1. Introduction
Human population increase, environmental degradation, and climate change pose significant challenges to sustainable food crop production. It is essential that the yield of crops is increased while minimizing negative impacts on the environment and ensuring economic sustainability. Selective plant breeding has increased crop productivity with the trade-off that the resulting varieties require substantial inputs of mineral fertilizers and pesticides to deliver high yields. Both traditional breeding and genetic engineering approaches for crop plant improvement have the disadvantages of long development times, technical limitations, and potential problems with public acceptance (reviewed in [1,2,3]).
An alternative approach to increasing plant productivity relies not on their genetic modification but on promoting their growth using plant growth-promoting microbes (PGPM) [4]. PGPM are part of the soil microbiome, which includes both microbes in the soil and their metabolic or structural elements, such as polysaccharides and secondary metabolites. PGPM species are adapted to interact with plant species to form relatively species-specific plant rhizosphere communities or rhizomicrobiomes. The PGPM component of the rhizomicrobiome includes mycorrhizal fungi, plant growth-promoting archaea, and plant growth-promoting rhizobacteria (PGPR). The PGPR are the most numerous members of the PGPM and enhance plant growth and stress resistance by increasing nutrient availability to the plant, producing phytohormones, stimulating plant defense responses against abiotic stresses and pathogens, and/or fixing nitrogen. These and other mechanisms underlying plant growth promotion by PGPR have been extensively reviewed [1,5,6,7,8,9,10,11] and will not be treated in detail here.
Much of the ability of PGPR to increase plant growth derives from their ability to modify plant root architecture and growth and thereby increase nutrient uptake [12]. These changes in the roots can be mediated by phytohormones (auxins, cytokinins, gibberellins) produced by PGPR, or by PGPR manipulating plant ethylene levels. PGPR affect gene expression for signaling and developmental pathways in plants, but the chemical signals involved are largely unknown [9]. Some PGPR protect plants against pathogens by niche competition, producing antibiotics and siderophores, and stimulating systemic defense systems against pathogens. Diazotrophic PGPR use the enzyme nitrogenase to reduce (fix) biologically inert atmospheric nitrogen to biologically usable ammonia. Reduced nitrogen is often the limiting nutrient in agriculture, and so diazotrophs are of great practical importance in the nitrogen nutrition of crop plants as well as having other plant-growth-promoting activities. For general reviews on PGPR in agriculture, see [1,11,13,14,15,16,17,18].
The presence of genes encoding plant growth-promoting traits in a given bacterium does not mean that it will promote plant growth. This stems from the differing degrees of plant–PGPR specificity and the existence of unknown factors involved in chemical signaling and nutrient exchange between the partners that result in growth promotion [9,19,20]. The complexity of interactions between different PGPR in synthetic communities (symcoms) that were inoculated onto corn was evident from the very dissimilar abilities of different syncoms to increase plant growth and disease resistance [21].
The gains in plant health and productivity brought about by PGPR have long shown practical benefits in agriculture through the application of commercial PGPR inoculants to certain crops (reviewed in [1,6,8,11,22]). The ability of PGPR inoculants to increase crop yields has been uneven in practice and the potential of the technology is underrealized. To overcome present limitations in PGPR efficiency and application, we need to learn more about how different PGPR species interact among themselves and with a variety of crop species under diverse environmental conditions. This knowledge can then be applied to developing more effective and reliable PGPR inocula, including formulations containing syncoms assembled using systems biology approaches [4,21,23].
For the establishment of plant–PGPR relationships, the bacteria best adapted to utilize the nutrients present in root exudates are generally the most successful at colonizing the roots. The root exudate composition thus shapes the microbial composition of the rhizosphere [20,24]. Well known examples of compounds involved in chemical communication include legume flavonoids, rhizobial nodulation factors, exopolysaccharides (EPS), and phytohormones produced by both plants and PGPR [8,15,17,25,26,27].
Polyamines (PAs) are primordial polycations with important or essential functions in all living things, such as physiological effectors and signal molecules in plant–microbe interactions. Pas are short hydrocarbons with two or more amino groups that are positively charged at physiological pH. The positive charge on PAs lets them associate with negatively charged macromolecules, such as nucleic acids, phospholipids, and some proteins, in the cell. These associations mediate cell physiology by changing the chemical or physical properties of the macromolecules with which they interact or by directly regulating their biological activities. PA catabolism generates reactive nitrogen and oxygen species that serve as signal molecules, and PAs can also be used by bacteria as sources of carbon and/or nitrogen [28].
The PAs commonly found in organisms include the diamine putrescine (H2N-(CH2)4-NH2), the triamine spermidine (H2N-(CH2)3-NH-(CH2)4-NH2), and the tetraamine spermine (H2N-(CH2)3-NH-(CH2)4-NH-(CH2)3-NH2). Spermidine (Spd) has an essential role in eukaryotes and archaea because it is required for the hypusine chemical modification of their IF5A translation factors. In eukaryotes, PAs are also important in cell growth and proliferation, while in archaea, less is known about their functions apart from their role in translation. PAs in bacteria can either be produced endogenously or transported from the environment. General reviews on PA metabolism and functions include [29,30,31,32].
Bacteria are especially versatile in their ability to synthesize many kinds of PAs. This may be because PAs in bacteria are involved in diverse processes, including growth, motility, gene expression, and biofilm formation [29,30,33,34,35]. PAs also condition pathogenic and mutualistic interactions between a wide range of bacteria and their animal or plant hosts (reviewed in [25,30,32,34,35,36,37,38,39,40,41]. In this review, we describe the production and function of PAs in PGPR and the plants with which they interact. In Section 2, we look at the basic ecology of PGPR, with an emphasis on those species that fix nitrogen in association with plants. In Section 3 and Section 4, we examine the metabolism and the roles of PAs in PGPR and plants, respectively. How PAs act as signal molecules and physiological effectors in the interaction of PGPR and plants and conclusions and perspectives are the subjects of Section 5 and Section 6.
2. The Rhizomicrobiome
The rhizosphere is broadly defined as the soil surrounding and chemically or physically affected by the root, including root tissues that are colonized by microbes [8]. The terms endorhizosphere, rhizoplane, and ectorhizosphere describe the spatial proximity and the intimacy of the interaction of microbes with the roots (Figure 1). The endorhizosphere is the innermost rhizosphere zone and is made up of the apoplastic (extracellular) fluid-filled spaces around root cells as well as their interior. The rhizoplane refers to the surface of the root, including mucilage. The ectorhizosphere begins at the outer limit of the rhizoplane and extends a short distance (millimeters) into the bulk soil [42] (Figure 1). Distinct species of free-living, associative, and endosymbiotic PGPR inhabit one or more of these rhizospheric niches [43].
Many species of free-living bacteria live in the bulk soil, but their greatest populations occur in the rhizoplane and ectorhizosphere, where root exudates act as selective chemoattractants and nutrients [14,24,44]. Free-living bacterial diazotrophs occur in the phyla Firmicutes, Cyanobacteria, and Proteobacteria. Rhizosphere-dwelling Azospirillum, Azotobacter, and cyanobacteria (mostly Anabaena and Nostoc) species increase plant growth by fixing nitrogen and/or producing auxin [17,45]. All of the diazogrophic archaea are methanogens in the phylum Euryarchaeota [20,26].
Associative diazotrophs have facultative interactions with plants and live in the rhizoplane or endophytically in the endorhizosphere. These microbes partner with a wide variety of plants, including cereals, ferns, and bryophytes (non-vascular plants) [46]. The root-colonizing alpha-proteobacterium Rhodopseudomonas palustris, for example, promotes the growth of Chinese cabbage by increasing auxin levels in leaves and nitrate uptake by roots [19]. Rhizoplane and endorhizospheric diazotrophic PGPR can improve plant growth by fixing nitrogen, increasing soil nutrient availability and/or uptake, secreting phytohormones and enhancing plant pathogen resistance by niche competition, or stimulating plant resistance responses [27]. Genome sequence analysis revealed genes for all of these traits in the rhizosphere bacterium Klebsiella variicola D5A. Genes for PA production were defined among those that confer PGPR fitness in this strain [47,48].
Endophytic nitrogen fixers like Azoarcus and Nostoc (Pseudomonadata and Cyanobacteria, respectively) directly access plant tissues via fissures at the sites where lateral roots emerge or through other natural openings. Diazotrophic endophytes remain outside of plant cells, fix nitrogen and contribute other PGPR activities to enhance the growth in crops like wheat, rice and corn [16,45]. Despite the tight association of endophytic species with plants, they do not damage plant tissues or elicit defense responses [46].
Intracellular diazotrophic endophytes reside inside root cells and fix atmospheric nitrogen for their hosts. Rhizobia is the collective term for bacteria that initiate a chemical dialogue with selected (mostly leguminous) hosts and induce the formation of specialized organs called nodules on the roots. Metabolically differentiated rhizobia living inside of plant cells of the nodule are called bacteroids, and they receive photosynthate-derived carbon sources from the host in exchange for reduced nitrogen. Rhizobia include alphaproteobacteria that elicit the formation of root nodules on legumes or the non-legume Parasponia and include species of Rhizobium, Sinorhizobium, Mesorhizobium, Bradyrhizobium and Allorhizobium [46,49,50]. Rhizobia belonging to the beta-proteobacteria include species in the genera Burkholderia and Cupriavidus, which nodulate Mimosa spp. and a few other legumes [51]. The Gram-positive actinobacterial genus Frankia forms nitrogen-fixing nodules on some actinorhizal plants and also fixes nitrogen in free life, unlike the majority of nodule-forming rhizobia. Azorhizobium caulinodans (phylum Pseudomonadota) forms both stem and root nodules on Sesbania rostrata and can also fix nitrogen asymbiotically [49]. In addition, some Rhizobium species are intercellular endophytes that reside in the stems of non-legumes like cottonwood trees and are thought to fix nitrogen as members of the PGPR consortia found there [52,53].
