It is estimated that 99% of the bacteria and archaea are able to produce at least one bacteriocin. Historically, lactic acid bacteria (LAB) were studied as main bacteriocin producers, mostly because of their long history of safe use in food fermentation (O’Sullivan et al., 2002). Nisin (Figure 3C), produced by Lactobacillus lactis subsp. lactis, was approved as a food additive in the 1960s and has since then been used in over 50 countries for its antimicrobial activity against Gram-positive pathogens such as Clostridium spp. and Bacillus spp. (Klaenhammer, 1988; Delves-Broughton, 1990). However, the search for new bioactive molecules has rapidly expanded to other bacteriocin-producing genera, with a particular attention, in the late 1990s, to the GRAS (generally recognized as safe) Bacillus species whose bacteriocin antimicrobial spectra were broader than those of LAB (Pedersen et al., 2002; Riley and Wertz, 2002; Sumi et al., 2015).

The generic biosynthetic pathway of Bacillus species bacteriocins includes several post-translational modifications, including the proteolytic cleavage of the leader peptide at the N-terminal end (McIntosh et al., 2009). The modifications of active peptides, its secretion and the immunity to the bacteriocin (as described below) vary depending on the bacteriocin class.

While many classifications have been suggested over the years, one reasonable way to cope with the diversity of the Bacillus bacteriocins is to sort them on the basis of their biosynthetic pathway as previously reported for Streptococcus spp. and Enterococcus spp. bacteriocins (Nes et al., 2007) and reviewed in Abriouel et al. (2011). Accordingly, three main classes subdivided into several subclasses can be distinguished for the B. subtilis group. As detailed in Table 1, Class I includes the post-translationally modified peptides such as the lantibiotics whereas the non-modified peptides are grouped in Class II; Class III involved bacteriocins larger than 10 kDa (Abriouel et al., 2011). Supplementary Table S1 summarizes the different RPs produced by the strains belonging to the B. subtilis group, as well as their reported antimicrobial activities.

Class I includes small AMPs (19–38 AA) with extensive post-translational modifications. Subclasses I.1, I.2, and I.3 have in common their lantibiotic structure, which refers to inter-residual thioester bonds made of modified AA residues. As illustrated in Figure 3, lantibiotics involve 2,3-didehydroalanine (Dha) and (Z)-2,3-didehydrobutyrine (Dhb), resulting from the dehydration of serine and threonine residues, respectively. The intra-molecular addition of Dha or Dhb on a cysteine residue leads to the respective formation of lanthionine and methyllanthionine bridges (Willey and Donk, 2007). Subtilin (Figure 3B), from subclass I.1, is one of the most studied bacteriocins from the B. subtilis group. Its structure shares several similarities with nisin A lantibiotics, shown in Figure 3C (Guder et al., 2000; Abriouel et al., 2011). Peptides from subclass I.4 undergo other types of modifications. For instance, subtilosin A is a head-to-tail cyclic peptide with unusual inter-residue linkages (i.e., Cys-Phe bond) (Marx et al., 2001; Kawulka et al., 2004).

Class II bacteriocins include small (<10 kDa), linear and non-modified peptides, resistant to heat and acido-basic treatments. They are divided in three subclasses based on a conserved AA motif near their N-terminus. The YGNGVXC (X is any AA) motif is associated to pediocin-like peptides from subclass II.1 whereas DWTXWSXL is specific to thuricin-like peptides from subclass II.2. Subclass II.3 comprises the small non-modified AMPs without any typical motif in their AA sequence (Abriouel et al., 2011). Finally, class III bacteriocins consist into large and heat labile molecules, generally characterized by a phospholipase activity (Cleveland et al., 2001).

