury, J.M.; Castagnone-Sereno, P.; Danchin, E.G.; Deleury, E.; Perfus-Barbeoch, L.; Anthouard, V.; Artiguenave, F.; Blok, V.C.; et al. Genome sequence of the metazoan plant-parasitic nematode Meloidogyne incognita. Nat. Biotechnol. 2008, 26, 909–915. [Google Scholar] [CrossRef][Green Version]
  2. Cheng, W.; Yang, J.; Nie, Q.; Huang, D.; Yu, C.; Zheng, L.; Cai, M.; Thomashow, L.S.; Weller, D.M.; Yu, Z.; et al. Volatile organic compounds from Paenibacillus polymyxa KM2501-1 control Meloidogyne incognita by multiple strategies. Sci. Rep. 2017, 7, 16213. [Google Scholar] [CrossRef] [PubMed]
  3. Lu, H.; Xu, S.; Zhang, W.; Xu, C.; Li, B.; Zhang, D.; Mu, W.; Liu, F. Nematicidal activity of trans-2-hexenal against southern root-knot nematode (Meloidogyne incognita) on tomato plants. J. Agric. Food Chem. 2017, 65, 544–550. [Google Scholar] [CrossRef]
  4. Poveda, J.; Abril-Urias, P.; Escobar, C. Biological control of plant-parasitic nematodes by filamentous fungi inducers of resistance: Trichoderma, mycorrhizal and endophytic fungi. Front. Microbiol. 2020, 11, 992. [Google Scholar] [CrossRef]
  5. Kyndt, T.; Fernandez, D.; Gheysen, G. Plant-parasitic nematode infections in rice: Molecular and cellular insights. Annu. Rev. Phytopathol. 2014, 52, 135–153. [Google Scholar] [CrossRef] [PubMed]
  6. Zhai, Y.; Shao, Z.; Cai, M.; Zheng, L.; Li, G.; Huang, D.; Cheng, W.; Thomashow, L.S.; Weller, D.M.; Yu, Z.; et al. Multiple modes of nematode control by volatiles of Pseudomonas putida 1A00316 from Antarctic soil against Meloidogyne incognita. Front. Microbiol. 2018, 9, 253. [Google Scholar] [CrossRef] [PubMed][Green Version]
  7. Li, J.; Zou, C.; Xu, J.; Ji, X.; Niu, X.; Yang, J.; Huang, X.; Zhang, K.Q. Molecular mechanisms of nematode-nematophagous microbe interactions: Basis for biological control of plant-parasitic nematodes. Annu. Rev. Phytopathol. 2015, 53, 67–95. [Google Scholar] [CrossRef] [PubMed]
  8. Abad, P.; Favery, B.; Rosso, M.N.; Castagnone-Sereno, P. Root-knot nematode parasitism and host response: Molecular basis of a sophisticated interaction. Mol. Plant Pathol. 2003, 4, 217–224. [Google Scholar] [CrossRef] [PubMed]
  9. Jones, J.T.; Haegeman, A.; Danchin, E.G.; Gaur, H.S.; Helder, J.; Jones, M.G.; Kikuchi, T.; Manzanilla-Lopez, R.; Palomares-Rius, J.E.; Wesemael, W.M.; et al. Top 10 plant-parasitic nematodes in molecular plant pathology. Mol. Plant Pathol. 2013, 14, 946–961. [Google Scholar] [CrossRef] [PubMed]
  10. Kihika, R.; Murungi, L.K.; Coyne, D.; Ng’ang’a, M.; Hassanali, A.; Teal, P.E.A.; Torto, B. Parasitic nematode Meloidogyne incognita interactions with different Capsicum annum cultivars reveal the chemical constituents modulating root herbivory. Sci. Rep. 2017, 7, 2903. [Google Scholar] [CrossRef]
  11. Abdel-Rahman, F.H.; Alaniz, N.M.; Saleh, M.A. Nematicidal activity of terpenoids. J. Environ. Sci. Health B 2013, 48, 16–22. [Google Scholar] [CrossRef]
  12. Riga, E. The effects of Brassica green manures on plant parasitic and free living nematodes used in combination with reduced rates of synthetic nematicides. J. Nematol. 2011, 43, 119–121. [Google Scholar]