Globally, endosymbiotic rhizobia fix more nitrogen than free-living or associative diazotrophs. Many legumes are important crops for human consumption or fodder or are used in crop rotation with nonlegumes to provide a partial replacement for chemical nitrogen fertilizer [11]. Rhizobia also promote the growth of non-legumes, such as rice and radish, by mechanisms other than nitrogen fixation, including siderophore production, phosphate solubilization and indole-3-acetic acid (IAA) production [54,55]. Non-rhizobial PGPR inoculated onto legumes in combination with compatible rhizobia can increase nodulation, nitrogen fixation, and crop yields [56,57].
Several genera of archaeal methanogens promote plant growth by mechanisms similar to those of the bacterial PGPR [18]. Methanogens that possess the gene encoding the iron protein of nitrogenase (nifH) are presumed to be nitrogen fixers and are common in many marine environments and wetlands [58]. Much less is known about their interactions with plants than for bacterial PGPR [26].
3. Polyamine Metabolism and Functions in PGPR
In this section, we consider the metabolism (Figure 2) and physiological functions of PAs in PGPR. Later sections will describe these aspects in plants and in PGPR–plant interactions.
3.1. Polyamine Biosynthesis in PGPR
The diamine putrescine (Put) is made by the decarboxylation of
Whether bacteria exclusively or principally use Odc or Adc to make Put may largely depend on which basic amino acid substrate was most available during their evolution [59]. In PGPR, Put production by Odc appears to be the more common route. This is true in nodule-forming rhizobia and in the sugarcane endophyte Gluconacetobacter diazotrophicus PAI 5 [36,60,61,62]. In contrast, the endophytic Klebsiella spp. LTGPAF-6F and D5A and the heterocyst-forming cyanobacterium Anabaena sp. PCC 7120 use Adc and agmatinase to produce Put from Arg [47,63,64]. Genome analysis of the endophytes Klebsiella pneumoniae 342 (Enterobacteriaceae) and Methylobacterium P1–11 (Methylobacteriaceae) and the endosymbiotic Tardiphaga P9–11 (Bradyrhizobiaceae) show both Adc and Odc as potential enzymes for Put biosynthesis [65,66].
Orn is an intermediate in the Arg biosynthetic pathway, where it is formed by the deacetylation of N-acetylornithine. The Medicago endosymbiont S. meliloti produces several non-homologous N-acetyl-ornithine deacetylase paralogs to produce Orn. Orn is also produced by the hydrolysis of Arg by arginase ArgI1 [67,68,69]. In S. meliloti Rm8530, about 90% of the free intracellular Put is made by a L/Odc designated Odc2 [60], with the remainder being of unknown origin. It seems that metabolism in S. meliloti is geared towards the production of Orn for making Put as well as Orn-based siderophores and Orn-containing lipids [69].
Homologs of the S. meliloti Odc2 occur in other species of nitrogen-fixing rhizobia and in A. tumefaciens [36]. The Odc activity of the S. meliloti Odc2 is ten-fold higher than its Ldc activity, and odc2 mutants of strain Rm8530 produce very low levels of Put, homospermidine (HSpd) and Spd [60]. An additional Odc in S. meliloti, called Odc1, is specific for Orn as a substrate but has very low specific activity compared to Odc2. The inactivation of odc1 does not alter PA production under any growth condition tested [60].
The diamine cadaverine (Cad) is produced by Ldcs that are specific for L-lysine (Lys) or by L/Odcs [33,70,71]. Ldcs are present in some diazotrophic actinobacteria, firmicutes, and proteobacteria [36,72]. Among associative or free-living diazotrophs, Azotobacter spp., including A. vinelandii, have high levels of Put and lower levels of Cad and Spd in cells from minimal medium cultures. Low amounts of 1,3-diaminopropane (DAP) were detected in less than one-third of the strains analyzed [73,74]. The Azosprillum brasilense PGPR commercial inoculant strains Az39 and Cd produced and excreted high levels of Spd (>100 nmoles/mL) and 5–20 nmoles/mL of Put, spermine (Spm) and Cad when grown in minimal medium. When the supernatants of cultures grown with exogenous Lys were analyzed, Spd content was lowered by more than 90%, and Cad increased about 3-fold. The increase in Cad production when the medium was supplemented with the Ldc substrate, Lys, is not unexpected. How Lys decreases Spd biosynthesis (via Spd synthase), increases its catabolism or export is not known [75,76] (Figure 2). Most of the Cad produced by S. meliloti Rm8530 is linked to macromolecules rather than being exported or present in free form in the cytoplasm [60].
The triamine Spd is made by one of two different pathways (Figure 2). In the canonical pathway, Spd synthase (EC 2.5.1.16) transfers the aminopropyl group from decarboxylated S-adenosylmethionine (dcSAM) to Put to produce Spd. The dcSAM required for this reaction is produced by S-adenosylmethionine decarboxylase (SAMdc, EC 4.1.1.50).
The canonical pathway exists in some bacteria and in many eukaryotes. The Klebsiella spp. LTGPAF-6F and D5A and Gluconacetobacter diazotrophicus Pal 5 are diazotrophic endophytes whose genomes encode the canonical Spd pathway enzymes SAMdc (SpeD) and Spd synthase (SpeE) [47,61,64].
The second, or “alternative”, Spd pathway converts Put and L-aspartate ß-semialdehyde to carboxyspermidine using carboxynorspermidine dehydrogenase (CANSDH, EC 1.5.1.43). Carboxyspermidine is then decarboxylated by carboxynorspermidine decarboxylase (CANSDC, EC 4.1.1.96) to produce Spd (Figure 2). This pathway is found in many species of Bacteroidetes, Firmicutes, and proteobacteria [34]. Put and Spd are the predominant intracellular pAs in the associative species Azoarcus rhizosphaerae, a nitrogen-fixing member of the Fiscus religiosa (sacred fig) rhizomicrobiome [77]. PA screening of a collection of 15 endophytic actinorhizal isolates from mangrove roots showed that two strains with nitrogenase activity produced Put, Spd, and the tetraamine Spm [78].
To make Spm, Spm synthase (EC 2.5.1.22) transfers the aminopropyl group from dcSAM to Spd. Thermospermine (TSpm), the structural isomer of Spm, is made in plants from the same substrates in a reaction catalyzed by TSpm synthase (EC 2.5.1.79) [33,79] (Figure 2).
S. meliloti produces Spd by the “alternative” pathway described above. Neither this pathway nor the canonical Spd synthase pathway is present in most other alpha-proteobacterial rhizobia [36,60,80], consistent with their lack of Spd production [36,81].
Essentially all rhizobia and many other PGPR produce the triamine HSpd, which is formed by the condensation of two Put molecules using homospermidine synthase (Hss, EC 2.5.1.44) [81,82] (Figure 2). The Hss enzymes of B. japonicum, R. leguminosarum bv. viciae and S. meliloti can also use Spd and Put as co-substrates to produce HSpd and DAP ([36,82,83], Dunn, unpublished). Strains of Frankia are unusual in that they do not produce HSpd, but instead synthesize mostly Put and lower amounts of Cad, Spd, and Spm [84].
In contrast to S. meliloti, most legume-nodulating rhizobia produce only Put and HSpd when grown in cultures. HSpd is often the major PA in free-living rhizobia and bacteroids [36,81,85]. The nodule-forming betaproteobacterial genera Burkholderia and Cupriavidus usually contain 2-hydroxyputrescine (hydroxy-Put) and Put rather than Put and HSpd [81,85,86]. Hydroxy-Put biosynthesis in these diazotrophic beta-proteobacteria may result from the action of Put hydroxylases like those of the non-diazotrophic ß-proteobacteria Bordetella bronchiseptica and Ralstonia solanacearum. Hydroxy-Put may function in iron binding, as it forms part of the structure of the alcaligin siderophore produced by some ß-proteobacterial species [87].
HSpd is made by some free-living and associative diazotrophs, including the cyanobacterium Anabaena [30,63,88]. In Anabaena sp. PCC 7120, the enzyme SpeY converts Put to HSpd. SpeY is a homolog of eukaryotic and archaeal deoxyhypusine synthase, required for using Spd to post-translationally modify the IF5A translation factor [63]. In the associative diazotrophs belonging to Azospirillum spp., Put, Spd, and HSpd are major intracellular PAs in cells grown in culture [81,86].
In addition to producing Put, Spd and HSpd, S. meliloti makes the triamine norspermidine (NSpd) (Figure 2). To initiate the NSpd biosynthesis pathway glutamate and L-aspartate ß-semialdehyde are converted to DAP by the sequential activities of diaminobutyric acid (DABA) aminotransferase (DABA AT, EC 2.6.1.76) and DABA decarboxylase (DABA DC, EC 2.6.1.76) [37,60] (Figure 2). NSpd is produced from DAP and L-aspartate ß-semialdehyde by the consecutive actions of CANSDH and CANSDC, the same enzymes that participate in the alternative Spd pathway [37,60,71,89] (Figure 2). In S. meliloti Rm8530 NSpd is principally bound to macromolecules rather than being in free form [60].