Because of their wide diversity, bacteriocins display different modes of action such as protoplasm vesicularization, pore formation or cell disintegration (Sumi et al., 2015). They are generally bactericidal with some exceptions that exhibit bacteriostatic activities (Gautam and Sharma, 2009). For most class I and II bacteriocins, the target of their activity is the bacterial envelope due to their amphiphilic or hydrophobic properties. For instance, lantibiotics from subclass I.1 have a dual mode of action. On the one hand, they can inhibit the cell wall synthesis of the targeted bacteria through binding to lipid II, the major transporter of peptidoglycan subunits across the inner cell membrane. On the other hand, lipid II can be used as a docking molecule to insert the lantibiotic in the membrane leading to pore formation and ultimately to cell death as well described in Chatterjee et al. (2005) and Cotter et al. (2005). This duality has been reported for subtilin, a class I bacteriocin which is active against a broad range of Gram-positive bacteria such as Staphylococcus simulans, B. subtilis, and Bacillus stearothermophilus (Linnett and Strominger, 1973; Parisot et al., 2008).

Many regulation systems mediate bacteriocin production, secretion and immunity. Bacteriocin production is usually linked to particular cellular events such as stress responses. For instance, subtilin production depends on cell density and is increased under starvation conditions (Abriouel et al., 2011). Lantibiotic production is also mediated by QS. For subtilin, it has been demonstrated that the peptide itself acts as an auto-inducer of its own production (Kleerebezem, 2004). The export of bacteriocins is generally ensured by a dedicated membrane-associated ATP-Binding Cassette (ABC) transporter. For some lantibiotics, the cleavage of the leader peptide often occurs in a proteolytic domain present in the ABC transporter as described in McAuliffe et al. (2001) and Cotter et al. (2005). The immunity of the producing strains to its own active bacteriocin(s) can be achieved by several mechanisms like the secretion of immunity proteins sequestering the peptide, the bacteriocin re-export through an ABC transporter system or the alteration of the targeted peptidoglycans bonds (e.g., modification of the cell wall or cytoplasmic membrane charge) (Cotter et al., 2005; Dubois et al., 2009).

Among the B. subtilis group, two major types of enzymes exhibit antagonistic activities (Supplementary Table S1): the lytic enzymes and those involved in quorum quenching (QQ). Several strains from the B. subtilis group have indeed been identified as capable to produce lytic enzymes with biocontrol potential (Herrera-Estrella and Chet, 1999; Kumar et al., 2012; Shafi et al., 2017). They include cellulases, glucanases, proteases and chitinases and are generally referred to as cell wall degrading enzymes (CWDE) (Ariffin et al., 2006; Alamri, 2015; Caulier et al., 2018). They are particularly active against fungi since chitin and glucan are the major constituents of their cell wall where various glycoproteins are embedded (Bowman and Free, 2006; Geraldine et al., 2013; Gomaa, 2012).

Quorum quenching is able to silence or block QS which is generally defined as the cell-to-cell communication mechanism through the production of signal molecules (Czajkowski and Jafra, 2009). N-acyl-homoserine lactones (AHLs), composed of a fatty acid side chain and a homoserine lactone (Figure 4) are the most characterized signal autoinducers in Gram-negative bacteria. When a bacterial population proliferates, concentration of AHLs increases so that all the cells coordinate their metabolic activities (e.g., biofilm formation, sporulation, virulence factors or antibiotic production) (Dong et al., 2004). As the QS system brings ecological advantages to a coordinate population, QQ is able to counteract QS. Four types of enzymes (i.e., lactonase, decarboxylase, acylase, and deaminase) are able to inactivate AHLs, as illustrated in Figure 4 (Czajkowski and Jafra, 2009). B. subtilis AHL-lactonases have for instance attracted interest for biocontrol since they affect the growth of deleterious microbial pest such as Pectobacterium carotovorum subsp. carotovorum causing potato soft rot (González and Keshavan, 2006).