  13. Schneider, S.M.; Rosskopf, E.N.; Leesch, J.G.; Chellemi, D.O.; Bull, C.T.; Mazzola, M. United States Department of Agriculture-Agricultural Research Service research on alternatives to methyl bromide: Pre-plant and post-harvest. Pest Manag. Sci. 2003, 59, 814–826. [Google Scholar] [CrossRef]
  14. Zasada, I.A.; Halbrendt, J.M.; Kokalis-Burelle, N.; LaMondia, J.; McKenry, M.V.; Noling, J.W. Managing nematodes without methyl bromide. Annu. Rev. Phytopathol. 2010, 48, 311–328. [Google Scholar] [CrossRef][Green Version]
  15. Tian, B.; Yang, J.; Zhang, K.Q. Bacteria used in the biological control of plant-parasitic nematodes: Populations, mechanisms of action, and future prospects. FEMS Microbiol. Ecol. 2007, 61, 197–213. [Google Scholar] [CrossRef][Green Version]
  16. Chen, W.; Wang, J.; Huang, D.; Cheng, W.; Shao, Z.; Cai, M.; Zheng, L.; Yu, Z.; Zhang, J. Volatile organic compounds from Bacillus aryabhattai MCCC 1K02966 with multiple modes against Meloidogyne incognita. Molecules 2021, 27, 103. [Google Scholar] [CrossRef]
  17. Geng, C.; Nie, X.; Tang, Z.; Zhang, Y.; Lin, J.; Sun, M.; Peng, D. A novel serine protease, Sep1, from Bacillus firmus DS-1 has nematicidal activity and degrades multiple intestinal-associated nematode proteins. Sci. Rep. 2016, 6, 25012. [Google Scholar] [CrossRef][Green Version]
  18. Meyer, S.L. United States Department of Agriculture-Agricultural Research Service research programs on microbes for management of plant-parasitic nematodes. Pest Manag. Sci. 2003, 59, 665–670. [Google Scholar] [CrossRef]
  19. Rajaofera, M.J.N.; Wang, Y.; Dahar, G.Y.; Jin, P.; Fan, L.; Xu, L.; Liu, W.; Miao, W. Volatile organic compounds of Bacillus atrophaeus HAB-5 inhibit the growth of Colletotrichum gloeosporioides. Pestic. Biochem. Physiol. 2019, 156, 170–176. [Google Scholar] [CrossRef]
  20. Syed-Ab-Rahman, S.F.; Carvalhais, L.C.; Chua, E.T.; Chung, F.Y.; Moyle, P.M.; Eltanahy, E.G.; Schenk, P.M. Soil bacterial diffusible and volatile organic compounds inhibit Phytophthora capsici and promote plant growth. Sci. Total Environ. 2019, 692, 267–280. [Google Scholar] [CrossRef]
  21. Ayaz, M.; Ali, Q.; Farzand, A.; Khan, A.R.; Ling, H.; Gao, X. Nematicidal volatiles from Bacillus atrophaeus GBSC56 promote growth and stimulate induced systemic resistance in tomato against Meloidogyne incognita. Int. J. Mol. Sci. 2021, 22, 5049. [Google Scholar] [CrossRef] [PubMed]
  22. El-Sappah, A.H.; Islam, M.; El-awady, H.; Yan, S.; Qi, S.; Liu, J.; Cheng, G.T.; Liang, Y. Tomato natural resistance genes in controlling the root-knot nematode. Genes 2019, 10, 925. [Google Scholar] [CrossRef] [PubMed][Green Version]
  23. Gao, H.; Qi, G.; Yin, R.; Zhang, H.; Li, C.; Zhao, X. Bacillus cereus strain S2 shows high nematicidal activity against Meloidogyne incognita by producing sphingosine. Sci. Rep. 2016, 6, 28756. [Google Scholar] [CrossRef] [PubMed]
  24. Nicol, J.M.; Turner, S.J.; Coyne, D.L.; Nijs, L.d.; Hockland, S.; Maafi, Z.T. Current nematode threats to world agriculture. In Genomics and Molecular Genetics of Plant-Nematode Interactions; Jones, J., Gheysen, G., Fenoll, C., Eds.; Springer: Dordrecht, The Netherlands, 2011; pp. 21–43. [Google Scholar] [CrossRef]
  25. Lima, F.S.; Correa, V.R.; Nogueira, S.R.; Santos, P.R. Nematodes affecting soybean and sustainable practices for their management. In Soybean—The Basis of Yield, Biomass and Productivity; Kasai, M., Ed.; IntechOpen: London, UK, 2017. [Google Scholar] [CrossRef][Green Version]
