LPs are surface-active molecules also called biosurfactants. Biosurfactants help to condition the producing strain’s environment by supporting various processes including bacterial motility, attachment and colonization of surfaces, biofilm development and access to nutrients and water (Nybroe and Sorensen, 2004; D'Aes et al., 2010; Raaijmakers et al., 2010; Geudens and Martins, 2018; Bricout et al., 2022).

Various lab-based studies show a strong correlation between LP production and swarming motility when comparing wild-type and LP-deficient mutant strains (de Bruijn et al., 2008; de Bruijn and Raaijmakers, 2009; Geudens and Martins, 2018). The ability of LPs to alter surface tension and viscosity is determined by their structural properties (D'Aes et al., 2010). In the amphisin producer Pseudomonas sp. DSS73, amsY and gacS mutant strains are unable to swarm on soft agar and as expected swarming motility is restored by the addition of amphisin but also viscosin, tensin and serrawettin from Serratia liquefaciens (Andersen, 2003). In contrast, synthetic surfactants were unable to complement the non-motile phenotype indicating that unique physiochemical properties relating to the chemical structure of LPs contribute to bacterial movement (Andersen, 2003). The diverse structures and physiochemical properties of LPs make them attractive alternatives to chemical surfactants for numerous industrial applications (Ceresa et al., 2023).

One of the most effective biosurfactants reported to date is surfactin from Bacillus subtilis capable of lowering the surface tension of water from 71 to 27 mN/n at a critical micelle concentration (CMC) of 20 µM (Yeh et al., 2005). Viscosin also shows strong surface activity reducing water surface tension to 25 mN/n at CMC of 4 μg/mL (Nybroe and Sorensen, 2004). Similarly, viscosinamide reduces water surface tension to 27 mN/n however no CMC values are available (Nybroe and Sorensen, 2004). Syringomycin and syringopeptin reduce water surface tension in ranges of 31–35 mN/n comparable to values of 31.5 mN/n and 38 mN/n recorded for putisolvin II and tolaasin, respectively (Hutchison and Johnstone, 1993; Nybroe and Sorensen, 2004; Janek et al., 2010). For most Pseudomonas LPs however information on physiochemical properties, e.g., surface reduction activities and CMC values, foaming capacity, emulsifying activity and compound stability are lacking.

In situ studies support the role of LPs in motility and colonization of specific habitats, e.g., plant material or fungal tissues. Massetolide A contributes to colonization of tomato roots by P. lactis (fluorescens) SS101, viscosin is required for colonizing broccoli florets by a pectolytic strain of P. fluorescens, cichofactin is essential for Pseudomonas cichorii to colonize lettuce leaves, while Pseudomonas sp. DSS73 uses amphisin to colonize sugar beet seeds and roots (Hildebrand et al., 1998; Nielsen and Sorensen, 2003; Tran et al., 2007; Pauwelyn et al., 2013). Other LPs, for example, poaeamide from Pseudomonas poae RE*1-1-13 and putisolvin from P. putida 257 do not positively contribute to rhizosphere competence as no differences in plant root colonization are observed between WT and biosurfactant deficient mutant strains (Kruijt et al., 2009; Zachow et al., 2015).

For many plant-associated LP-producers, biosurfactant-assisted motility is a key determinant for successful rhizosphere and phyllosphere colonization, where in addition to plant material, LPs also mediate interactions with fungi. In the phytopathogen Ralstonia solanacearum, the LP ralsolamycin (RM) facilitates fungal tissue invasion by inducing the formation of fungal survival structures known as chlamydospores in Aspergillus flavus (Spraker et al., 2016). An RM deficient rmyA mutant cannot induce chlamydospore formation and shows reduced hyphal invasion (Spraker et al., 2016). Subsequently (Venkatesh et al., 2022), demonstrated that compared to the rmyA mutant, WT R. solanacearum is internalized in chlamydospores during co-culture with A. flavus and shows increased fitness under starvation and cold stress. Phylogenetically distinct bacteria (not associated with endofungal lifestyles) including the nitrogen-fixing bacterium Herbaspirillium seropedicae were also shown to colonize chlamydospores of A. flavus when treated with WT supernatants further confirming the role of RM in facilitating fungal invasion (Venkatesh et al., 2022). Such knowledge could be translated to stimulate diverse endofungal interactions for improved ecosystem services, e.g., nitrogen fixation (Venkatesh et al., 2022). While a number of other LPs including the Pseudomonas CLPs viscosinamide and tensin can induce fungal survival structures, their role in fungal invasion and endosymbiosis is currently unknown (Nielsen et al., 1999; Nielsen et al., 2000; Venkatesh et al., 2022). More information on interactions with fungi is described in Section 3.2.2.

Biosurfactants also regulate biofilm development with a number of CLPs shown to promote biofilm formation (massetolide A, sessilin and xantholysin) and others involved in biofilm dispersal (arthrofactin, orfamide and putisolvin) (Kuiper et al., 2004; Bonnichsen et al., 2015). In P. sessilinigenes CMR12a, orfamides are indispensable for swarming motility, while sessilin is important for biofilm formation (D’aes et al., 2014). Based on contrasting studies, viscosin is proposed to mediate both biofilm formation and dispersal (de Bruijn et al., 2007; Bonnichsen et al., 2015). Viscosin-mediated biofilm dispersal is dependent on carbon starvation and by microscopic analysis it was observed that cells exhibiting high viscA (required for viscosin biosynthesis) expression levels were leaving biofilms, further supporting the role of LPs in motility. However, information on the mechanisms of LP-mediated biofilm development and dispersal, particularly in situ is limited and could benefit from more temporal studies examining LP function during biofilm lifecycles. For the producing bacteria, the ability to disperse is an important escape function under unfavorable nutrient conditions to support their spread throughout the environment and enable the colonization of new niches. From an industrial perspective, LPs displaying roles in biofilm formation could be used for bulk chemical production using biofilm fermentations whereas dispersal functions could be exploited as surface-coating agents or used in disinfectant formulations (Bonnichsen et al., 2015; Leonov et al., 2021).