Genome sequences of diazotrophic PGPR often encode multiple putative transport systems to take up or export PAs, virtually none of which have been characterized experimentally. These include multi-subunit ABC-type transporters and single-component PA-basic amino acid antiporters. The expression of smc01652 in S. meliloti, annotated as encoding the substrate binding protein of a Put/agmatine ABC transporter, was induced by both agmatine and Put, consistent with the annotation [90]. PA transporters would allow intracellular PA levels to be modulated via PA import and export as well as the dispatch and receipt of PAs as chemical signals (Section 5, [37]). In addition, PA/basic amino acid antiporters might function in acid stress resistance, as described in Section 3.2.2.
In rhizobia and other bacteria, alternative and sometimes compensatory PAs are produced under certain growth conditions. In the plant pathogen A. tumefaciens, only Put and Spd are normally found in cultures of wild-type strain C58. When strain C58 is grown in the presence of the Odc inhibitor difluoromethylornithine (DFMO), it produces Spm. When Spd biosynthesis is mutationally inactivated, it makes HSpd [80,91]. Growing S. meliloti in the presence of exogenous NSpd boosts the level of DAP and reduces levels of Spd, HSpd, and Put [60]. These collateral effects complicate the interpretation of experiments requiring the addition of PAs to cultures, for example, in the chemical complementation of PA biosynthesis mutants.
3.2. Functions of Polyamines in PGPR
As mentioned in Section 1, PAs affect growth, biofilm formation, and motility in many bacteria. PAs in enteric bacteria also affect the translation of mRNAs for some global transcriptional regulators and are important in DNA supercoiling [29,92]. It is not known if either of these phenomena occurs in PGPR. Some of the known effects of PAs on PGPR are summarized in Table 1 and are further described in the following sections.
3.2.1. Requirement for Polyamines for Growth and Development
The requirement for endogenous PA synthesis for bacterial growth varies from dispensable to essential, depending on the species and growth conditions. For example, Bacillus subtilis does not require PA biosynthesis for growth [70], while the plant pathogen A. tumefaciens does not grow if unable to synthesize a PA having a 1,3-diaminopropane moiety such as DAP, Spd, or NSpd [80]. The majority of bacteria simply grow more slowly when PA biosynthesis genes are inactivated, or the enzymes are pharmacologically inhibited [30,93].
The growth of R. leguminosarum bv. viciae 3841 is significantly slowed in cultures that contain the irreversible Odc inhibitor DFMO. Odc activity, intracellular PA levels, and normal growth were largely restored by adding exogenous Put or Spd to the DFMO-containing cultures [82,83] (Table 1). Similarly, a S. meliloti Rm8530 odc2 mutant that produced low levels of PAs and grew at 60% of the wild-type velocity grew normally if cultures were supplemented with Put or Spd [60] (Table 1).
Some PAs inhibit the growth of diazotrophic bacteria when added to cultures. In S. meliloti Rm8530, exogenous 1 mM Put, Spd, or NSpd added to cultures are transported into and accumulate in the cells and modestly reduce the growth velocity of the cultures (Table 1). This may be a direct effect of increasing the intracellular level of the exogenously added PA or due to changes in the levels of other PAs in the cell that result from supplementation with exogenous PAs, as mentioned in Section 3.1.
A recent transposon sequencing (Tn-seq) study identified the hss genes in B. japonicum USDA110 and R. palustris DGA009 as being essential for the growth of these strains in rich medium [94]. A PA profile for R. palustris DGA009 has not been reported, but other strains of the species contain HSpd as their major PA [81,86,95]. B. japonicum USDA110 also produces relatively high levels of HSpd [36,85]. To confirm the results of the Tn-seq analysis, growth experiments with purposely constructed hss null mutants should be done.
In contrast to the apparent essentiality of hss in R. palustris and B. japonicum, Tn-seq analyses of S. meliloti 1021 and R. leguminosarum bv. viciae 3841 showed that none of their respective PA synthesis genes were essential for growth [62,96,97]. The lack of a requirement for hss in S. meliloti is consistent with the normal growth rate of a strain 1021 hss mutant (Becerra-Rivera and Dunn, unpublished). This mutant might grow normally because it produces the potentially compensatory triamines Spd and NSpd [60]. In contrast, HSpd is the only triamine produced by R. leguminosarum bv. viciae 3841 [36]. The generation and phenotypic testing of a hss mutant is needed to confirm the importance of HSpd in this strain [82].
Under nitrogen-limited growth conditions, vegetative cells in filaments of the cyanobacterium Anabaena differentiate into nitrogen-fixing heterocysts. Heterocyst formation in Anabaena sp. PCC 7120 requires HSpd synthesis. As mentioned in Section 3.1, HSpd in Anabaena is synthesized by SpeA (Adc), SpeB (agmatinase), and SpeY (deoxyhypusine synthase-like Hss). Inactivation of the gene encoding any of these enzymes prevents HSpd synthesis and impairs (∆speB and ∆speY mutants) or prevents (∆speA mutant) diazotrophic growth in a medium with N2 as a nitrogen source. Nitrogen fixation does not occur in the speA mutant because its vegetative cells are unable to form heterocysts. The need for HSpd in heterocyst differentiation may arise from its use as a building block of the cell wall or because the lack of its production impacts nitrogen metabolism or signaling pathways necessary for heterocyst differentiation [63].
3.2.2. Functions of Polyamines in PGPR Abiotic Stress Resistance
PGPR in the bulk soil and rhizosphere may encounter salinity, drought, pH, temperature, or oxidative stress. These stresses can significantly reduce populations of PGPR and decrease their ability to interact with plants despite a number of stress defense strategies present in bacteria. The involvement of PAs in resistance to these stresses is described in the following sections, while general stress resistance mechanisms were reviewed by [98,99,100,101,102,103,104,105,106]. For most PGPR, studies on stress resistance have dealt only with how they confer stress tolerance to a plant partner. In rhizobia, stress resistance has been extensively studied ex planta, and the following sections deal exclusively with this work.
Oxidative Stress
Reactive oxygen species (ROS) such as hydrogen peroxide (H2O2), hydroxyl radical (·OH), and superoxide anion (·O2−) are produced from oxygen during normal aerobic metabolism and at higher levels under abiotic stress [107]. In addition to their ability to damage macromolecules, increase mutation rates, and slow growth, ROS also have important signaling and functional roles in the rhizobia-legume symbiosis. Signaling by H2O2 and general mechanisms by which rhizobia contend with oxidative stress have been reviewed [39,104,108,109]. Like other bacteria, rhizobia fight oxidative stress mostly with enzymatic defenses, including superoxide dismutases, peroxidases, and catalases [110,111,112]. PAs represent one of the non-enzymatic defenses against ROS in bacteria [32,113].
TolC encodes a bacterial membrane protein required for exporting toxic substances like antibiotics, disinfectants, and eukaryotic defense compounds out of the cell. The inactivation of tolC in S. meliloti appears to increase the levels of oxidative stress based on the higher expression of genes involved in enzymatic and non-enzymatic oxidative stress resistance mechanisms. Genes whose expression is also induced in the tolC mutant include those encoding Orn decarboxylase odc1 and the Arg/Lys/Orn decarboxylase sma0682 [114]. This indicates that the tolC mutant may be synthesizing additional quantities of PAs in response to oxidative stress. In comparison to the S. meliloti wild-type Rm8530, an odc2 mutant that produces low levels of PAs grows more slowly than the wild type and reaches less than half its cell density when grown in a minimal medium containing 0.33 mM H2O2. However, we hypothesized that the greater peroxide sensitivity of the mutant was at least partly due to its inability to make wild type levels of EPS [115], which protects S. meliloti from stress caused by exogenous H2O2 [116]. A metabolome analysis of the oxidative stress response caused by treating R. leguminosarum bv. viciae 3841 with the synthetic auxin 2,4-dichlorophenoxyacetic acid showed an increase in Cad and Put levels [117], indicating a possible direct role of these PAs in oxidative stress resistance.
The oxidative stress response in archaea is similar to that of bacteria with respect to antioxidant enzymes [118,119], but the involvement of PAs in resisting oxidative stress is not reported.
Osmotic Stress
About 20% of the earth’s arable land is prone to salinity problems. High concentrations of salts cause ionic osmotic stress, while elevated levels of non-ionic solutes like sugars cause non-ionic osmotic stress. PGPR in soil may be exposed to either or both kinds of osmotic stress, especially in the rhizosphere or during plant colonization or infection [102]. PGPR show a wide range of hyperosmotic stress tolerances, and PAs participate in this by acting as chaperones to physically protect macromolecules from damage or possibly as a signal to increase the intracellular concentration of compatible solutes. The ability of PAs themselves to act as compatible solutes is controversial, mainly because they are present in much lower amounts than the well-established compatible solutes like proline, gamma-aminobutyric acid (GABA), trehalose, or sucrose [106,120,121,122].