  26. Elling, A.A. Major emerging problems with minor Meloidogyne species. Phytopathology 2013, 103, 1092–1102. [Google Scholar] [CrossRef][Green Version]
  27. Coyne, D.L.; Cortada, L.; Dalzell, J.J.; Claudius-Cole, A.O.; Haukeland, S.; Luambano, N.; Talwana, H. Plant-parasitic nematodes and food security in Sub-Saharan Africa. Annu. Rev. Phytopathol. 2018, 56, 381–403. [Google Scholar] [CrossRef][Green Version]
  28. Janssen, T.; Karssen, G.; Verhaeven, M.; Coyne, D.; Bert, W. Mitochondrial coding genome analysis of tropical root-knot nematodes (Meloidogyne) supports haplotype based diagnostics and reveals evidence of recent reticulate evolution. Sci. Rep. 2016, 6, 22591. [Google Scholar] [CrossRef][Green Version]
  29. Tapia-Vazquez, I.; Montoya-Martinez, A.C.; De Los Santos-Villalobos, S.; Ek-Ramos, M.J.; Montesinos-Matias, R.; Martinez-Anaya, C. Root-knot nematodes (Meloidogyne spp.) a threat to agriculture in Mexico: Biology, current control strategies, and perspectives. World J. Microbiol. Biotechnol. 2022, 38, 26. [Google Scholar] [CrossRef]
  30. Gheysen, G.; Mitchum, M.G. How nematodes manipulate plant development pathways for infection. Curr. Opin. Plant Biol. 2011, 14, 415–421. [Google Scholar] [CrossRef]
  31. Castagnone-Sereno, P. Genetic variability and adaptive evolution in parthenogenetic root-knot nematodes. Heredity 2006, 96, 282–289. [Google Scholar] [CrossRef][Green Version]
  32. Castagnone-Sereno, P.; Danchin, E.G.; Perfus-Barbeoch, L.; Abad, P. Diversity and evolution of root-knot nematodes, genus Meloidogyne: New insights from the genomic era. Annu. Rev. Phytopathol. 2013, 51, 203–220. [Google Scholar] [CrossRef]
  33. Saucet, S.B.; Van Ghelder, C.; Abad, P.; Duval, H.; Esmenjaud, D. Resistance to root-knot nematodes Meloidogyne spp. in woody plants. New Phytol. 2016, 211, 41–56. [Google Scholar] [CrossRef][Green Version]
  34. Caillaud, M.C.; Dubreuil, G.; Quentin, M.; Perfus-Barbeoch, L.; Lecomte, P.; de Almeida Engler, J.; Abad, P.; Rosso, M.N.; Favery, B. Root-knot nematodes manipulate plant cell functions during a compatible interaction. J. Plant Physiol. 2008, 165, 104–113. [Google Scholar] [CrossRef]
  35. Opperman, C.H.; Bird, D.M.; Williamson, V.M.; Rokhsar, D.S.; Burke, M.; Cohn, J.; Cromer, J.; Diener, S.; Gajan, J.; Graham, S.; et al. Sequence and genetic map of Meloidogyne hapla: A compact nematode genome for plant parasitism. Proc. Natl. Acad. Sci. USA 2008, 105, 14802–14807. [Google Scholar] [CrossRef][Green Version]
  36. Szitenberg, A.; Salazar-Jaramillo, L.; Blok, V.C.; Laetsch, D.R.; Joseph, S.; Williamson, V.M.; Blaxter, M.L.; Lunt, D.H. Comparative genomics of apomictic root-knot nematodes: Hybridization, ploidy, and dynamic genome change. Genome Biol. Evol. 2017, 9, 2844–2861. [Google Scholar] [CrossRef][Green Version]
  37. Danchin, E.G.; Rosso, M.N.; Vieira, P.; de Almeida-Engler, J.; Coutinho, P.M.; Henrissat, B.; Abad, P. Multiple lateral gene transfers and duplications have promoted plant parasitism ability in nematodes. Proc. Natl. Acad. Sci. USA 2010, 107, 17651–17656. [Google Scholar] [CrossRef][Green Version]