Lipopeptides are usually synthetized through a NRPS sequential addition of AA residues, either in an iterative or non-iterative way. Similarly to PKSs, NRPSs have a modular organization implementing the initiation, elongation, and termination modules (Figure 6B). Each module is subdivided in core domains whose catalytic and carrier domains slightly differ from PKSs, as shown in Figure 7B. The biosynthesis which was previously summarized in Ongena and Jacques (2008) and Raaijmakers et al. (2010) starts with an adenylation domain (A domain) that recruits and phosphorylates an AA monomer into an aminoacyl adenylate intermediate. The intermediate is then linked to the corresponding peptidyl carrier protein or thiolation domain (PCP or T domain) through a thioester bond. The PCP acts as a bridge and ensures the link with the condensation domain (C domain) that forms the C–N bond between the recruited aminoacyl and the peptide acyl chain in formation. The termination module contains a thioesterase domain (TE) that catalyzes the release of the final peptide acyl chain (Ongena and Jacques, 2008; Raaijmakers et al., 2010). The elongation modules can be supplemented with accessory domains such as cyclization domain (Cy), epimerization domain (E) and methylation domain (M). Those domains are able to modify the growing peptide chain which leads to diverse mature compounds structure (Cane and Walsh, 1999; Challis and Naismith, 2004).

Since the LP biosynthetic pathways are highly flexible, the range of produced LPs is extremely heterogeneous. Among LPs produced by Bacillus spp., four main families have been distinguished: kurstakins, surfactins, iturins, and fengycins (Jacques, 2011). Each family shares the same structural features based on the nature and organization of the peptide moiety or fatty acid tail, as summarized in Table 4. Strains from the B. subtilis group produce surfactins, iturins and fengycins whereas kurstakins are produced by B. thuringiensis strains (Béchet et al., 2012). Among the three LP families produced by B. subtilis, at least eight fengycins, 13 surfactins and 14 iturins variants have been described so far, as detailed in Supplementary Table S2.

For each LP family, the compounds production is mainly regulated by environmental factors such as carbon sources, oxygen availability, pH and temperatures (Yakimov et al., 1995; Kim et al., 1997; Cosby et al., 1998). Warm temperature (≥37°C) and anaerobic conditions increase the production of surfactins while lower temperatures (25–37°C) and aerated bioreactors favor fengycins and iturins family metabolites (Jacques, 2011). The production of surfactins by B. subtilis is also QS-dependent and involves ComX and PhrC. These pheromones trigger complex cascades regulating cell density-dependent processes such as sporulation and competence (Hamoen et al., 2003; Ongena et al., 2005).

Iturins and fengycins are mainly known for their strong antifungal activity against several plant and human pathogenic fungi (Supplementary Table S2). In addition, iturin-like mycosubtilin, bacillomycin R, subtulene A and eumycin show antibacterial properties (Besson et al., 1976; Leclere et al., 2005; Thasana et al., 2010). Contrary to iturins and fengycins, surfactins mainly display antiviral and antibacterial activities (Ongena and Jacques, 2008). Their antiviral activity essentially targets enveloped viruses (e.g., herpes simplex or porcine epidemic diarrhea viruses). They also inhibit pathogenic bacteria such as Legionella pneumophila, Listeria monocytogenes, R. solanacearum or X. oryzae (Naruse et al., 1990; Yakimov et al., 1995; Sabaté and Audisio, 2013; Loiseau et al., 2015; Luo et al., 2015). However, some surfactins are able to control important fungal plant and human pathogens such as Botrytis cinerea, Candida albicans, F. oxysporum or Rhizoctonia solani (Jenny et al., 1991; Lee et al., 2007; Qi et al., 2010; Dimkić et al., 2013; Romano et al., 2013).

The mere composition of LPs, where a peptide moiety is bound to a lipid tail, gives them an amphiphilic property. This nature makes them excellent surfactants and plays a significant role in their biological functions and antimicrobial properties. Indeed, LPs are able to destabilize the plasma membrane via a pore forming activity leading to the cell death of the target microbes. Their antiviral activity is the result of a similar disintegration of the bi-lipid envelope of virions explaining the weak LPs activity against plant viruses among which very few are enveloped (Ongena and Jacques, 2008).

Bacillus spp. LPs have many other biological and ecological functions as fully documented by Raaijmakers et al. (2010). They are also known to impact other metabolic mechanisms such as biofilm formation, motility, virulence, plant root colonization, and plant defenses. Moreover, it has been suggested that their participation to the degradation of hydrophobic substrates could be used for polluted soils bioremediation (Mulligan et al., 2001). Although some lipopetides have already been exploited as food biopreservatives or crop protection products, the industrial interest for LPs in specific applications is unsurprisingly continuously growing.