Additional biological properties of LPs include the chelation of metal ions and xenobiotic degradation, e.g., petroleum hydrocarbons and pesticides (Nybroe and Sorensen, 2004; Raaijmakers et al., 2010; de Cássia et al., 2014; Raj et al., 2021). Research on biosurfactants as bioremediation agents to clean up contaminated soils is largely limited to rhamnolipids or Pseudomonas-leachates containing uncharacterized LPs (Sekhon Randhawa and Rahman, 2014; Sun et al., 2021). The CLPs viscosin, amphisin, massetolide A and putisolvin can emulsify alkane hydrocarbons such as n-hexadecane (Bak et al., 2015). Viscosin was also shown to stimulate alkane mineralization by a diesel-degrading bacterial consortium however the activity of the CLP was short-lived due to rapid (likely microbial) degradation (Bak et al., 2015). While LPs are proposed to chelate heavy metals and degrade insoluble hydrocarbons to increase their bioavailability and/or detoxify polluted soils for protection against toxicants, roles of LPs in such processes remain unclarified (Raaijmakers et al., 2010; Gutiérrez-Chávez et al., 2021).

Pseudomonas-CLPs display broad-spectrum antimicrobial properties exerting effects against bacteria, fungi, oomycetes and viruses as previously reviewed by (Nielsen et al., 2002; Geudens and Martins, 2018; Girard et al., 2020; Oni et al., 2022).

LPs also possess anti-proliferative activities against different cancer cell lines including viscosin (breast and prostate cancer cell lines) (Siani et al., 2008), xantholysin A (Pascual et al., 2014) and MDN-0066 (Cautain et al., 2015) (kidney tumor cell lines), pseudofactin II (melanoma cell lines) (Janek et al., 2013) and nunamycin/nunapeptin (mantle cell lymphoma, melanoma cell lines, T-cells leukemia) (Michelsen et al., 2015a). Accurate comparison of LP cytotoxic activities is challenged by the lack of standardization across assays. Similar to antimicrobial testing crude extracts over purified compounds are often used making the interpretation of results difficult as the bioactivities observed may derive from other compounds or synergistic activities between compounds. This has been observed in P. nunensis In5, where increased cytotoxic activity was seen when purified nunamycin and nunapeptin are mixed instead of applied individually (Michelsen et al., 2015a).

Pseudomonas LP producers are implicated in both positive and negative interactions with plants. Phytopathogenic strains typically co-produce phytotoxic CLPs from the Mycin and Peptin family, which act as virulence factors and form pores in plant membranes causing electrolyte leakage and necrosis. They usually also co-produce a third LLP or CLP not directly involved in virulence but instead aiding in plant tissue colonization. Phytopathogenic Pseudomonas LP producing strains taxonomically belong to the P. syringae group, or to the P. asplenii and P. corrugata subgroup of the P. fluorescens group (Girard et al., 2020). Phytotoxic CLPs from the Mycin and Peptin family also have strong antimicrobial activity (see above), demonstrating a dual role in pathogenicity and antagonism against competitors. Strains belonging to the P. corrugata subgroup can behave as plant pathogens, causing pit necrosis on tomato and pepper, but also show strong biological control activity against plant pathogens. They are often isolated from the roots and rhizosphere of non-diseased plants and from bulk soil (Catara, 2007; Gislason and de Kievit, 2020). Cormycin and corpeptin produced by P. corrugata double up as phytotoxic compounds and antimicrobial molecules against bacterial and fungal pathogens. Pseudomonas sp. SH-C52 (Mendes et al., 2011) (see above) also belongs to the P. corrugata subgroup and produces Mycin and Peptin-type CLPs with antifungal and anti-oomycete activity, indicating that there is no clear-cut line between plant pathogens and beneficials in these groups (Girard et al., 2020).

Disease suppressive soils are a rich source of LP-producing Pseudomonas strains. The potent biocontrol agents P. nunensis In5 (Michelsen and Stougaard, 2011) and Pseudomonas sp. SH-52 have been obtained from R. solani suppressive soils. Irrigation is known to protect potato tubers against the scab pathogen S. scabies. Microbiome analysis revealed that irrigated potato field had a larger proportion of Pseudomonadales bacteria than a non-irrigated potato field and that the presence of biosynthetic gene clusters encoding CLPs was positively correlated with disease suppression. Tensin, an 11:9 amphisin family CLP proved to be key determinant of in planta inhibition of potato scab in glasshouse trials (Pacheco-Moreno et al., 2021). Likewise, Pseudomonas strains able to produce CLPs belonging to 11 different families are dominant in the rhizosphere of cocoyam plants grown in a tropical soil suppressive to the cocoyam root rot disease caused by P. myriotylum (Oni et al., 2019a; Oni et al., 2020b).