  38. Rybarczyk-Mydlowska, K.; Maboreke, H.R.; van Megen, H.; van den Elsen, S.; Mooyman, P.; Smant, G.; Bakker, J.; Helder, J. Rather than by direct acquisition via lateral gene transfer, GHF5 cellulases were passed on from early Pratylenchidae to root-knot and cyst nematodes. BMC Evol. Biol. 2012, 12, 221. [Google Scholar] [CrossRef][Green Version]
  39. Haegeman, A.; Jones, J.T.; Danchin, E.G. Horizontal gene transfer in nematodes: A catalyst for plant parasitism? Mol. Plant Microbe Interact. 2011, 24, 879–887. [Google Scholar] [CrossRef][Green Version]
  40. Sun, X.; Zhang, X.; Zhang, S.; Dai, G.; Han, S.; Liang, W. Soil nematode responses to increases in nitrogen deposition and precipitation in a temperate forest. PLoS ONE 2013, 8, e82468. [Google Scholar] [CrossRef][Green Version]
  41. Armenteros, M.; Rodriguez-Garcia, P.; Perez-Garcia, J.A.; Gracia, A. Diversity patterns of free-living nematode assemblages in seagrass beds from the Cuban archipelago (Caribbean Sea). Biodivers. Data J. 2020, 8, e58848. [Google Scholar] [CrossRef]
  42. Chen, J.; Li, Q.X.; Song, B. Chemical nematicides: Recent research progress and outlook. J. Agric. Food Chem. 2020, 68, 12175–12188. [Google Scholar] [CrossRef]
  43. Jang, J.Y.; Le Dang, Q.; Choi, G.J.; Park, H.W.; Kim, J.C. Control of root-knot nematodes using Waltheria indica producing 4-quinolone alkaloids. Pest Manag. Sci. 2019, 75, 2264–2270. [Google Scholar] [CrossRef] [PubMed]
  44. Bali, S.; Zhang, L.; Franco, J.; Gleason, C. Biotechnological advances with applicability in potatoes for resistance against root-knot nematodes. Curr. Opin. Biotechnol. 2021, 70, 226–233. [Google Scholar] [CrossRef] [PubMed]
  45. Park, M.G.; Choi, J.; Hong, Y.S.; Park, C.G.; Kim, B.G.; Lee, S.Y.; Lim, H.J.; Mo, H.H.; Lim, E.; Cha, W. Negative effect of methyl bromide fumigation work on the central nervous system. PLoS ONE 2020, 15, e0236694. [Google Scholar] [CrossRef]
  46. Gharibi, H.; Entwistle, M.R.; Schweizer, D.; Tavallali, P.; Thao, C.; Cisneros, R. Methyl-bromide and asthma emergency department visits in California, USA from 2005 to 2011. J. Asthma 2020, 57, 1227–1236. [Google Scholar] [CrossRef]
  47. Bulathsinghala, A.T.; Shaw, I.C. The toxic chemistry of methyl bromide. Hum. Exp. Toxicol. 2014, 33, 81–91. [Google Scholar] [CrossRef] [PubMed]
  48. Methyl Bromide. In Encyclopedia of Toxicology, 3rd ed.; Weber, D.V.A.M. (Ed.) Elsevier: Amsterdam, The Netherlands, 2014; pp. 270–273. [Google Scholar]
  49. Yoshida, S.; Yamada, H.; Sugawara, I.; Takeda, K. Effect of dibromochloropropane (DBCP) on the hormone receptors of the male rat reproductive system. Biosci. Biotechnol. Biochem. 1998, 62, 479–483. [Google Scholar] [CrossRef] [PubMed][Green Version]
  50. Krzastek, S.C.; Farhi, J.; Gray, M.; Smith, R.P. Impact of environmental toxin exposure on male fertility potential. Transl. Androl. Urol. 2020, 9, 2797–2813. [Google Scholar] [CrossRef] [PubMed]
  51. Clark, H.A.; Snedeker, S.M. Critical evaluation of the cancer risk of dibromochloropropane (DBCP). J. Environ. Sci. Health C Environ. Carcinog. Ecotoxicol. Rev. 2005, 23, 215–260. [Google Scholar] [CrossRef]
  52. United States Congress; Senate Committee on Foreign Relations. Amendment to the Montreal Protocol on Substances That Deplete the Ozone Layer: Report. 1993. Available online: (accessed on 14 June 2022).