Itoic acid is a mono-peptide composed of a 2,3-dihydroxybenzoate (DHB) molecule bound to a glycine. It is used as a precursor by trimodular NRPS machinery to produce bacillibactin which is obtained after a condensation of three units of DHB-glycine-threonine (May et al., 2001). The synthesis of the final hexapeptide is catalyzed by a terminal thioesterase domain leading to the production of a methylated trilactone ring link to three catecholates moieties. It is this cyclic structure that enables the sequestration of the metal atom (Dertz et al., 2006). Itoic acid and bacillibactin are both catecholic siderophores that chelates iron reducing its bioavailability. This is limited access to iron that allows B. subtilis to antagonize the growth of other surrounding microbes such as, for instance, F. oxysporum f. sp. capsici (Yu et al., 2011).

Bacteria from the B. subtilis group are also able to synthesize other antimicrobial NRPs through non-thiotemplate mechanism. Rhizocticins are di- and tri-phosphono-peptides. They are constituted of a L-2-amino-5-phosphono-3-cis-pentenoic acid (APPA) linked to an arginine (rhizocticin A). They can be supplemented with an additional valine (rhizocticin B), isoleucine (rhizocticine C) or leucine (rhizocticine D). After their integration into the target microbes, their cleavage by host cell peptidases releases the fungitoxic L-APPA moiety that interferes with threonine metabolism in fungal cells. Interestingly, rhizocticin A has also an antagonistic activity against nematodes such as Caenorhabditis elegans (Kugler et al., 1990).

In addition to rhizocticin compounds, two other dipeptide NRPs are produced by B. subtilis: bacilysin (also known as tetaine) and its chlorinated derivative, chlorotetain. They contain L-alanine (or chlorine-L-alanine) bound to the non-proteinogenic L-anticapsin (Kenig and Abraham, 1976; Rapp et al., 1988). Despite their simple composition, these bioactive compounds display strong antibacterial activity mediated by the anticapsin moiety that inhibits the glucosamine-6-phosphate synthase. Its inhibition suppresses the biosynthesis of peptidoglycans that are the main constituents of bacterial cell wall (Steinborn et al., 2005; Mahlstedt and Walsh, 2010). For the fungi, it has been proposed that because anticapsin is able to inhibit the production of chitin and fungal membrane mannoproteins, bacilysin and chlorotetain exhibit antifungal activity against Aspergillus fumigatus or C. albicans (Milewski et al., 1986; Rapp et al., 1988).

Finally, bacitracin and mycobacillin are two cyclic polypeptides produced by B. subtilis. Bacitracins are dodecapeptides containing a cyclic heptapeptide linked to a thiazoline ring (Johnson et al., 1945). They are mostly active against Gram-positive bacteria where they inhibit the bacterial cell-wall biosynthesis by preventing the lipid carrier from re-entering in the reaction cycle of peptidoglycan synthesis (Siewert and Strominger, 1967). Besides this primary mode of action, bacitracin might also act through other mechanisms affecting membrane functions, hydrolytic enzymes and/or the biosynthesis of ubiquinone precursors (Konz et al., 1997). Mycobacillin is an antifungal cyclic tridecapeptide altering the membrane of fungi like Aspergillus niger (Majumdar and Bose, 1958). Interestingly, its biosynthesis is rather peculiar. Although it is catalyzed by a large NRPS complex, it is divided in three fractions (A, B, and C) and does not use a thio-template mechanism (Zuber et al., 1993). Each fraction of the enzymatic complex contains a single enzyme polypeptide that catalyzes the polymerization of a first pentapeptide (A), a second nonapeptide (B) and the final tridecapeptide.