Growth under saline conditions has opposing effects on intracellular PA levels in Rhizobium tropici CIAT899 and S. meliloti Rm8530, although both species are similarly salt tolerant. In the R. tropici wild type, HSpd levels and Odc activity increased a few-fold under salt stress. A CIAT899 hss mutant that lacked HSpd grew slower than the wild type under salt stress (0.2 M NaCl in rich medium). The growth of the mutant was partially restored when 0.1 mM HSpd was added to the cultures. These results indicate that HSpd is important for osmotic stress resistance in R. tropici [123].
Growing S. meliloti Rm8530 in a minimal medium with 0.3 M NaCl did not alter the Spd concentration but significantly reduced Put and HSpd levels and odc2 gene transcription [60]. Salt shock in E. coli causes a rapid uptake of potassium ions and an increase in endogenous glutamate synthesis. Put, but not Spd, is rapidly exported from E. coli cells to prevent an overload of cations in the cytoplasm [124]. In salt-stressed S. meliloti, Put and HSpd export, along with odc2 downregulation, may account for the low intracellular levels of these PAs. In S. meliloti 1021, salt stress decreased the expression of odc2 and the genes for the PotD and PotF substrate binding proteins of Put and Spd-preferential ABC transporters, respectively [125]. The homologous transporters in E. coli function in Put and Spd import, respectively, rather than in their export. These results suggest that less endogenous PA synthesis and import from the environment lead to the low levels of Put and HSpd found in salt-stressed S. meliloti [60,125]. Furthermore, an S. meliloti Rm8530 hss mutant grew like wild type under salt stress, while a cansdh grew more slowly. Exogenous Spd restored the growth of the cansdh mutant under salt stress, indicating that Spd is important in salt stress resistance in S. meliloti (M. Dunn, unpublished), perhaps by acting as a molecular chaperone. In contrast, the levels of HSpd and Put may be decreased to achieve cytoplasmic ionic balancing.
As in S. meliloti [60,125], the expression of the R. etli CE3 odc2 homolog is lower under salt stress [126]. Changes in PA levels in salt-stressed R. etli have not been reported to see if they correlate with the changes in odc2 gene transcription.
The acid- and salt-tolerant S. fredii strain P220 accumulates intracellular glutamate and potassium and loses HSpd under salt stress. The failure to detect HSpd in supernatants of the cultures indicates that it was not exported from the cells [127], but we calculate that HSpd in the culture supernatants, if analyzed directly without concentration, would likely be undetectable. It was proposed that the reduction in HSpd levels might aid in cytoplasmic ionic charge balancing under osmotic stress [127].
In contrast to most rhizobia, PAs levels in the cyanobacterium Synechocystis sp. PCC 6803 increased under salt or osmotic stress [128], where it was proposed that they act as compatible solutes [121]. Adc mRNA in this organism was more stable under salt stress, although total Adc activity was unchanged between control and high salt conditions [128].
Acid Stress
Acid soils, defined as those with a pH of 5.5 or less, comprise as much as 40% of arable lands worldwide. In many rhizobia-legume combinations, both nodulation and nitrogen fixation are reduced in acid soils [100]. Rhizobia are also exposed to acidic conditions in plant rhizospheres and the symbiosome space surrounding bacteroids in root nodules [102], and so these bacteria have multiple acid resistance mechanisms [100,101,129].
The HSpd that accumulates in acid-stressed S. fredii P220 was proposed to act as a chaperone to protect macromolecules from acid-induced degradation [127]. PA synthesis also consumes protons and alkalinizes the cytoplasm. Bacteria such as E. coli have acid-inducible isozymes of Adc, Odc, and Ldc paired with cognate basic amino acid-PA antiporters. The basic amino acid decarboxylation reactions consume protons and thus raise the intracellular pH in cells under acid stress. The PA produced in the decarboxylation reaction is exported from the cell by the antiporter in exchange for a molecule of the decarboxylase’s basic amino acid substrate, thus constituting a cycle of decarboxylation, product export, and substrate import [130]. One region of the S. meliloti symbiotic plasmid (pSyma) encodes a putative Put/Orn antiporter next to the odc1 gene and located close by are a possible agmatine/Arg antiporter and putative Adc [36]. S. meliloti encodes two other possible basic amino acid/PA antiporters in other regions of pSyma, one of which was shown to be essential for symbiosis by Tn-seq analysis [62]. The function of the decarboxylase and antiporter genes on the symbiotic plasmid suggests a role in symbiosis, most logically in acid stress resistance. Nevertheless, an S. meliloti odc1 mutant grew like wild type at pH 5.5, suggesting that Odc1 is not required for resistance to this relatively mild acidity [60]. Further testing of this mutant and of the other pSyma genes potentially involved in acid resistance is being done in our laboratory.
The expression of a Spd/Put ABC transporter was reduced under acid growth conditions in Rhizobium freirei PRF 81, which belongs to the “R. tropici group” of bean nodulating rhizobia [129]. In S. meliloti 1021, potD encodes the substrate binding protein of a putative TRAP-type Put transporter and was progressively and permanently downregulated over the course of short-term growth at low pH [131]. In the case of acid-stressed S. meliloti, decreased import of external Put from the environment could prevent its interfering with endogenous Put biosynthesis and the proton consumption that is accomplished as part of the acid resistance mechanism described above. However, the R. freirei PRF81 genome sequence does not appear to contain basic amino acid/PA antiporters nor odc homologs of those on the S. meliloti pSym.
The growth of S. meliloti Rm8530 at pH 5.5 caused a several-fold decrease in the level of Put, while the amounts of Spd and HSpd were unaffected. An odc2 mutant that made very low levels of Put, HSpd, and Spd grew much more slowly than the wild type at pH 5.5. Growth of the mutant at pH 5.5 was completely restored by exogenous Spd and partially restored by Put or NSpd (HSpd was not tested due to lack of a commercially available product) [60]. These results suggest that Put (itself or as a precursor of Spd), NSpd, and Spd are able to lessen the impact of acid stress on growth and may be functionally interchangeable.
In S. meliloti 2011, grown at different pHs in chemostat cultures, intracellular Put levels increased at pHs above (7.4) or below (6.1) neutrality. However, transcriptomic and proteomic analyses did not show changes in genes for PA biosynthesis or transport [132]. The acid-induced decrease in Put in S. meliloti Rm8530 batch cultures and its increase in strain 2011 chemostat cultures are probably due to differences in the experimental approaches and the fact that strains 2011 and 1021 (the latter is the parent strain of Rm8530) do show important phenotypic differences despite being derived from S. meliloti strain SU47 [133].
Temperature Stress
Global warming is increasing heat stress on PGPR and plants. High temperatures cause protein misfolding and inactivation [98,121]. Heat shock in R. etli CE3 increased the expression of genes encoding chaperones and proteases that aid in protein folding and in degrading misfolded proteins, respectively. These responses do not occur in R. etli exposed to salt shock [126].
Little has been reported on PAs as related to heat stress resistance in PGPR. Growing S. meliloti Rm8530 at 37 °C rather than its “optimal” temperature of 30 °C significantly increased both its growth rate and its intracellular content of Spd. A Rm8530 odc2 unable to synthesize normal levels of Spd, Put, or HSpd grew at about half the rate of the wild type at 37 °C, while the mutant genetically complemented with the odc2 gene grew like wild type. This suggests that Spd or other PAs are important for the growth of S. meliloti under heat stress (V. Becerra-Rivera, unpublished).
Cold stress creates problems for bacteria by increasing membrane rigidity and decreasing the rate of enzyme reactions [98,121]. The general response of Mesorhizobium sp. strain N33 to cold shock was to adjust gene expression to achieve growth reduction or cessation. In addition, strain N33 under cold stress upregulated expression of the gene for the ATPase subunit of an ABC-type PA transporter, possibly indicating increased transport of PAs as protective agents [134].
3.2.3. The Importance of Polyamines in Motility and Biofilm Formation
PAs present in the environment markedly affect chemotaxis, EPS production, biofilm formation, and motility in some PGPR [36,38]. Chemotaxis to root exudates is an essential first step for the establishment of PGPR–plant interactions: Bacteria that arrive first at the root will have the widest choice of nutrients and hence a growth advantage [24]. Put, Cad, and Spd are chemoattractants for Pseudomonas putida KT2440, which can utilize these PAs as carbon or nitrogen sources for growth in cultures. All 3 of these PAs bind to the McpU ligand binding domain, while Spm, Orn, GABA, or proteinogenic amino acids do not. A P. putida KT2440 mcpU mutant was severely compromised in its ability to colonize corn roots [44]. The McpU homolog in S. meliloti allows chemotaxis towards proline, which is found in alfalfa seed exudates, but it is not known if it is also a chemoreceptor for PAs [135].
The second messenger bis-(3′-5′) cyclic diguanosine monophosphate (c-di-GMP) regulates motility and biofilm formation in many bacteria. In S. meliloti, the PA signal detection/transduction system comprised of the NspS and MbaA proteins affects biofilm, EPS production, and motility in response to specific exogenous PAs. The NspS-MbaA-dependent phenotypic responses to PAs differ depending on whether the quorum sensing (QS) system is functional. This is consistent with the multiple interacting regulatory circuits, including QS, that regulate biofilm formation, EPS production and motility in S. meliloti. We speculate that the NspS-MbaA system modulates these symbiotically important phenotypes in S. meliloti in response to host plant-produced PAs [136].
Swimming motility in an S. meliloti Rm8530 hss null mutant was reduced by one-third relative to the wild type, while a cansdh mutant was unaffected in swimming. Swimming by a hss cansdh double mutant was reduced by nearly 60% compared to the wild type (V. Becerra-Rivera and M. Dunn, unpublished results). The more pronounced phenotype of the double mutant probably results from its inability to produce either of the partially compensatory triamines Spd and HSpd.