  53. Oliver, J.E. Montreal Protocol. In Encyclopedia of World Climatology; Oliver, J.E., Ed.; Springer: Dordrecht, The Netherlands, 2005; p. 516. [Google Scholar] [CrossRef]
  54. Qiao, K.; Shi, X.; Wang, H.; Ji, X.; Wang, K. Managing root-knot nematodes and weeds with 1,3-dichloropropene as an alternative to methyl bromide in cucumber crops in China. J. Agric. Food Chem. 2011, 59, 2362–2367. [Google Scholar] [CrossRef]
  55. Martin, F.N. Development of alternative strategies for management of soilborne pathogens currently controlled with methyl bromide. Annu. Rev. Phytopathol. 2003, 41, 325–350. [Google Scholar] [CrossRef][Green Version]
  56. Holmes, G.J.; Mansouripour, S.M.; Hewavitharana, S.S. Strawberries at the Crossroads: Management of soilborne diseases in California without methyl bromide. Phytopathology 2020, 110, 956–968. [Google Scholar] [CrossRef][Green Version]
  57. Eilenberg, J.; Hajek, A.; Lomer, C. Suggestions for unifying the terminology in biological control. BioControl 2001, 46, 387–400. [Google Scholar] [CrossRef]
  58. Leneveu-Jenvrin, C.; Charles, F.; Barba, F.J.; Remize, F. Role of biological control agents and physical treatments in maintaining the quality of fresh and minimally-processed fruit and vegetables. Crit. Rev. Food Sci. Nutr. 2020, 60, 2837–2855. [Google Scholar] [CrossRef]
  59. Sethuraman, A.; Janzen, F.J.; Weisrock, D.W.; Obrycki, J.J. Insights from population genomics to enhance and sustain biological control of insect pests. Insects 2020, 11, 462. [Google Scholar] [CrossRef]
  60. Singh, J. Natural bioactive products in sustainable agriculture. In Natural Bioactive Products in Sustainable Agriculture; Joginder Singh, A.N.Y., Ed.; Springer Nature: Berlin/Heidelberg, Germany, 2020. [Google Scholar] [CrossRef]
  61. Lahlali, R.; Ezrari, S.; Radouane, N.; Kenfaoui, J.; Esmaeel, Q.; El Hamss, H.; Belabess, Z.; Barka, E.A. Biological control of plant pathogens: A global perspective. Microorganisms 2022, 10, 596. [Google Scholar] [CrossRef]
  62. Tilocca, B.; Cao, A.; Migheli, Q. Scent of a killer: Microbial volatilome and its role in the biological control of plant pathogens. Front. Microbiol. 2020, 11, 41. [Google Scholar] [CrossRef][Green Version]
  63. Forghani, F.; Hajihassani, A. Recent advances in the development of environmentally benign treatments to control root-knot nematodes. Front. Plant Sci. 2020, 11, 1125. [Google Scholar] [CrossRef]
  64. Manan, A.; Bazai, Z.A.; Fan, J.; Yu, H.; Li, L. The Nif3-family protein YqfO03 from Pseudomonas syringae MB03 has multiple nematicidal activities against Caenorhabditis elegans and Meloidogyne incognita. Int. J. Mol. Sci. 2018, 19, 3915. [Google Scholar] [CrossRef][Green Version]
  65. Jamal, Q.; Cho, J.Y.; Moon, J.H.; Munir, S.; Anees, M.; Kim, K.Y. Identification for the first time of Cyclo(d-Pro-l-Leu) produced by Bacillus amyloliquefaciens Y1 as a nematocide for control of Meloidogyne incognita. Molecules 2017, 22, 1839. [Google Scholar] [CrossRef][Green Version]
  66. Adam, M.; Heuer, H.; Hallmann, J. Bacterial antagonists of fungal pathogens also control root-knot nematodes by induced systemic resistance of tomato plants. PLoS ONE 2014, 9, e90402. [Google Scholar] [CrossRef][Green Version]
  67. Oliveira, D.F.; Santos Junior, H.M.; Nunes, A.S.; Campos, V.P.; Pinho, R.S.; Gajo, G.C. Purification and identification of metabolites produced by Bacillus cereus and B. subtilis active against Meloidogyne exigua, and their in silico interaction with a putative phosphoribosyltransferase from M. incognita. An. Acad. Bras. Cienc. 2014, 86, 525–538. [Google Scholar] [CrossRef] [PubMed][Green Version]