Pseudomonas LP-producers demonstrating fungal and/or oomycete antagonism have gained considerable interest as candidates for controlling plant diseases. A detailed overview of Pseudomonas LP-mediated biocontrol is given by (D'Aes et al., 2010; Raaijmakers et al., 2010; Höfte, 2021; Oni et al., 2022). However, it is important to note that many studies are centered on the collection of data from lab-based experiments with only a handful of reports linking lab data to microcosm- or field-based studies. For example, for the viscosinamide producer P. fluorescens DR54 (Nielsen and Sorensen, 2003) a strong correlation between the ability of DR54 to inhibit growth of R. solani and P. ultimum in co-culture and during colonization of the rhizosphere of germinating sugar beet using plant-soil microcosms was observed. The study highlights the multifunctionality of CLPs with antibiotic and surfactant properties enabling the producing strain to condition its environment for successful rhizosphere colonization.

As wetting agents LPs can increase the solubility of nutrients and hydrophobic substrates for the producing strain (Nybroe and Sorensen, 2004; D'Aes et al., 2010; Mavrodi et al., 2010). For example, Bunster et al. (1989) reported that compared to a non-surface active Pseudomonas strain, surface-active isolates of P. fluorescens and P. putida increased the wetness of wheat leaves. Likewise, syringafactins are strong biosurfactants exerting hygroscopic activities to attract water vapor from the atmosphere increasing water availability and reducing water stress in P. syringae pv. syringae B728a on dry leaves and the apoplast of bean (Burch et al., 2014). Increasing the availability of nutrients and water potentially offers LP-producers a competitive advantage against other microbes including phytopathogens and thus may indirectly contribute to plant disease management by reducing pathogen growth delaying disease onset in the host plant (Nybroe and Sorensen, 2004).

A number of CLPs including massetolide A (Tran et al., 2007), sessilin, orfamide (Ma et al., 2016b; Ma et al., 2017), WLIP, lokisin and entolysin (Omoboye et al., 2019b) are involved in the induction of systemic resistance in plants. This type of resistance is systemically expressed rendering plants less susceptible to subsequent infection with pathogens (Pršić and Ongena, 2020). These studies are typically conducted with CLP mutants and CLP crude extracts with only a few studies demonstrating induction of ISR by pure compounds. Application of purified massetolide A at a concentration of 44 µM to tomato leaves or roots reduced the lesion area caused by Phytopthora infestans, but did not reduce disease incidence (Tran et al., 2007). Pure orfamide triggered ISR against R. solani web blight in bean at concentrations ranging from 0.001 to 0.1 µM (Ma et al., 2016b), while 25 µM orfamide was needed to elicit resistance to Cochliobolus miyabeanus in rice (Ma et al., 2017) suggesting different mechanisms involved.

While protists stimulate beneficial plant-microbe interactions and contribute important functions, e.g., nutrient cycling and pathogen removal they are major bacterial predators (Gao et al., 2019; Bahroun et al., 2021; Guo et al., 2021; Hawxhurst et al., 2023). One predation defense mechanism used by bacteria involves LPs. Using a combination of wild-type and mutant P. fluorescens strains, Mazzola et al. (2009) showed that massetolide and viscosin protect bacteria against predation by the amoeba Naegleria americana, with the predator showing a greater sensitivity to viscosin. LP-producers showed better persistence and protection in soil against the predator but the effect was only temporal (Mazzola et al., 2009). P. nunensis 4Aze was co-isolated with the social amoeba Polyspondyllium pallidum RM1 from forest soil. This strain produces keanumycin D, and nunapeptin B and C with suppressive activities against amoebal predators and the bacterivorous nematode Oscheius myriophilus, highlighting the broad-spectrum activity of LPs and underexplored anti-predator function of these compounds (Pflanze et al., 2023) Also the keanumycins produced by Pseudomonas sp. QS1027 have strong amoebicidal activity (Götze et al., 2023). More information on the influence of predator interactions and predator-derived molecules on LP regulation is needed.

Antiparasitic activities are documented for viscosin against the human parasitic protozoan Trypanosoma cruzi, the causal agent of Chagas disease. A viscosin-like LP from Pseudomonas sp. H6 was active against the fish parasitic ciliate Ichthyophthirius multifiliis and showed inhibitory effects against green algae, crustaceans, cyanobacteria and zebrafish embryos (Raaijmakers et al., 2006; Korbut et al., 2022).

LPs also mediate insect interactions with orfamides, sessilin and viscosin shown to possess insecticidal properties (Geudens and Martins, 2018). Insecticidal activity appears a multifactorial process involving LPs and other metabolites, e.g., Fit toxin, rhizoxin and HCN wherein the role of LPs appears to be strain-specific (Jang et al., 2013; Flury et al., 2017).

The main mode of action of LPs is membrane disruption through pore-formation causing membrane leakage and cell death (Geudens and Martins, 2018). The majority of mechanistic studies rely on model cell membranes that enable simpler and well-controlled experiments however they lack the complexity of real biological membranes. Consequently, the detailed mechanism of action underlying LP pore-formation including membrane selectivity, is largely unknown. As biological organisms are capable of altering their lipid membrane composition in response to external signals in order to adapt to their physical environment (Chwastek et al., 2020), it is likely that LPs have not one but multiple modes of action and display context-dependent activities. This may also contribute to the low resistance towards LPs in the environment despite their ubiquitous nature (Peschel and Sahl, 2006; Steigenberger et al., 2023). Future work should determine the influence of basic membrane parameters, e.g., membrane thickness in addition to physical membrane properties and lipid composition of different membrane types, e.g., bacterial, fungal and mammalian on LP activity. This was recently highlighted by (Ferrarini et al., 2022a) wherein the bioactivities of tolaasin and sessilin were reduced against oomycetes when membrane sterol composition is altered.