Volatile inorganic compounds synthesized by microorganisms are mainly by-products of primary metabolism. They are carbonated, hydrogenated, sulfur or nitrogen-containing compounds such as CO₂, CO, H₂, HCN, H₂S, N₂, NH₃ and NO. Nitrogen-containing compounds are mostly released in aerated upper sediments layers by denitrifying bacteria. In this process, nitric oxide is enzymatically produced by the nitric-oxide reductase or the nitric-oxide synthase (Adak et al., 2002). The range of antimicrobial activities exhibited by VIC nitrogen-containing compounds from the B. subtilis group is wide. For instance, NO is able to induce systemic acquired resistance (SAR) in plants against bacterial pathogens such as R. solanacearum (Wang et al., 2005). A contrario, ammonia, a secondary metabolite from the catabolism of the amino acids L-aspartate, is known to be active against soil-borne Oomycetes such as Pythium spp. (Howell et al., 1988). Hydrogen cyanide, derived from the glycine catabolism, shows a direct antagonistic activity against aerobic microorganisms by inhibiting metal-containing enzymes such as the cytochrome c oxidase active in the respiration chain (Cherif-Silini et al., 2016).

Deeper in the soil, under low oxygen concentration, bacteria tend to produce different VICs such as H₂ or H₂S. Those compounds can serve as electron acceptors, AA precursors or antimicrobial metabolites. Hydrogen sulfide could be produced by B. subtilis from sulfate reduction or as a by-product of L-methionine and L-cysteine catabolism via a direct cleavage of L-methionine or a transamination followed by reductive demethiolations (Even et al., 2006; Schulz and Dickschat, 2007). It is known to exhibit antifungal activity against several plant pathogens such as A. niger or Penicillium italicum but also against some food-borne bacteria or human pathogens (Fu et al., 2014). Curiously, it is also known to act as a bacterial defense mechanism against antibiotics (Shatalin et al., 2011). Interestingly, ammonia increases the resistance of several Gram-negative and Gram-positive bacteria to antibiotics too (Bernier et al., 2011).

Volatile organic compounds are small compounds with fewer than 20 carbon atoms and are characterized by low molecular mass (100–500 Da), high vapor pressure, low boiling point and a lipophilic moiety. These features ensure an easy evaporation and a long distance distribution which is convenient in a complex matrix like soil (Schmidt et al., 2015). Their diffusion and production by soil-borne microbes are strongly dependent on various factors such as nutrient and oxygen availability, temperature, pH, physiological state of microorganisms, soil moisture, texture and architecture (McNeal and Herbert, 2009; Insam and Seewald, 2010; Effmert et al., 2012). The majority of VOCs derives from glucose oxidation involving glycolysis and the subsequent cycles such as the tricarboxylic acid cycle (TCA) as it has been well summarized in Korpi et al. (2009) and Schmidt et al. (2015). However, their production can also result from various other pathways such as aerobic heterotrophic carbon metabolism, fermentations, AA degradation, terpenes synthesis or sulfur reduction (Peñuelas et al., 2014). Based on previous reviews presented in Schulz and Dickschat (2007); Peñuelas et al. (2014) and Audrain et al. (2015), five categories of VOCs can be distinguished: (1) fatty acids and derivatives, (2) terpenoids, (3) nitrogen-containing VOCs, (4) sulfur-containing VOCs, and (5) metalloid- or halogenated-containing VOCs. To date, about 2,000 compounds produced by almost 1,000 species of microorganisms have been listed in the mVOC 2.0 database (Lemfack et al., 2018). According to this database, almost 70% of recorded Bacillus VOCs are fatty acids derivatives (alcohols, ketones, alkanes, aldehydes, alkenes, and acids) followed by sulfur- and nitrogen-containing compounds. Supplementary Table S3 displays the VOCs produced within the B. subtilis group and their antimicrobial activity.

Since many volatile fatty acids and their derivatives result from the glucose metabolism, their precursors mostly derive from the Embden-Meyerhof (glycolysis), Entner-Doudoroff, heterolactic and homolactic fermentation pathways (Peñuelas et al., 2014). B. subtilis bacteria, for instance, ferment pyruvate to produce ketone compounds such as acetoin (3-hydroxy-2-butanone) or 2,3-butanedione under anaerobic conditions (Ryu et al., 2003). Other intermediates coming from fatty acid biosyntheses or their β-oxydations are also used as precursors by microbes and transformed into VOCs through a decarboxylation reaction or a reduction of their carboxyl group (Schulz and Dickschat, 2007). They provide essential hydrocarbons but also other fatty acid derivatives. An oxidative deamination of several amino acids can lead to the production of aldehyde, ketone or alcohol volatile too. For instance, the degradation of L-phenylalanine or L-tyrosine can be the first step of the aromatic volatile compounds synthesis such as benzene or its carbohydrate derivatives. Finally, benzenoid volatiles can also be synthesized by microbes through the shikimate pathway that leads to the formation of chorismate, a natural precursor of aromatic amino acids (Bentley and Haslam, 1990). Degradation of intermediates from the shikimate pathway or aromatic amino acids can also lead to the production of benzenoid volatiles (Dickschat et al., 2005).