  68. Zhang, L.; Gleason, C. Enhancing potato resistance against root-knot nematodes using a plant-defence elicitor delivered by bacteria. Nat. Plants 2020, 6, 625–629. [Google Scholar] [CrossRef] [PubMed]
  69. Migunova, V.D.; Tomashevich, N.S.; Konrat, A.N.; Lychagina, S.V.; Dubyaga, V.M.; D’Addabbo, T.; Sasanelli, N.; Asaturova, A.M. Selection of bacterial strains for control of root-knot disease caused by Meloidogyne incognita. Microorganisms 2021, 9, 1698. [Google Scholar] [CrossRef] [PubMed]
  70. Koch, M.S.; Ward, J.M.; Levine, S.L.; Baum, J.A.; Vicini, J.L.; Hammond, B.G. The food and environmental safety of Bt crops. Front. Plant Sci. 2015, 6, 283. [Google Scholar] [CrossRef]
  71. Zhang, F.; Peng, D.; Ye, X.; Yu, Z.; Hu, Z.; Ruan, L.; Sun, M. In vitro uptake of 140 kDa Bacillus thuringiensis nematicidal crystal proteins by the second stage juvenile of Meloidogyne hapla. PLoS ONE 2012, 7, e38534. [Google Scholar] [CrossRef] [PubMed]
  72. Guo, S.; Liu, M.; Peng, D.; Ji, S.; Wang, P.; Yu, Z.; Sun, M. New strategy for isolating novel nematicidal crystal protein genes from Bacillus thuringiensis strain YBT-1518. Appl. Environ. Microbiol. 2008, 74, 6997–7001. [Google Scholar] [CrossRef] [PubMed][Green Version]
  73. Wei, J.Z.; Hale, K.; Carta, L.; Platzer, E.; Wong, C.; Fang, S.C.; Aroian, R.V. Bacillus thuringiensis crystal proteins that target nematodes. Proc. Natl. Acad. Sci. USA 2003, 100, 2760–2765. [Google Scholar] [CrossRef] [PubMed][Green Version]
  74. Li, X.Q.; Wei, J.Z.; Tan, A.; Aroian, R.V. Resistance to root-knot nematode in tomato roots expressing a nematicidal Bacillus thuringiensis crystal protein. Plant Biotechnol. J. 2007, 5, 455–464. [Google Scholar] [CrossRef]
  75. Yu, Z.; Xiong, J.; Zhou, Q.; Luo, H.; Hu, S.; Xia, L.; Sun, M.; Li, L.; Yu, Z. The diverse nematicidal properties and biocontrol efficacy of Bacillus thuringiensis Cry6A against the root-knot nematode Meloidogyne hapla. J. Invertebr. Pathol. 2015, 125, 73–80. [Google Scholar] [CrossRef]
  76. Geng, C.; Liu, Y.; Li, M.; Tang, Z.; Muhammad, S.; Zheng, J.; Wan, D.; Peng, D.; Ruan, L.; Sun, M. Dissimilar crystal proteins Cry5Ca1 and Cry5Da1 synergistically act against Meloidogyne incognita and delay Cry5Ba-based nematode resistance. Appl. Environ. Microbiol. 2017, 83, e03505-16. [Google Scholar] [CrossRef][Green Version]
  77. Liu, C.; Timper, P.; Ji, P.; Mekete, T.; Joseph, S. Influence of root exudates and soil on attachment of Pasteuria penetrans to Meloidogyne arenaria. J. Nematol. 2017, 49, 304–310. [Google Scholar] [CrossRef][Green Version]
  78. Kokalis-Burelle, N. Pasteuria penetrans for Control of Meloidogyne incognita on tomato and cucumber, and M. arenaria on Snapdragon. J. Nematol. 2015, 47, 207–213. [Google Scholar]
  79. Phani, V.; Somvanshi, V.S.; Rao, U. Silencing of a Meloidogyne incognita selenium-binding protein alters the cuticular adhesion of Pasteuria penetrans endospores. Gene 2018, 677, 289–298. [Google Scholar] [CrossRef]
  80. Sun, X.; Zhang, R.; Ding, M.; Liu, Y.; Li, L. Biocontrol of the root-knot nematode Meloidogyne incognita by a nematicidal bacterium Pseudomonas simiae MB751 with cyclic dipeptide. Pest Manag. Sci. 2021, 77, 4365–4374. [Google Scholar] [CrossRef]