The best studied global regulatory system associated with LP production is the Gac/Rsm signal transduction pathway (Haas and Défago, 2005) (Figure 5). The cascade is initiated by the GacS/GacA two-component system composed of the membrane-bound GacS sensor kinase and the cognate GacA response regulator located in the cytoplasm. The GacS sensor kinase, initially called LemA, was first described in the bean pathogen P. syringae pv. syringae strain B728a and found essential to produce the CLP syringomycin (Hrabak and Willis, 1992; Hrabak and Willis, 1993). At high cell densities autophosphorylation of the GacS sensor kinase is triggered by an unknown chemical signal in the periplasm and the phosphate group is then transferred to the GacA response regulator via a phospho-relay mechanism. Phosphorylated GacA triggers the expression of small RNA genes. The resulting small RNAs (rsmX, rsmY, rsmZ) specifically bind to post-transcriptional repressor proteins (RsmA, RsmE) thereby relieving the translation repression exerted by these proteins at the ribosomal binding site of mRNAs encoding genes (Sonnleitner and Haas, 2011) involved in the biosynthesis or regulation of bioactive molecules including LPs. In most LP-producing strains mutations in gacS or gacA lead to a complete loss of LP production (Koch et al., 2002; de Bruijn et al., 2007; de Bruijn et al., 2008; Song et al., 2014; Olorunleke et al., 2017). Likewise, a rsmXYZ mutant in P. protegens CHA0 is no longer able to produce the CLP orfamide (Sobrero et al., 2017). Mutations in both rsmY and rsmZ resulted in loss of massetolide production in P. lactis SS101, while a double mutation in rsmE and rsmA restored massetolide production in a gacS mutant. In this strain, the most likely target of the RsmE and RsmA repressor proteins is the LuxR-type regulator MassAR (Song et al., 2015b).

Targeting global regulators is an effective strategy to activate silent gene clusters as demonstrated in several Streptomyces studies. In P. fluorescens Pf0-1, a silent gene cluster encoding a novel CLP (gacamide) was identified by genome mining and subsequently activated by repairing a defective gacA through complementation (Jahanshah et al., 2019).

LP mono-producers are found in the P. fluorescens, P. putida and P. syringae group and possess roles in surface motility, biofilm formation or break down, solubilization of nutrients, protection against competitors and predators, induction of systemic resistance in plants, and interactions with insects (D'Aes et al., 2010; Raaijmakers et al., 2010; Götze and Stallforth, 2020; Oni et al., 2022).

The organization of the BGCs encoding CLPs in mono-producers is very well conserved with usually three NRPS genes arranged in either one operon, or in a split configuration in which the first NRPS gene is located elsewhere in the genome (Cesa-Luna et al., 2023) (Figure 6).

The genomic regions flanking the NRPS gene clusters encoding CLPs typically contain well conserved regulatory and transporter genes. Pseudomonas CLPs are secreted by the PleABC tripartite efflux system, homologous to the MacAB-TolC ABC-type multidrug efflux pump that is found in many Gram-negative bacteria (Fitzpatrick et al., 2017). The PleABC machinery is composed of the inner membrane protein PleB (MacB), the periplasmic adapter PleA (MacA) and the outer membrane protein PleC (TolC) (Girard et al., 2022) (Figure 5). The pleC gene is usually located upstream of the first NRPS gene and preceded by a regulatory gene (luxR1) encoding a protein of the LuxR family. These LuxR proteins contain a typical DNA-binding helix-turn-helix (HTH) motif but lack an N-acylhomoserine lactone (N-AHL)-binding domain. The pleA and pleB genes are positioned downstream of the third NRPS gene and are often, but not always, followed by a luxR-type regulatory gene (luxR2) transcribed in the opposite direction (Figure 6). Recently is was shown that PleB is suitable as a diagnostic sequence for genome mining allowing the detection and/or typing of Pseudomonas LP producers (Girard et al., 2022).

Mutation of either luxR1 (viscAR) or luxR2 (viscBCR) results in loss of viscosin production in P. fluorescens strain SBW25 (de Bruijn and Raaijmakers, 2009). A viscAR mutation in this strain could be complemented with the luxR1 (massAR) regulatory gene of the massetolide producer P. lactis (fluorescens) SS101 (de Bruijn and Raaijmakers, 2009). Mutation of luxR1 (arfF) in Pseudomonas sp. MIS38 likewise leads to loss of arthrofactin production (Washio et al., 2010). Mutation of psoR1 leads to loss of putisolvin production in P. putida PCL1445, while mutation of pleA (macA) or pleB (macB) in this strain leads to reduced putisolvin production (Dubern et al., 2008). In P. protegens CHA0 it has been shown that translation of the LuxR-type regulatory genes orfR1 and orfR2 located up- and downstream of the orfamide BGC are under the direct control of the Gac/Rsm pathway (Sobrero et al., 2017).