Inactivation of the hss gene in R. etli CNPAF512 abolished its ability to swarm, indicating that HSpd is essential for this type of motility [137]. Like R. etli strain CFN42 [36], strain CNPAF512 probably lacks the ability to produce Spd which might compensate for the lack of HSpd in the mutant.
4. A Brief Review of the Roles of Polyamines in Plants
While higher plants have both the Odc and Adc pathways for Put production [33,138], Adc is usually the sole or more important pathway [139]. Put is a precursor of Spd and Spm in all plants and of TSpm in some plants. These PAs are synthesized by Spd, Spm, and TSpm synthases, respectively, as described in Section 3.1 (Figure 2). The triamine NSpd is found in non-vascular plants and some higher plants [140,141,142]. The production of Cad by Ldc is restricted to Leguminosae and Solanaceae [143].
Broadly generalizing, Spd is required in plants for growth, and Spm and Put are important in stress resistance [139]. Manipulating the PA content in plants by exogenous application or metabolic engineering has shown that PAs are involved in embryogenesis, fruit development and maturation, flower and chloroplast development, organogenesis, and senescence and stress responses. TSpm, for example, controls stem elongation in A. thaliana by promoting the expression of upstream open reading frames (uORFs) that control translation of genes involved in xylem differentiation, which affects stem elongation [79,144]. In fact, some genes required for plant PA biosynthesis are regulated by uORFs, probably in response to specific PAs [145]. Spm and its degradation products trigger plant defense reactions to pathogens (for reviews, see [39,40,146,147].
In legumes, L/Odcs are the starting points for the production of Cad and HSpd for making alkaloids that provide chemical defense against herbivorous insects [143,148,149]. In the genus Crotalaria (“rattlepods”, Fabaceae), HSpd is incorporated into the structure of the pyrrolizidine alkaloid monocrotaline. Plant-encoded Hss produces HSpd for monocrotaline synthesis, although the plants need to be nodulated by Bradyrhizobium spp. to produce the alkaloid. The restriction of alkaloid synthesis to nodulated plants might ensure their sufficiently high nitrogen status for alkaloid production [149].
PAs protect plants from abiotic stresses by acting as molecular chaperones to protect macromolecules from physical and chemical damage. PAs in plants also stabilize plasma membranes and significantly affect the activity of various ion transporters, both of which are important for growth and stress resistance [150,151]. A recent study found that treatments affecting the formation of conjugated forms of Spd and Spm correlated with drought resistance, root plasma membrane stability, and higher H+-ATPase activity [152]. Pas also interact with phytohormones to promote stress resistance (Section 5.2.2).
PA catabolism regulates plant stress response pathways by producing nitric oxide (NO), H2O2, and GABA, all of which also have important signaling functions in plant–microbe interactions [41]. These signals arise as products of PA catabolism by diamine and PA oxidases [153]. Plant defense responses triggered by different biotic and abiotic stresses increase ROS levels. These ROS also serve as chemical signals during the interaction of legumes and rhizobia and are required for the development of an efficient symbiosis [99,102]. PAs are also effective scavengers of ROS, increase the activity of antioxidant enzymes, and trigger ROS-defensive signaling pathways (for reviews, see [105,122,146,154,155]). Thus, in addition to being the agents of oxidative stress and associated damage, ROS in plants act as key physiological signaling and regulatory molecules. In terms of PA metabolism, a balance must be struck between the protective effects of PAs against ROS and ROS production by PA catabolism [104,146,156].
Analyses of abiotic stress-tolerant legume genotypes have revealed changes in their PA content in comparison to less tolerant lines. For example, a drought-tolerant line of Lupinus luteus (yellow lupin) had a high Spd level and was characterized by higher plant and seed biomass than non-tolerant lines under drought stress [157]. Salt-stress tolerant genotypes of M. truncatula [158] and G. max [159] had high levels of Spm and Put, and Spd and Spm, respectively, when grown under salt stress. In both cases, PA catabolism by PAO was diminished, and in G. max, antioxidant enzyme activities increased [159]. Because many different kinds of abiotic stress ultimately trigger oxidative stress, the ability of PAs to increase a plant’s resistance to diverse stress conditions is, in part, a measure of their ability to resist oxidative stress.
PAs also aid in abiotic stress resistance in plants by increasing the production of GABA, which moderates the negative effects of abiotic stress by affecting osmoregulation, antioxidant defense, and molecular signaling [160,161]. GABA synthesis in plants results from the degradation of PAs by diamine oxidase (DAO, EC 1.4.3.6) and by the action of glutamate decarboxylase (GDC, EC 4.1.1.15). Salt stress imposed on soybean seedlings caused a rapid increase in DAO activity and dao gene transcription, and GABA content, with a concomitant decrease in Put and Spd levels. The inhibition of DAO with the specific inhibitor aminoguanidine reduced seedling GABA content by nearly 40%, suggesting that this proportion of total GABA is produced by DAO [162]. Fang and co-workers [163] showed that exogenous Spd alleviated the inhibition of soybean seedling growth caused by salt stress. Spd treatment was accompanied by an increase in GABA content and a reduction in malondialdehyde (an indicator of membrane oxidative damage) and H2O2 levels, concomitant with an increase in ROS-detoxifying enzyme activities [163]. Similarly, exogenous Put prevented the growth inhibition of soybean seedlings under salt stress and increased superoxide dismutase, catalase, and peroxidase activities to near the levels found in unstressed plants. Exogenous Put reduced the levels of H2O2 and superoxide as well as that of malondialdehyde [164].
5. Polyamines in PGPR–Plant Interactions
We have seen that endogenous PAs or those taken from the environment can affect diazotrophs and plants living separately, but what effects do they have during the interaction of plants with PGPR? The fact that PAs modulate the interaction of plant pathogens with their hosts is well documented, to the extent that phytopathogens are known to alter plant PA metabolism to their own benefit [39,165,166,167,168]. The roles of PAs in mutualistic plant–PGPR interactions are much less clear, and most work done in the area has been with rhizobia and legumes [37,39,105]. In this section, we examine how PA exchange between symbiotic partners affects developmental processes and stress resistance in both plants and PGPR.
5.1. Polyamines Found in Root Nodules
The free PAs commonly found in legume root nodules include Put, HSpd, Spd, Cad, and Spm. Put in nodules is synthesized by both symbionts while Spd, Cad, and Spm are usually made by the plant partner [36,37,120]. In contrast, HSpd in nodules is contributed by the microsymbiont, although some legumes are able to synthesize it using a deoxyhypusine synthase-like Hss that transfers an aminobutyl group from Spd to Put to produce HSpd [33,140,143,169].
In nodules formed by R. legminosarum bv. viciae 3841 on pea mutants unable to support nitrogen fixation, plant odc gene expression was highly induced in the ineffective nodules relative to the effective nodules formed on the wild-type pea line. The transcriptional expression of adc was similar in both effective and ineffective nodules. The higher odc expression in ineffective nodules indicates that it may function as part of a defense response against the formation of non-functional nodules, while the Adc serves for the housekeeping synthesis of Put [170] (Section 4).
The uncommon PA 4-aminobutylcadaverine (AbCad) is found in nodules of adzuki bean and common bean formed by B. japonicum A1017 and R. tropici CIAT899, respectively. The synthesis of AbCad in nodules on these species depends on their containing high levels of Cad, to which the bacteroid Hss adds an aminobutyl group from Put to form AbCad [82,123,171,172]. Bacteroids of an R. tropici CIAT899 hss mutant failed to produce HSpd and AbCad in bean nodules, while the wild type made high amounts of each [123]. These findings confirm the bacterial origin of HSpd in bean nodules and the role of Hss in producing AbCad.
Bacteroids in nodules are notable for their high HSpd content [123,172,173,174]. Nodules from soybean, faba bean, and kidney bean contain roughly equal amounts of HSpd and Spd, while those of cowpea, mung bean, siratro, and Sesbania spp. have high HSpd to Spd ratios. The fact that different PA patterns occur in nodules of different species of plants able to host the same Rhizobium or Bradyrhizobium species indicates that the PA content in nodules is mostly controlled by the plant host and not the microsymbiont [174]. It would be interesting to further explore this using a host plant that is nodulated by rhizobia that differ in the types of PAs they produce.
Unlike legumes, the non-legume shrub Alnus spp. has extremely low quantities of PAs in its nodules [174]. Nodules from different Alnus species contained Spd and occasionally Cad and Put but lacked HSpd [84,174]. The low levels of PAs in these nodules may relate to their high content of lignified tissue, which in senescent nodules correlates with a low PA content [84,174].
A R. leguminosarum bv. viciae strain was modified by replacing its native symbiotic plasmid with that from an R. leguminosarum bv. phaseoli strain, thus allowing it to form indeterminate nodules on pea while the native bv. viciae strain formed determinate nodules on bean. Transcriptome comparisons of bacteroids from both hosts showed that genes for Put transporter components, a L/Odc and Hss were upregulated in bacteroids isolated from determinate nodules but not in bacteroids from indeterminate nodules [175]. The upregulation of the Put transporter genes suggests the presence of sufficient quantities of Put around the bacteroids in determinate nodules to induce the transporter [90], while upregulation of the two genes for PA biosynthetic enzymes suggests that more PAs are produced by the bacteroids in these nodules than in the indeterminate ones [125].