  81. Meyer, S.; Roberts, D.; Chitwood, D.; Carta, L.; Lumsden, R.; Mao, W. Application of Burkholderia cepacia and Trichoderma virens, alone and in combinations, against Meloidogyne incognita on Bell pepper. Nematropica 2001, 31, 75–86. [Google Scholar]
  82. Rahul, S.; Chandrashekhar, P.; Hemant, B.; Chandrakant, N.; Laxmikant, S.; Satish, P. Nematicidal activity of microbial pigment from Serratia marcescens. Nat. Prod. Res. 2014, 28, 1399–1404. [Google Scholar] [CrossRef]
  83. Sharifi, R.; Ryu, C.M. Sniffing bacterial volatile compounds for healthier plants. Curr. Opin. Plant Biol. 2018, 44, 88–97. [Google Scholar] [CrossRef]
  84. Monson, R.K. Reactions of biogenic volatile organic compounds in the atmosphere. In The Chemistry and Biology of Volatiles; John Wiley & Sons: Hoboken, NJ, USA, 2010; pp. 363–388. [Google Scholar] [CrossRef]
  85. Kessler, A.; Morrell, K. Plant volatile signalling: Multitrophic interactions in the headspace. In The Chemistry and Biology of Volatiles; John Wiley & Sons: Hoboken, NJ, USA, 2010; pp. 95–122. [Google Scholar] [CrossRef]
  86. Schalchli, H.; Tortella, G.R.; Rubilar, O.; Parra, L.; Hormazabal, E.; Quiroz, A. Fungal volatiles: An environmentally friendly tool to control pathogenic microorganisms in plants. Crit. Rev. Biotechnol. 2016, 36, 144–152. [Google Scholar] [CrossRef]
  87. Mercier, J.; Jiménez, J.I. Control of fungal decay of apples and peaches by the biofumigant fungus Muscodor albus. Postharvest Biol. Technol. 2004, 31, 1–8. [Google Scholar] [CrossRef]
  88. Song, G.C.; Ryu, C.M. Two volatile organic compounds trigger plant self-defense against a bacterial pathogen and a sucking insect in cucumber under open field conditions. Int. J. Mol. Sci. 2013, 14, 9803–9819. [Google Scholar] [CrossRef][Green Version]
  89. Parafati, L.; Vitale, A.; Restuccia, C.; Cirvilleri, G. Performance evaluation of volatile organic compounds by antagonistic yeasts immobilized on hydrogel spheres against gray, green and blue postharvest decays. Food Microbiol. 2017, 63, 191–198. [Google Scholar] [CrossRef] [PubMed]
  90. Audrain, B.; Farag, M.A.; Ryu, C.M.; Ghigo, J.M. Role of bacterial volatile compounds in bacterial biology. FEMS Microbiol. Rev. 2015, 39, 222–233. [Google Scholar] [CrossRef] [PubMed][Green Version]
  91. Lemfack, M.C.; Gohlke, B.O.; Toguem, S.M.T.; Preissner, S.; Piechulla, B.; Preissner, R. mVOC 2.0: A database of microbial volatiles. Nucleic Acids Res. 2018, 46, D1261–D1265. [Google Scholar] [CrossRef] [PubMed][Green Version]
  92. Lin, Y.T.; Lee, C.C.; Leu, W.M.; Wu, J.J.; Huang, Y.C.; Meng, M. Fungicidal Activity of volatile organic compounds emitted by Burkholderia gladioli Strain BBB-01. Molecules 2021, 26, 745. [Google Scholar] [CrossRef]
  93. Bui, H.X.; Desaeger, J.A. Volatile compounds as potential bio-fumigants against plant-parasitic nematodes—A mini review. J. Nematol. 2021, 53, 1–12. [Google Scholar] [CrossRef]
  94. Lazazzara, V.; Perazzolli, M.; Pertot, I.; Biasioli, F.; Puopolo, G.; Cappellin, L. Growth media affect the volatilome and antimicrobial activity against Phytophthora infestans in four Lysobacter type strains. Microbiol. Res. 2017, 201, 52–62. [Google Scholar] [CrossRef]
  95. Schulz-Bohm, K.; Martin-Sanchez, L.; Garbeva, P. Microbial Volatiles: Small molecules with an important role in intra- and inter-kingdom interactions. Front. Microbiol. 2017, 8, 2484. [Google Scholar] [CrossRef]
  96. Huang, Y.; Xu, C.; Ma, L.; Zhang, K.; Duan, C.; Mo, M. Characterisation of volatiles produced from Bacillus megaterium YFM3.25 and their nematicidal activity against Meloidogyne incognita. Eur. J. Plant Pathol. 2010, 126, 417–422. [Google Scholar] [CrossRef]