LuxR2-type regulatory proteins are lacking in BGCs encoding xantholysin (Li et al., 2013) and entolysin (Vallet-Gely et al., 2010), while the situation is variable for WLIP producers (Figure 6). A luxR2 gene is present downstream of the WLIP BGC in P. yamanorum LMG27247 (wlyR2), P. fluorescens LMG5329 (wipR2) and two WLIP producers from the P. putida group (P. xanthosomae COR54 (wlfR2) and P. fakonensis COW40 (wlfR2)), but absent in P. wayambapalatensis RW10S2 (Rokni-Zadeh et al., 2012) (Figure 6). The LuxR1 regulators XltR, EltR, WlpR and WipR are required for xantholysin, entolysin and WLIP production in the respective strains P. mosselii BW11M1, Pseudomonas entomophila L48, P. wayambapalatensis RW10S2, and P. fluorescens LMG5329 (Vallet-Gely et al., 2010; Rokni-Zadeh et al., 2012; Li et al., 2013; Rokni-Zadeh et al., 2013). Intriguingly, all LP BGCs that lack luxR2 are produced by strains that taxonomically belong to the P. putida group (Girard et al., 2021). The reason for this is unknown but could symbolize a different ecological role.

Mono-producers of the LLPs syringafactin and cichofactin are described in various phylogroups of the P. syringae group (Bricout et al., 2022; Cesa-Luna et al., 2023) and in some isolates of the P. putida group (Cesa-Luna et al., 2023). LLPs are encoded by two NRPS genes arranged in an operon. Homologues of luxR1 and luxR2 are present upstream and downstream of the BGC encoding syringafactin in P. syringae DC3000, and cichofactin in P. putida 4A7, but the pleC transporter gene is absent in P. syringae DC3000 (Berti et al., 2007) and no transporter genes are present in the BGC encoding cichofactin in P. putida 4A7 (Figure 6). Mutation of luxR1 (also called syfR), but not of luxR2 (pspto2833) leads to loss of syringafactin production in P. syringae DC3000 (Berti et al., 2007). P. bijieensis L22-9 is a thanafactin mono-producer. The thanafactin BGC lacks regulatory genes, and a major facilitator superfamily (MFS) transporter is located downstream of the BGC (Figure 6).

Phylogenetic analysis shows that LuxR1 and LuxR2 proteins from CLP and LLP mono-producers form two distinct phylogenetic groups (Figure 7).

Regulation of LP production in mono-producers in response to cell density by autoinducers, also called quorum sensing regulation (Venturi, 2006), has to our knowledge only been described for the viscosin producer P. fluorescens 5064 (Cui et al., 2005) and in the putisolvin producer P. putida PCL1445. P. fluorescens 5064, an opportunistic soft rot pathogen of broccoli, produces the 9:7 CLP viscosin that is important for plant surface colonization. The N-acyl homoserine lactone (HSL) quorum sensing signal 3-OH-C₈-HSL regulates viscosin production in this strain (Cui et al., 2005). P. putida PCL1445 was isolated from grass roots grown in soil polluted with polyaromatic hydrocarbons. The strain produces the 12:4 CLPs putisolvin I and II. These compounds inhibit biofilm formation and break down existing Pseudomonas biofilms (Kuiper et al., 2004). The ppuI-rsaL-ppuR quorum sensing system is involved in putisolvin production and mutants impaired in either ppuI or ppuR show a severe reduction in putisolvin production (Dubern et al., 2006). The quorum-sensing signals 3-oxo-C₁₀-N-acyl homoserine lactone (3-oxo-C₁₀-AHL) or 3-oxo-C₁₂-AHL induce expression of the biosynthesis genes activating production of putisolvin I and II in this strain (Dubern et al., 2006).

Additional compounds involved in LP regulation in mono-producers include heat shock proteins, Clp proteases and enzymes involved in amino acid metabolism (Figure 5). The Hsp70 class heat shock protein DnaK regulates putisolvin production together with DnaJ at low temperature in P. putida PCL1445 (Dubern et al., 2005). DnaK is also involved in the regulation of massetolide in P. lactis SS101 (Song et al., 2014). Mutation of the gene encoding the Hsp90 class heat shock protein HtpG leads to loss of arthrofactin synthesis, while arthrofactin biosynthesis genes are normally expressed suggesting a role in posttranscriptional processes (Washio et al., 2010). Heat shock proteins may be required for the proper folding of positive (LuxR-type?) transcription factors or for assembly of the NRPS complex (Song et al., 2014). In P. putida, DnaK is under the control of the Gacs/GacA regulatory system, but this may not be the case in P. lactis SS101. In P. lactis SS101 the serine protease ClpP and its chaperone ClpA are required for massetolide biosynthesis. ClpP is an ATP-dependent serine protease that associates with different ATPases, including ClpA. ClpA selects target proteins for degradation by ClpP. Transcriptomic and proteomic analyses suggest that the ClpAP complex regulates massetolide biosynthesis via the LuxR1 transcriptional regulator MassAR, the heat shock proteins DnaK and DnaJ and via proteins involved in the citric acid cycle (Song et al., 2015a) (Figure 5). Additional regulators identified in P. lactis SS101 include D-3-phosphoglycerate dehydrogenase (Phgdh) and the antisigma factor PrtR (Song et al., 2014). Phgdh is involved in the biosynthesis of L-serine, an amino acid that makes up two of the nine amino acids in massetolide. PrtR interacts with extra-cytoplasmic function sigma factors of the sigma 70 family and regulates the expression of luxR1 and luxR2 by an unknown mechanism (Figure 5).

Concerning environmental signals that regulate LP production in mono-producers Mazzola et al. (2009) showed that the massetolide and viscosin biosynthesis genes, massABC and viscABC respectively, in P. lactis SS101 and P. fluorescens SBW25 are upregulated upon protozoal exposure conferring protection to each strain against predation. Interestingly, the authors observed that physical contact between prey and predator was not necessary to activate the massABC and viscABC genes. It would be interesting to test extracts from different protists in order to identify specific protist signals that trigger CLP production.