Another example of host-dependent expression of PA biosynthesis genes was found by proteome analysis of R. leguminosarum bv. viciae UPM791 bacteroids isolated from indeterminate nodules formed on pea and lentil. Those from pea showed much higher expression of putative Ldc, Odc, and DABA AT enzymes in pea relative to lentil [176]. Bacteroids of strain UPM791 from both pea and lentil contained Put, NSpd, Spd, and HSpd, while cells grown in culture contained only HSpd and Put. Unexpectedly, the deletion of gene encoding DABA AT did not alter the PA profile of bacteroids from either host or of cells grown in culture [177].
Nodules on the non-legume Parasponia andersonii formed by Bradyrhizobium spp. CP-283 contained similar levels and types of PAs as found in most legume nodules, predominantly Put and HSpd, with much lower levels of Spd, Cad, and Spm [174]. A comparison of the PAs in stem and root nodules on the non-legume Sesbania rostrata inoculated with A. caulinodans ORS571 showed they were similar in both types of nodules [174], despite their different morphologies and the much higher level of nitrogen fixation and metabolic activity in stem nodules [178].
5.2. Involvement of PAs in PGPR–Plant Interactions
5.2.1. Effects of PAs on Growth, Differentiation, and Metabolism of the Symbiotic Partner
Plants treated with exogenous PAs generally show increased growth, accelerated development, higher photosynthetic efficiency, and greater stress resistance [40,153,170,179]. Examples of how the manipulation of PA levels in plants affects their interaction with PGPR are summarized in Table 1, and some examples are further described here.
Put added to the nutrient solution of chickpea or vetch inoculated with indigenous or commercial inoculum strains of R. leguminosarum significantly increased plant nodule and shoot biomass, chlorophyll, and total nitrogen [180]. In the R. galegae-goat’s rue symbiosis, 10 or 50 µM Put included in the agar growth medium increased nodule numbers and biomass per plant, and nitrogen-fixing activity [181], possibly by increasing acid stress resistance and the root-binding ability of R. galegae [182]. Treatment of the G. orientalis plants with 100 µM Put or Spd reduced nodule numbers and nodule biomass per plant to below the levels of the untreated control. Spm at this concentration had little effect [181]. It was hypothesized that the reduction in nodulation caused by 100 µM Put was due to this concentration being inhibitory for the growth and root attachment ability of R. galegae [181].
Exogenous Put (0.1 mM) increased the number of nodules formed and acetylene reduction activity in pea inoculated with R. leguminosarum bv. viciae 3841, while 1 mM Put significantly decreased nitrogenase activity but not nodulation. Wild-type plants treated with 5 mM Put were very sparsely nodulated and fixed no nitrogen. However, nodulation and nitrogen fixation in transgenic plants with greatly reduced levels of DAO activity were not inhibited by 5 mM Put. Possibly, the high Put levels inhibit bacteroid growth or development via an oxidative burst from its catabolism by DAO. This would not occur at such high levels in transgenic plants with lower DAO activities [183].
Cucumber inoculation with A. baldaniorum Sp 245 (formerly A. brasilense Sp 245) increased Put and H2O2 levels and DAO activity and lowered the content of cell wall phenolics. The higher Put content may allow DAO to produce more H2O2 and other ROS that, along with the reduced concentrations of cell wall phenolics, promote growth of the roots [184]. In wheat, inoculation with A. baldaniorum Sp 245 reduced root superoxide levels by nearly one-third [185].
PAs may also control the growth of some diazotrophs during their interaction with plants. Species of Frankia in free life fix nitrogen in vesicles, which develop from the hyphae. Hyphal growth and integrity and vesicle formation were significantly inhibited by exogenous Spd or Spm used as a nitrogen source by Frankia grown in cultures [84].
In B. japonicum, the regrowth of bacteroids isolated from soybean nodules was inhibited by micromolar concentrations of Spd or Spm [186]. While Spm is absent or present at very low levels in soybean nodules, Spd accounts for about one-third of their total PA content [174,187], indicative of a concentration sufficient to retard bacteroid growth.
PAs likely participate in controlling nodule formation in the B. japonicum/soybean interaction apart from their direct effects on bacteroid growth. The leaves, roots, and nodules of the supernodulating soybean mutant En6500 had a strikingly lower Spd/Put ratio than the parental cv. Enrei. This suggested that there was less conversion of Put to Spd in the supernodulating line. Foliar application of the SAMdc inhibitor MDL74038 significantly reduced Spd levels in the parental cv. Enrei and doubled the number of nodules formed. Consistent with this, the foliar application of Spd or Spm drastically reduced the number of nodules formed by the supernodulating En6500 mutant. This suggests that the total amount or ratio of different PAs modulate the number of nodules formed on soybean, in conjunction with phytohormones called brassinosteroids. The hypothesis from this work is that low levels of brassinosteroids in the supernodulating line suppress Spd synthesis and lead to excessive nodulation [187,188]. Our current understanding of autoregulation of nodulation is incomplete, and the process is controlled by a great number of plant signal receptors that respond to nitrogen availability, phytohormones, and chemical signals generated during the symbiosis. While PAs have only been indirectly implicated in autoregulation of nodule numbers in soybean (see above), they could also conceivably participate in modulating autoregulation control points like mRNA stability and translation efficiency [189].
Gene mutagenesis and Tn-Seq experiments showed that two arginase homologs and the Orn decarboxylase homologs Odc1 and Odc2 were required for an efficient symbiosis between S. meliloti and M. truncatula [62]. The inactivation of Odc2 in S. meliloti Rm8530 also decreases its symbiotic efficiency on M. sativa [115]. The Tn-Seq experiments also showed that the S. meliloti genes encoding two Arg transporters, four Put ABC transporters, Hss and DABA AT were required for an efficient symbiosis with M. truncatula [36,62]. The symbiotic requirement for Hss and DABA AT indicates a role for endogenously produced HSpd and NSpd in bacteroids, and the increased transporter activity might serve to import plant-produced PAs. Tn-Seq analysis of R. leguminosarum bv. viciae in symbiosis with pea plant revealed that the hss gene was essential in both undifferentiated and bacteroid forms of the microsymbiont isolated from nodules but dispensable for growth in the rhizosphere or for root colonization [190].
The PGPR B. subtilis OKB105 increased tobacco plant growth by exporting endogenously synthesized Spd, which increased the expression of genes for expansions and decreased expression of the ACO1 gene for ethylene biosynthesis. Lower levels of ethylene favor plant growth. Strain OKB105 null mutants in speB (agmatinase) or a transporter required for Spd export (yecA) failed to stimulate plant growth. Exogenous Spd or genetically complemented mutants were effective in promoting plant growth [191]. Bacillus megaterium BOFC15 secretes Spd in vitro and, when inoculated onto Arabidopsis, increased plant Spd, Spm and abscisic acid (ABA) levels and drought stress resistance. The hypothesis from these results is that Spd secreted by B. megaterium caused an increase in ABA levels in the plant, and this imparts higher drought tolerance [192].
A PA ABC transporter is required for the normal development of nitrogen-fixing heterocysts in Anabaena sp. PCC 7120. The transporter may be required to maintain PA homeostasis or for transferring HSpd, which is required for heterocyst development, from vegetative cells to the heterocysts [63].
5.2.2. Polyamines in PGPR–Plant Associations under Abiotic Stress
The ability of PGPR to enhance the abiotic stress resistance of plants has been amply demonstrated and was recently reviewed [18,27,154]. In general, PGPR potentiate host responses that allow them to better tolerate abiotic and biotic stresses and promote better growth in general, which itself increases the stress-fighting ability of plants.
The PGPR Stenotrophomonas rhizophila DSM14405 grown in the presence of oilseed rape root exudates had several-fold higher transcription of the mdtI and mdtJ genes, which encode a Spd transport system [193]. In E. coli, MdtIJ activity is required to prevent Spd toxicity in mutants unable to acetylate and thus inactivate “excess” endogenous Spd [194]. Spd export from S. rhizophila via MdtIJ was not experimentally demonstrated but was hypothesized to be involved in this PGPR’s ability to increase plant growth and stress resistance [193]. S. rhizophila DSM14405 co-inoculated with B. japonicum onto soybean increases the symbiotic effectiveness of the nitrogen-fixing interaction under salt stress. The export of Spd or glucosylglycerol (an osmoprotectant) by DSM14405 is a possible mechanism for the synergistic effects of co-inoculation with the two species [195].
The experimentally supported metabolic model proposed by Christen and co-workers asserts that Arg and succinate produced by M. truncatula are actively transported into S. meliloti bacteroids and co-metabolized to generate energy and reduce power for nitrogen fixation [62]. The catabolism of Arg to Put by the Odc and Adc/agmatine routes releases ammonium and consumes protons, which is hypothesized to protect the bacteroids against acid conditions normally present in the symbiosomes [62,99,102]. As mentioned above (Section 3.2.2 and Section 5.2.1), and consistent with this role for PAs in acid stress resistance, gene mutagenesis, and Tn-Seq experiments showed that two arginase homologs and the Orn decarboxylase homologs odc1 and odc2 were required for an efficient symbiosis between S. meliloti and M. truncatula. [62].