This wide variety of volatile fatty acids and their derivatives make them the most important group of VOCs produce by microbes and represent up to 87% of known antimicrobial VOCs produced by B. subtilis bacteria (Supplementary Table S3). They can be divided in two main categories: hydrocarbons (alkanes, alkenes, alkynes) or carbohydrates (acids, alcohols, aldehydes, esters, furans, ketones, lactones, benzenoids). Among them, benzenoids is the most represented sub-category followed by alkanes, aldehydes, ketones, acids, and alcohols. Even though benzenoids could be considered as an individual category, they can also be seen as fatty acids derivatives because a large majority of antimicrobial benzenoid volatile produced by B. subtilis harbor a benzene core linked to a fatty acid derivatives.

There is an important diversity of benzenoids, sometimes linked with carbohydrate chains containing nitrogen, sulfur or both. Most of these antimicrobial volatile exert fungicidal activities but some have been characterized for their antibacterial or nematicidal abilities, too. Their mode of action is rarely fully characterized. For instance, morphological abnormalities on fungal and bacterial cells have been documented after an exposition to B. subtilis VOCs (Tahir et al., 2017). Volatile such as 1,3-butadiene or 2,3-butanediol are also known to induce modifications in the expression of genes linked to the pathogenicity of R. solanacearum and Pectobacterium carotovorum (Marquez-Villavicencio et al., 2011; Tahir et al., 2017). In addition to direct antimicrobial activities, fatty acids volatile have also several other biological functions. For instance, acetoin and 2-butanone have the ability to stimulate plant defenses or to induce plant stress tolerance which then promote plant growth (Ryu et al., 2003; Ryu et al., 2004; Ryu, 2015). They are essentially produced by strains of B. amyloliquefaciens, B. velezensis or B. subtilis (Audrain et al., 2015).

Terpenes and their derivatives (also known as terpenoids or isoprenoids) are among the most abundant secondary metabolites found in living systems (Fisher et al., 2001; Gershenzon and Dudareva, 2007). They originate from two main precursors: isopentenyl pyrophosphate (IPP) and its allylic isomer the dimethylallyl pyrophosphate (DMAPP) (Schulz and Dickschat, 2007). IPP and DMAPP are also the end-products of the deoxy-xylulose phosphate pathway (DOXP) starting with pyruvate and glyceraldehyde-3-phosphate originating from the glucose metabolism (Fisher et al., 2001). Terpenoids can be synthesized from isoprene molecules too. Julsing et al. (2007) showed that, in B. subtilis, isoprene is not formed by the MVA or DOXP pathways but, as in plant systems, might be a product of the methylerythritol phosphate (MEP) pathway (Guan et al., 2015).

Isoprenoid compounds are produced by all living organisms for essential physiological functions such as electron transport, membrane fluidity, light harvesting, photoprotection, anchoring of molecules to specific membranes and signaling (Fisher et al., 2001). The signaling ability is particularly important and is associated with several antagonistic, mutualistic or multi-trophic interactions (Shrivastava et al., 2015). More than 25,000 terpenic compounds have been listed and, for the vast majority, their biological functions and roles remain unknown (Buckingham, 1997). Volatile terpenes are generally recognized for their ability to inhibit bacteria (Scortichini and Rossi, 1991), fungi (Hammer et al., 2003; Dambolena et al., 2008), nematodes (Gu et al., 2007) or insects (Lee et al., 2003; Justicia et al., 2005). They can be classified in three categories: isoprene, monoterpenes (C₁₀) and sesquiterpenes (C₁₅) (Schmidt et al., 2015).