So, for LP mono-producers, quorum sensing regulation is not common, the LuxR1 regulator is always present (with the notable exception of the thanafactin producer P. bijieensis L22-9) and essential for LP production, while LuxR2 is sometimes lacking and when present, not always essential for LP production. LuxR1 and in some cases also LuxR2 are under translational control of the Gac/Rsm regulon. LP production is additionally regulated by heat shock proteins and the ClpAP complex.

LP poly-producers belonging to the P. syringae group, or P. corrugata and P. mandelii subgroup within the P. fluorescens group, typically co-produce CLPs from the Peptin and Mycin family and often produce an additional LLP of the Factin family, while poly-producers of the P. asplenii subgroup produce an additional CLP of the Asplenin family (Girard et al., 2020). Strains that produce a CLP of the Tolaasin family belonging to the P. fluorescens subgroup co-produce a CLP of the Viscosin family, while some members of the P. protegens group co-produce sessilin (a member of the tolaasin family) and orfamide (Cesa-Luna et al., 2023). Regulation of LP production in poly-producers is more complex than in mono-producers and can be quorum sensing dependent or independent.

Owing to the complexity of natural environments, a significant proportion of the Pseudomonas-LP research performed to date is only based on lab-based analyses. Production of several CLPs in the environment notably amphisin, tensin and viscosinamide has been detected, for example, in the sugar beet rhizosphere (Nielsen and Sorensen, 2003). Interestingly in this study no LPs were found in bulk soil signifying production is niche specific (Nielsen and Sorensen, 2003). Production of LPs by other isolates was also quantified on beet seeds and found in ranges of 0.22–0.65 µg CLP per seed. The P. fluorescens strain DSS73 (originally isolated from the sugar beet rhizosphere) was shown to produce amphisin on germinating sugar beet seeds in soil correlating with lab-based findings that unknown components of a sugar beet extract induce amsY expression needed for amphisin production (Koch et al., 2002). The environmental conditions required for amphisin production thus appear to reflect the producing strain’s specific habitat. A similar observation was made for P. syringae pv syringae where conditions required for lab-based production of syringomycin reflect the environmental conditions required for pathogenesis (Gross, 1985). As amphisin has antifungal properties it would be interesting to determine whether components of fungal extracts also induce amsY expression. Interestingly, the concentration of LPs detected over time remained similar in sterile soil whereas levels in non-sterile soil were rapidly reduced suggesting degradation by indigenous microbes (Nielsen and Sorensen, 2003).

While numerous studies have indicated that LPs are susceptible to degradation in the environment, the detailed mechanisms underpinning this process remain obscure. However, a key outcome of such research to date is the finding that microbial degradation of LPs can result in structural changes that alter their biological activities. For example, in Bacillus, degradation of surfactin generates a linear surfactin which can no longer induce systemic resistance (ISR) in tobacco (Rigolet et al., 2023). Linear surfactant also displays a reduced ability to lower surface tension when compared to cyclic surfactin (Liu et al., 2015). In the common button mushroom, protective helper bacteria disarm the causal agent of brown blotch P. tolaasii by enzymatic linearization of the toxin tolaasin and surfactant pseudodesmin, required to colonize the mushroom cap, to yield inactive linear forms of the CLPs (Hermenau et al., 2020). The linearization of CLPs is thought to be a resistance mechanism against competing bacterial species (Hermenau et al., 2020). More recently (Hansen et al., 2023), reported the cooperative degradation of orfamide A by Rhodococcus globerulus D757 and Stenotrophomonas indicatrix D763 to protect orfamide-sensitive members of a synthetic community during co-culture with the orfamide-producer P. protegens. It has been proposed that degradation of CLPs could be a strategy deployed by competitors to prevent CLP producers from performing “critical” functions in their environment such as biofilm formation, enhancing motility or colonization of specific niches (Rigolet et al., 2023). In another recent study (Zhang et al., 2021), present a new role of LPs in mediating bacterial cooperation to evade amoebal predation. Synthesis of syringafactins by Pseudomonas sp. SZ57 induces peptidase production in Paenibacillus sp. SZ31 resulting in partial LP-degradation yielding a mix of modified natural products that become amoebicidal (Zhang et al., 2021). These findings expand current knowledge on the ecological functions of LPs and demonstrate how interactions with other microbes can be exploited to unlock production of new compounds. Additionally, the identification of specific compounds capable of natural product modification could be exploited to select for the synthesis of LPs with specific functions during, for example, lab or large-scale cultivations (see Section 6.4).

Going forward it will be critical to quantify production of LPs in situ and correlate expression of LP-associated genes to LP production. Moreover, information on the stability of LPs in the environment, the mechanisms by which they are degraded and importantly the impact of degradation on LP structure and function is key for developing applications in environmental biotechnology and potentially opens a new avenues for natural product discovery.