In Galega orientalis [181] and M. truncatula [196], treatments with exogenous PAs increased nodulation and nitrogen fixation, as well as the biomass of shoots, roots, and nodules under normal conditions and salt stress. The conclusion from these studies is that changes in the physiology or metabolism of the nodule are not caused by PAs at that site but result from secondary signals generated by PA action in the shoots. PAs have been shown to have regulatory interactions with hormones like ethylene, ABA, salicylic acid, and brassinosteroids [26,161,196,197]. Exogenous Spd or Spm in the nutrient solution supplied to M. truncatula inoculated with S. meliloti 1021 increased the expression of brassinosteroid synthesis genes and increased symbiotic efficiency under salt stress [196]. Treatment of salt-stressed M. sativa inoculated with S. meliloti GR4 without or with ABA pre-treatment increased Spd and Spm levels in nodules, increased antioxidant enzyme activities, and improved their nitrogen fixation capacity [198].
The positive effect of PAs on both plant and PGPR abiotic stress resistance is one reason for the generally beneficial effects of PGPR on plant growth. In addition, the development of Rhizobium-legume interactions may require signaling pathways or other metabolic changes that result from responses to different abiotic stresses that occur during the normal course of symbiosis. Both the micro- and macrosymbiont participate in these stress-induced responses [102], some of which are known to be modulated by PAs.
Bacteroids of wild type R. tropici CIAT899 isolated from bean plants grown under salt stress had higher levels of HSpd than bacteroids from unstressed plants. A CIAT899 hss null mutant grew more slowly than the wild type under salt stress in vitro. Plants inoculated with the hss mutant had significantly lower nodule fresh weights than those of plants inoculated with the wild type under both control and salt stress conditions [123]. The beneficial effect of the R. tropici hss on nodule mass and salt resistance agree with their importance in the symbioses of S. meliloti and R. leguminosarum with their respective hosts described in Section 5.2.1.
In the A. brasilense commercial inoculant strains Cd and Az39, Cad exported from the cells was implicated in plant growth promotion and increased osmotic stress resistance [75,76]. Exogenous Cad applied to rice seedlings at a concentration similar to that found in A. brasilense culture supernatants promoted root and shoot growth, similar to the effect of treating seedlings with A. brasilense Az39. The growth-promoting effects of exogenous Cad and inoculation with strain Az39 also occurs in plants under osmotic stress [75]. However, for other strains of A. brasilense inoculated onto wheat or Arabidopsis, auxin production by the bacteria and inhibition of plant ethylene biosynthesis appears to be responsible for plant growth promotion [199,200].
6. Conclusions and Perspectives
In PGPRs and plants, endogenous PA biosynthesis and the uptake of PAs from the environment are important in determining intracellular PA levels and, accordingly, their physiological effects [201]. A summary of the general effects of PAs on plant–PGPR interactions is shown in Figure 3, although it is conceptually and experimentally difficult to separate the effects of PAs on symbiotic partners separately versus effects that occur only during their interaction. PAs excreted into the environment can also serve as signal molecules in PGPR–plant interactions, although this aspect has not been explored in great depth. The possible or proven mechanisms by which PAs exported by PGPR or nodule-forming rhizobia affect plant growth and stress resistance include stabilizing plant plasma membranes, affecting the activity of plant ion transporters, decreasing oxidative damage to cells by increasing antioxidant defenses or by interacting with phytohormones to promote stress resistance (Section 4). PAs made by legumes also participate in limiting bacteroid proliferation and regulation of nodule numbers (Section 5.2.1). In PGPR, the perception of environmental PAs is important in promoting chemotaxis, while both endogenous and exogenous PAs affect motility, biofilm formation, and EPS production (Section 3.2.3). Because PAs in either plant or PGPR may act as signals, their ability to trigger physiological or morphological changes or increase stress resistance may not require remarkable changes in their levels, a potentially misplaced criterium for their importance in many studies.
PA biosynthesis or availability of PGPR in soil or in their interaction with plants are important as modulators of stress resistance. In soil, the ability to survive for long periods is important in maintaining PGPR populations and giving competitive advantages in the rhizosphere to specific PGPR species [202], and PAs have an important role in this.
The efficient design of effective bioinoculants containing synthetic communities of PGPR requires detailed knowledge of the physiological and genetic traits of the organisms and their interaction with one another and the environment, including different plant hosts, and how to translate laboratory studies done on a few model organisms to a broader range of plants in the field [203]. Could manipulating the kinds or quantities of PAs exported by PGPR enhance their ability to increase the growth of specific crops? As described in Section 3.1, PGPR produce a range of different PAs, although there is considerable overlap in the PA profiles within some species and even in some genera and classes (Section 3.1). It is not known if differences in the types of PAs produced by PGPR function as determinants of host specificity: This would depend largely on the Pas excreted by the PGPR and how they are perceived by different plants. We suspect that Pas produced by PGPR are not major determinants of host specificity since many plants interact promiscuously with PGPR or rhizobia that produce different types of Pas, while specific wide host range PGPR can interact with many different plant hosts [43,50,204].
Table 1Effects of polyamines on PGPR in free life or in association with plants.
| PGPR | Experimental System or Treatment | Results | Interpretation | References |
|---|---|---|---|---|
| Anabaena sp. | Diazotrophic growth of speA (Adc), speB (agmatinase), or speY (hss synthase homolog) null mutants | Reduced diazotrophic growth. speA mutant cannot form heterocysts | HSpd is needed for heterocyst differentiation as a cell wall component, or its absence affects nitrogen metabolism and/or signaling processes | [63] |
| Azospirillum |
Inoculated on cucumber | Inoculation correlated with increased Put and H2O2 levels, diamine oxidase (DAO) activity, and reduced cell wall phenolics content | Higher Put content allows DAO to make more H2O2 and other reactive oxygen species (ROS). This and the lower concentration of phenolics promote root growth | [184] |
| Azospirillum |
Rice inoculated with strain Az39 or treated with exogenous Cad, with or without osmotic stress (mannitol) | Significantly increased osmotic stress resistance and abscisic acid (ABA) production for both treatments | Cad excreted by strain Az39 promoted growth and osmotic stress resistance in rice | [75] |
| Bacillus megaterium BOFC15 | Inoculated on Arabidopsis | BOFC15 secretes Spd in vitro. When inoculated on Arabidopsis, it increased plant Spd, Spm, and ABA levels, and drought stress resistance | Spd secreted by B. megaterium increases plant ABA levels, thus increasing drought tolerance | [192] |
| Bacillus subtilis OKB105 | Inoculated on tobacco | The strain exports Spd, causing increased expression of genes for expansins and decreased expression of the ACO1 gene for ethylene biosynthesis. Spd biosynthesis mutants do not have this effect | Spd exported by strain OKB105 results in lower ethylene content in the plants and favors growth | [191] |
| Bradyrhizobium diazoefficiens 110spc4, Rhodopseudomonas palustris CGA009, Rhizobium tropici CIAT899T | Growth in cultures following Tn-seq mutagenesis (B. j., R. p.) or hss inactivation (R. t.) | Few hss mutants recovered (Tn-seq mutagenesis), or slower growth of hss mutant | HSpd important for growth | [94,123] |
| Bradyrhizobium japonicum 138NR | Growth in culture with exogenous PAs | Regrowth of bacteroids isolated from soybean nodules, but not of culture-grown cells, is prevented by 0.1 mM Spd or Spm, but not by Put | Spd and Spm inhibit proliferation of bacteroids in nodules | [186] |
| Bradyrhizobium japonicum USDA 110 (now B. diazoefficiens USDA 110T [205]) | Inoculation on soybean super-nodulating and parent cultivar, without or with foliar application of Spd or Spm, or the SAMdc inhibitor MDL74038 | PA treatment decreased the number of nodules formed on the super-nodulating cv. and decreased root growth. Application of MDH74038 to the parent cv. increased nodule number | PAs function as a systemic regulator of nodulation by affecting the plant rather than the microsymbiont | [187,188] |
| Frankia spp., various strains | Cultures grown with exogenous Spd or Spm | Formation of vesicles (where nitrogenase is located) is inhibited, hyphal degradation occurs | PAs inhibit growth and vesicle formation | [84] |
| Pseudomonas putida KT2440 | Inoculated on corn | Ability of a mcpU PA chemotaxis receptor mutant to colonize plants was greatly reduced | Chemotaxis towards PAs favors root colonization | [44] |
| Rhizobium etli CNPAF512 | hss null mutant | Mutant unable to swarm | HSpd required for swarming motility | [137] |
| Rhizobium galegae HAMBI 540T | Inoculated on goat’s rue, plants grown without or with 10 or 50 µM exogenous Put or Spd | Plant acid stress resistance and binding of R. galegae to roots enhanced by PA treatment | The exogenous PAs increase root attachment and colonization | [181] |