The mode of action of these compounds might be linked to their lipophilic nature allowing them to destabilize the cell membrane integrity (Cox et al., 2000; Inoue et al., 2004). To our knowledge, only two terpenes produced by B. subtilis show antimicrobial abilities: isoprene and monoterpene α-terpineol exhibit antagonistic activities against cyanobacteria and nematodes (Wright and Thompson, 1985; Gu et al., 2007).

Little is known about the biosynthetic pathways of nitrogen-containing VOCs. Nevertheless, it is accepted that two main routes can be used: a non-enzymatic amination of acyloins, that can lead to the formation of pyrazines (Schulz and Dickschat, 2007) or derived from α-aminoketone intermediates resulting from AA catabolism (Owens et al., 1997; Zhu et al., 2010).

Nitrogen-containing VOCs can be distinguished based on their cyclization rate. Within non-cyclic compounds, three groups are identified (amides, amines and imines) while there are five categories of cyclic compounds (azoles, pyrazines, pyridines, pyridazines, and pyrimidines). Pyrazines are strongly represented among microbial volatile and are separated in two classes: lower-alkylated and higher-alkylated pyrazines (Schulz and Dickschat, 2007). These compounds are characterized by a strong odor and several B. subtilis coming from the rhizosphere or from food fermentations have already been recognized as pyrazines producers (Sugawara et al., 1985; Kosuge and Kamiya, 1962; Larroche et al., 1999; Leejeerajumnean et al., 2001). Pyrazines from B. subtilis strains are known to exhibit antifungal and nematicidal activities (Gu et al., 2007; Chen et al., 2008; Chaves-López et al., 2015; Haidar et al., 2016). For instance, tetramethylpyrazine inhibits the growth of Moniliophthora perniciosa and F. oxysporum f. sp. lactucae. Additionally, it acts on sporulation and elongation of the germ-tube of B. cinerea (Chen et al., 2008; Chaves-López et al., 2015). It is interesting to note that B. subtilis pyrazines can also exhibit antibacterial activities such as pulcherriminic acid which inhibits the growth of S. aureus, E. coli and Proteus vulgaris (Coutts et al., 1965). Beside pyrazines, strains from the B. subtilis group are able to produce other nitrogen VOCs such as 1H-imidazole,1-ethyl showing antifungal activities against numerous soil-borne phytopathogens (Lupetti et al., 2002; Liu et al., 2008; Snelders et al., 2009; Schmidt et al., 2015).

Microbial VOCs containing sulfur (VSCs) derive from two main pathways originated from inorganic or organic sources (Schulz and Dickschat, 2007): inorganic sulfate reduction in methylated inorganic sulfides compounds or, for some microbial VSCs, originate from catabolism of AA such as L-methionine or more rarely, L-cysteine (Schulz and Dickschat, 2007). Some VSCs are produced as secondary volatiles via the production of hydrogen sulfide or methanethiol. Indeed, these two compounds are important precursors for subsequent VSCs synthesis (Schulz and Dickschat, 2007; Sourabié et al., 2012). Within the B. subtilis group, multiple VSCs such as dimethyl disulfide (DMDS), dimethyl trisulfide (DMTS), S-methyl thioacetate or S-methyl butanethioate have been characterized for their antifungal and nematicidal activities (Coosemans, 2005; Gerik, 2005; Gu et al., 2007; Kai et al., 2009; Wang et al., 2009; de Vrieze et al., 2015; Schmidt et al., 2015; Velivelli et al., 2015; Gotor-Vila et al., 2017). A putative antibacterial effect of DMDS is not to exclude. Indeed, DMDS is known to affect the bacterial cell-to-cell communications through a decrease in the amount of N-acyl homoserine lactone (AHL) mediating QS (Chernin et al., 2011).

Other volatile organic compounds such as halogenated, metalloids, tellurium or selenium compounds have also been described. However, at the time of writing, no B. subtilis strains have been proved to produce these type of VOCs (Schulz and Dickschat, 2007), although related bacteria, like Bacillus arsenicoselenatis, have been shown to generate them (Switzer Blum et al., 1998).