LP production is influenced by growth phase in addition to abiotic factors (e.g., temperature, pH, oxygen) and nutritional factors (e.g., carbon, nitrogen and phosphorus sources, trace elements) (Raaijmakers et al., 2006). However, comparable studies are limited and the information available is scattered across a handful of strains (Nybroe and Sorensen, 2004). Optimization of growth conditions required for LP production is best described for syringomycin in P. syringae (Gross and DeVay, 1977; Gross and DeVay, 1977; Gross, 1985; Grgurina et al., 1996) showed higher levels of syringomycin production in still potato dextrose broth cultures (PDB) compared to aerated cultures. Viscosinamide is also produced in still cultures or under carbon, nitrogen or phosphorus starvation (Nybroe and Sorensen, 2004). Whereas syringomycin and syringopeptin are produced in the stationary phase, production of viscosinamide, tensin and amphisin is growth-coupled and occurs in the exponential phase (Nielsen et al., 1999; Nielsen et al., 2000; Koch et al., 2002). Syringomycin is regulated by iron (>2 µM), requires L-histidine as a nitrogen source and is repressed by inorganic phosphate (Gross, 1985). In plant tissues, levels of iron are high whereas concentrations of inorganic phosphate are not sufficient to inhibit the phytotoxin demonstrating that the environmental conditions required to support syringomycin production correlate with those necessary for pathogenesis (Gross, 1985).

While pH had no effect on syringomycin production, temperature was identified as an important factor with optimal production recorded at 24°C (Gross, 1985). In P. nunensis In5, regulation of nunamycin and nunapeptin is also temperature-dependent with optimal production at 15°C. Antifungal activity of In5 decreases with increasing temperature correlating with a reduction in production of both LPs (Michelsen and Stougaard, 2011; Christiansen et al., 2020). Production of putisolvin in P. putida PCL1445 is likewise temperature dependent with the highest production at 11 °C (Dubern et al., 2005). Also, syringafactin production in P. syringae pv. syringae is thermoregulated with much higher production at 20°C than at 30°C (Hockett et al., 2013). The molecular mechanisms underpinning the effects of temperature on CLP production are not well known. It has been suggested that temperature may directly impact synthetase formation and thereby alter iron uptake required for specialized metabolism (Gross, 1985). For P. putida PCL1445 it was shown that low temperature positively regulates putisolvin production during the late exponential phase via the DnaK stress response system (Dubern et al., 2005).

Screening for cultivation conditions inducing nunamycin and nunapeptin production revealed that nunapeptin is produced on a range of media both liquid and agar-based personal communication. In contrast, nunamycin production occurs on select agar-based media or in defined liquid minimal media supplemented with either glucose, glycerol and trehalose as carbon source (Christiansen et al., 2020). Glycerol also supports viscosin production by P. antarctica and has been used as a carbon source for rhamnolipid production (Zhao et al., 2021; Ciurko et al., 2023). Carbon source has been shown to influence tensin production in P. fluorescens, phytotoxin production in P. syringae and fungitoxin production in P. nunensis In5 (Nielsen et al., 2000; Woo et al., 2002; Christiansen et al., 2020). In contrast to nunapeptin, production of nunamycin appears to be tightly regulated occurring only under select conditions at low quantities. Nunamycin is a potent antimicrobial peptide and therefore potentially toxic to In5. Interestingly, production of syringomycin which displays a similar structure to nunamycin is not toxic to P. syringae at concentrations naturally produced (Gross, 1985). Gross (1985) observed that high syringomycin titers did not negatively impact bacterial growth as comparable cell densities were recorded for producers and non-producers. Finally, culture optimization has also been reported for thanamycin production in P. fluorescens SH-C52 resulting in a 3.3-fold product increase sufficient to recover 40 mg/120 L culture for NMR studies however the specific parameters altered were not specified (Johnston et al., 2015).

One major obstacle in natural product research is activating silent BGCs and/or improving the production of those expressed at low levels. For Pseudomonas LPs the challenge typically resides in low titers rather than no production. Several approaches can be used to activate or enhance BGC expression including cultivation-based approaches, molecular based techniques or synthetic biology strategies and combinatorial chemistry (Reen et al., 2015).

Cultivation-based approaches involve optimization of cultivation conditions, e.g., nutrients, carbon source, aeration, pH, temperature or using environmental cues, e.g., chemicals to induce compound production. While some information on optimal conditions needed for LP production exist, more systematic screening of culture conditions and in particular the identification of specific environmental signals that positively (or negatively) regulate expression of LP-associated genes is needed. In the well-studied and prolific producers of SMs actinomycetes, a number of key triggers have been identified to access antibiotic production including chemicals, microbial metabolites, interactions with microbes, environmental factors and enzymes that could serve as a useful starting point for studies in Pseudomonas (Seyedsayamdost, 2014; Zhu et al., 2014; Rosen and Seyedsayamdost, 2017; Zong et al., 2022). Information on the regulatory pathways and environmental signals influencing LPs can then be integrated into cultivation-based approaches and depending on the application, combined with other strategies, e.g., strain engineering to further enhance LP production.

Pseudomonas spp. are becoming increasingly attractive cell factories for the production of high-value chemicals including native and non-native SMs in part due to their capacity to utilize cheap carbon sources (Nikel et al., 2014; Wang et al., 2020). Using renewable resources, e.g., agricultural and food waste would greatly increase the commercialization potential of CLPs making bioprocessing of these compounds more sustainable (Ceresa et al., 2023). Importantly, Pseudomonas spp. are naturally competent and typically well-suited to genetics and molecular research. When selecting strains (particularly environmental isolates) for improved LP production it will be necessary to determine their genetic tractability and consider their origin, e.g., beneficial or pathogen. Select strains can then be engineered using traditional techniques, e.g., homologous recombination or next-generation CRISPR-Cas9 based technologies for increased production of target LPs either by introducing genes that promote LP production, deleting genes that inhibit LP biosynthesis, or overexpressing genes using native or synthetic promoters (Batianis et al., 2020). Depending on the complexity of the regulation, simple single-gene mutations to more complex multi-gene knockouts and/or insertions may be required.