  97. Yin, N.; Liu, R.; Zhao, J.L.; Khan, R.A.A.; Li, Y.; Ling, J.; Liu, W.; Yang, Y.H.; Xie, B.Y.; Mao, Z.C. Volatile organic compounds of Bacillus cereus strain Bc-cm103 exhibit fumigation activity against Meloidogyne incognita. Plant Dis. 2021, 105, 904–911. [Google Scholar] [CrossRef]
  98. Zhou, B.; Ye, L.; Wang, J.-Y.; Liu, X.-F.; Guan, Q.; Dou, N.-X.; Li, J.; Zhang, Q.; Gao, Y.-M.; Wang, M.; et al. Nematicidal activity of volatile organic compounds produced by Bacillus altitudinis AMCC 1040 against Meloidogyne incognita. Eur. PMC 2022. [Google Scholar] [CrossRef]
  99. Huang, D.; Yu, C.; Shao, Z.; Cai, M.; Li, G.; Zheng, L.; Yu, Z.; Zhang, J. Identification and Characterization of nematicidal volatile organic compounds from deep-sea Virgibacillus dokdonensis MCCC 1A00493. Molecules 2020, 25, 744. [Google Scholar] [CrossRef][Green Version]
  100. Xu, Y.Y.; Lu, H.; Wang, X.; Zhang, K.Q.; Li, G.H. Effect of volatile organic compounds from bacteria on nematodes. Chem. Biodivers. 2015, 12, 1415–1421. [Google Scholar] [CrossRef]
  101. Yang, T.; Xin, Y.; Liu, T.; Li, Z.; Liu, X.; Wu, Y.; Wang, M.; Xiang, M. Bacterial volatile-mediated suppression of root-knot nematode (Meloidogyne incognita). Plant Dis. 2022, 106, 1358–1365. [Google Scholar] [CrossRef]
  102. Bui, H.X.; Hadi, B.A.R.; Oliva, R.; Schroeder, N.E. Beneficial bacterial volatile compounds for the control of root-knot nematode and bacterial leaf blight on rice. Crop Prot. 2020, 135, 104792. [Google Scholar] [CrossRef]
  103. Costa, L.S.A.S.; Campos, V.P.; Terra, W.C.; Pfenning, L.H. Microbiota from Meloidogyne exigua egg masses and evidence for the effect of volatiles on infective juvenile survival. Nematology 2015, 17, 715–724. [Google Scholar] [CrossRef]
  104. Wolfgang, A.; Taffner, J.; Guimaraes, R.A.; Coyne, D.; Berg, G. Novel strategies for soil-borne diseases: Exploiting the microbiome and volatile-based mechanisms toward controlling Meloidogyne-based disease complexes. Front. Microbiol. 2019, 10, 1296. [Google Scholar] [CrossRef]
  105. Gómez-Tenorio, M.A.; Zanón, M.J.; de Cara, M.; Lupión, B.; Tello, J.C. Efficacy of dimethyl disulfide (DMDS) against Meloidogyne sp. and three formae speciales of Fusarium oxysporum under controlled conditions. Crop Prot. 2015, 78, 263–269. [Google Scholar] [CrossRef]
  106. Warnock, N.D.; Wilson, L.; Patten, C.; Fleming, C.C.; Maule, A.G.; Dalzell, J.J. Nematode neuropeptides as transgenic nematicides. PLoS Pathog. 2017, 13, e1006237. [Google Scholar] [CrossRef][Green Version]
  107. Sharifi, R.; Ryu, C.M. Revisiting bacterial volatile-mediated plant growth promotion: Lessons from the past and objectives for the future. Ann. Bot. 2018, 122, 349–358. [Google Scholar] [CrossRef][Green Version]
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Diyapoglu, A.; Oner, M.; Meng, M. Application Potential of Bacterial Volatile Organic Compounds in the Control of Root-Knot Nematodes. Molecules 2022, 27, 4355.

AMA Style

Diyapoglu A, Oner M, Meng M. Application Potential of Bacterial Volatile Organic Compounds in the Control of Root-Knot Nematodes. Molecules. 2022; 27(14):4355.

Chicago/Turabian Style

Diyapoglu, Ali, Muhammet Oner, and Menghsiao Meng. 2022. "Application Potential of Bacterial Volatile Organic Compounds in the Control of Root-Knot Nematodes" Molecules 27, no. 14: 4355.

Article Metrics

Back to TopTop