In Pseudomonas LP-producers, genetic manipulation of regulatory genes has been limited to functional genomics studies and not used to improve LP titers. Interestingly, a study investigating the regulation of massetolide and viscosin demonstrated that the massA gene encoding the first NRPS needed for massetolide production can be heterologously expressed to complement a viscA mutant deficient in the first NRPS needed for viscosin production (de Bruijn and Raaijmakers, 2009). However, for the LuxR-type regulators only massAR (luxR1) and not massBCR (luxR2) could restore viscosin production in the viscAR (luxR1) mutant (de Bruijn and Raaijmakers, 2009) indicating that while some LP genes can be exchanged among different Pseudomonas strains, differences in the functionality of structural and regulatory genes may be at play.

Metabolic engineering can be used to increase LP production using strong indigenous or artificial promoters to increase the copy number of biosynthesis and/or regulator genes or alternatively using inducible promoters to overexpress entire gene clusters. For example, the use of promoter systems to improve LP production has been successful in the rhamnolipid producing strains P. aeruginosa, Burkholderia kururiensis and P. chlororaphis (Chong and Li, 2017). Overexpression of the rhlAB operon required for rhamnolipid biosynthesis under the tac promoter in B. kururiensis yielded a mixture of over 50 rhamnolipid congeners (Tavares et al., 2013). Similarly, overexpression of the rhamnolipid biosynthesis gene rhlC in P. chlororaphis resulted in the synthesis of di-rhamnolipids instead of mono-rhamnolipids (Solaiman et al., 2015). These findings demonstrate the potential of metabolic engineering-based approaches not only to increase product titer but equally to alter the structure and function of LPs.

Synthetic biology can serve to overcome challenges in engineering native strains for large scale production, for example, by refactoring BGCs to reduce the complexity of LP regulation (Temme et al., 2012; Wang et al., 2021). However, cloning and heterologous expression of large NRPS gene clusters that can span over 100 Kb (Meleshko et al., 2019) is difficult. Drawbacks include PCR amplification or synthesis of large DNA fragments, decrease in cloning efficiency, stability of vectors and/or successful integration onto the chromosome. Moreover, compared to Pseudomonas spp., classical hosts such as E. coli and yeast are not natural producers of LPs and may, depending on the LP encounter toxicity issues. Furthermore, with the rapid development of advanced gene engineering techniques, e. g., CRISPRi toolbox precise editing of model and natural Pseudomonas genomes for the controllable manipulation of gene expression will greatly facilitate strain engineering strategies for improved production in native hosts or other Pseudomonas strains (Batianis et al., 2020; Wirth et al., 2020).

For pharmaceutical or health-related applications where high purity grade compounds are needed it may be more desirable to synthesize LPs in vitro. While total chemical synthesis of the Pseudomonas CLPs from the Viscosin, Bananamide and Entolysin family has been achieved, the synthesis of larger LP molecules, for example, found in the poly-producers is more challenging (De Vleeschouwer et al., 2014; De Vleeschouwer et al., 2016; De Roo et al., 2022; Ji et al., 2023; Muangkaew et al., 2023). Chemical synthesis is also used to increase the chemical diversity of molecules, for structure-function studies, and to elucidate the stereochemistry of CLPs (Steigenberger et al., 2021; De Roo et al., 2022; Muangkaew et al., 2023).

Limited studies on optimization of the conditions necessary to induce synthesis and improve production of LPs exist. Consequently, no Pseudomonas-derived LPs are currently commercially available with the exception of rhamnolipids used in broad-spectrum applications and produced at industrially viable yields (Soberón-Chávez et al., 2021).

Bioreactor-scale production of LPs has only been reported for pseudofactins (PFs), 8:6 CLPs from the Bananamide family in P. fluorescens BD5 (Biniarz et al., 2018; Biniarz et al., 2020). Optimal production of PFs requires high glycerol (80 g/L) and tryptone (15 g/L) concentrations with high culture aeration (30 L/min) to achieve 7.2 g/30 L yield of PFs (Biniarz et al., 2018). LPs are often produced as a mixture of LPs comprising variants with minor structural changes that can greatly impact their bioactivity (Dufour et al., 2005; Eeman et al., 2006; Biniarz et al., 2020). Biniarz et al. (2018) observed that media supplementation with valine and leucine causes a shift in the ratio of PFs produced and can be used to select structural LP variants. The selective production of LPs will greatly benefit the purification of LPs for structure-function studies and potentially provide opportunities to identify new LP variants with novel biological activities.

Going forward it will be important to determine the limiting factors of LP production across scales (lab-bioreactor-technical scale). Additional parameters influencing growth and metabolic activity to optimize for include temperature, pH, oxygen, agitation, speed or vessel type (Guez et al., 2021). More basic studies on media and culture conditions required for LP production are needed to (i) routinely produce LPs of interest at lab scale for biological and chemical characterization, and (ii) to scale production from lab-to-bioreactor to develop an efficient bioprocess for LP production. Moreover (Gross and DeVay, 1977), observed that syringomycin production varied considerably across different P. syringae strains. Thus, it will be important to determine the specific impact of growth phase, carbon source, nutrients and inducer molecules on LP production in individual strains and to tailor culture conditions accordingly.