| Rhizobium galegae HAMBI 540T | Inoculated on goat’s rue, plants grown without or with 100 µM exogenous Put, Spd or Spm | Nodule number and biomass per plant with Put or Spd treatment lower than those of untreated control. Spm had little effect | High concentration of Put or Spd reduces R. galegae growth and root attachment ability | [181] |
| Rhizobium leguminosarum indigenous or commercial inoculum strains | Inoculation on chickpea or vetch with or without single-dose Put treatment | Put treatment significantly increased nodule and shoot biomass, chlorophyll, and total nitrogen | Stimulation of plant growth with Put treatment not dependent on R. leguminosarum strain. Mechanism of growth stimulation unknown | [180] |
| Rhizobium leguminosarum bv. viciae 3841 | Growth in cultures without or with 2,4-dichlorophenoxyacetic acid | 2,4-dichlorophenoxyacetic acid provokes oxidative stress | Intracellular levels of Put and Cad increase as a protective response against oxidative stress | [117] |
| Rhizobium leguminosarum bv. viciae 3841 | Inoculated on pea plants grown with 0, 0.1, 0.5, 1, or 5 mM exogenous Put | Exogenous Put at 0.1 mM gives a slight increase in nodule number and a 50% increase in nitrogenase activity per plant. Put at ≥1 mM decrease these parameters relative to control without Put | Endogenous Put at too high a level is inhibitory to nodule function | [183] |
| Rhizobium leguminosarum bv. viciae 3841 | Inoculated on transgenic pea lines with normal or severely reduced DAO activity, under control conditions or with exogenous Put | Under control conditions, no correlation between nodule number or nitrogen fixation and DAO activity, but with exogenous Put, small increase in nodule number and big increase in nitrogen fixation. | DAO activity reduces inhibitory levels of PAs in nodules | [183] |
| Rhizobium leguminosarum bv. viciae 3841 | Growth in cultures without or with the Odc inhibitor DFMO | Inhibitor reduces PA biosynthesis and growth: Exogenous Put or Spd largely overcomes inhibition | PA biosynthesis required for wild type growth | [82,83] |
| Rhizobium leguminosarum bv. Viciae with native viciae pSym or with introduced pSym from bv. Phaseoli | Strains with bv. Viciae and bv. Phaseoli pSyms form determinate and indeterminate nodules, respectively, on bean | Put transporter components, l/odc and hss, upregulated in bacteroids from determinate but not indeterminate nodules | PA transport and endogenous synthesis higher in bacteroids from determinate versus indeterminate nodules | [175] |
| Rhizobium tropici CIAT899T | Growth of wild type and hss mutant under salt stress | Slower growth of the mutant partially restored by 0.1 mM exogenous HSpd | HSpd important for salt stress resistance | [123] |
| Sinorhizobium meliloti (currently Ensifer meliloti [206]) strains Rm8530 and 1021 | Phenotypic characterization of wild types and nspS null mutants | Altered biofilm formation, EPS production, and motility in presence of selected PAs, dependent on functionality of quorum sensing system | Exogenous PAs affect symbiotically relevant phenotypes in S. meliloti | [136] |
| Sinorhizobium meliloti (currently Ensifer meliloti [206]) Rm8530 | Swimming motility of hss cansdh double mutant | Swimming significantly reduced versus wild type and hss and cansdh single mutants | Spd, HSpd, and/or NSpd are important for swimming motility | V. Becerra and M. Dunn, unpublished. |
| Sinorhizobium meliloti (currently Ensifer meliloti [206]) 1021 | Wild type and tolC (outer membrane protein) mutant grown in cultures | tolC mutation provokes oxidative stress and higher expression of odc1 and the Arg/Lys/Orn decarboxylase sma0682. | Cells of mutant attempt to counter oxidative stress by synthesizing more PAs | [114] |
| Sinorhizobium meliloti (currently Ensifer meliloti [206]) 1021 | Inoculated on M. truncatula without or with salt stress and 0.l mM exogenous Spd or Spm | Treatments with Spd or Spm increased the expression of brassinosteroid synthesis genes and lowered oxidative stress provoked by high osmolarity | Cross-talk of PAs and the anti-stress phytoharmone brassinosteroid increase resistance to abiotic stress | [196] |
| Sinorhizobium meliloti (currently Ensifer meliloti [206]) GR4 | In combination with M. sativa under salt stress without or with ABA pre-treatment | ABA pre-treatment increased Spd and Spm levels in nodules, increased antioxidant enzyme activities, and improved nitrogen fixation | ABA induces nodule antioxidant defenses mediated by Spd and Spm and other non-enzymatic antioxidants | [198] |
| Sinorhizobium meliloti (currently Ensifer meliloti [206]) Rm8530 | Cultures with exogenous 1 mM Put, Spd or NSpd | Growth rate reduced up to 20% vs. control cultures | PAs moderately toxic at this concentration | [60] |
| Sinorhizobium meliloti (currently Ensifer meliloti [206]), various strains | Growth in culture or in symbiosis with M. sativa or M. truncatula | Sharply decreased intracellular PA content in odc2 mutant or when Odc activity is inhibited with DFMO | PA biosynthesis required for wild type growth, (especially under abiotic stress) and full symbiotic efficiency. Chemical complementation with PAs or PA biosynthesis genes largely restores wild-type phenotypes | [60,62,82,83,115,207,208] |
| Stenotrophomonas rhizophila DSM14405T | Co-inoculated with B. japonicum on soybean | Synergistic effect that increases the symbiotic effectiveness of the nitrogen-fixing interaction under salt stress | Export of Spd or glucosylglycerol (an osmoprotectant) by DSM14405 may participate in plant growth promotion | [195] |
| Stenotrophomonas rhizophila DSM14405T | Growth with oilseed rape root exudates | Several-fold higher transcription of mdtIJ genes encoding a probable Spd export system | Spd exported by MdtIJ could promote plant growth and stress resistance | [193] |
| Synechocystis sp. PCC 6803 | Growth under salt (NaCl) or osmotic (sorbitol) stress | Increased levels of PAs with osmotic shock, Adc mRNA stability greater under both stresses | PAs may balance ionic charges or increase Adc mRNA stability | [128] |
V.A.B.-R. drafted the initial manuscript. M.F.D. and V.A.B.-R. edited and modified subsequent versions. All authors have read and agreed to the published version of the manuscript.
No new data were created or analyzed in this review.
The authors declare no conflict of interest.
Footnotes
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.
Enlarge this image.Figure 1. The rhizosphere can be divided into different parts. The innermost zone is known as the endorhizosphere, which encompasses both the interior of root cells and the apoplastic (extracellular) fluid-filled spaces surrounding them. Adjacent to the endorhizosphere is the rhizoplane, consisting of the root epidermis and the mucilage layer. Moving further outward, we encounter the ectorhizosphere, extending from the rhizoplane into the surrounding bulk soil. The accompanying figure was sourced from the Nature Education Knowledge Project (https://www.nature.com/scitable/knowledge/library/the-rhizosphere-roots-soil-and-67500617/, accessed on 9 March 2023).
Enlarge this image.Figure 2. Polyamine biosynthesis pathways. Red arrows denote the canonical pathways present in plants and some bacteria, blue arrows indicate additional or alternative pathways present in some bacteria. Abbreviations not defined in the figure: MTA, 5′-methylthioadenosine; SAM, S-adenosyl methionine. The figure was made with Microsoft Powerpoint for Mac, version 16.74.
Enlarge this image.Figure 3. Processes influenced by polyamines in plant–PGPR interactions, with reference to relevant sections of this review. Clockwise from left: PAs in both plants and PGPR increase resistance to biotic and abiotic stress (Section 3.2.2, Section 4 and Section 5.2.2). Both endogenous PAs and those present in the environment influence PGPR growth, motility, and possibly chemotaxis (Section 3.2.1, Section 3.2.3 and Section 5.2.1). PAs are important or essential for root colonization and/or biofilm formation (Section 3.2.3). Nodule formation and symbiotic efficiency are significantly affected by PAs in both legumes and rhizobia (Section 5.2). The combined effects of PAs on these processes are an important factor in plant growth promotion. Created with BioRender.com.
References
1. Backer, R.; Rokem, J.S.; Ilangumaran, G.; Lamont, J.; Praslickova, D.; Ricci, E.; Subramanian, S.; Smith, D.L. Plant growth-promoting rhizobacteria: Context, mechanisms of action, and roadmap to commercialization of biostimulants for sustainable agriculture. Front. Plant Sci.; 2018; 9, 14763.
2. Pankievicz, V.C.S.; Irving, T.B.; Maia, L.G.S.; Ané, J.-M. Are we there yet? The long walk towards the development of efficient symbiotic associations between nitrogen-fixing bacteria and non-leguminous crops. BMC Biol.; 2019; 17, 99.
3. Villalobos-López, M.A.; Arroyo-Becerra, A.; Quintero-Jiménez, A.; Iturriaga, G. Biotechnological advances to improve abiotic stress tolerance in crops. Int. J. Mol. Sci.; 2022; 23, 12053.
4. Fagorzi, C.; Passeri, I.; Cangioliu, L.; Vaccaro, F.; Mangoni, A. When biodiversity preservation meets biotechnology: The challenge of developing synthetic microbiota for resilient sustainable crop production. J. Sustain. Agric. Environ.; 2023; 2, pp. 5-15. [DOI: https://dx.doi.org/10.1002/sae2.12038]
5. Afzal, I.; Shinwari, Z.K.; Sikandar, S.; Shahzad, S. Plant beneficial endophytic bacteria: Mechanisms, diversity, host range and genetic determinants. Microbiol. Res.; 2019; 221, pp. 36-49. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30825940]
6. Aloo, B.N.; Tripathi, V.; Makumba, B.A.; Mbega, E.R. Plant growth-promoting rhizobacterial biofertilizers for crop production: The past, present, and future. Front. Plant Sci.; 2022; 13, 1002448. [DOI: https://dx.doi.org/10.3389/fpls.2022.1002448]
7. Banerjee, S.; van der Heijden, M.G.A. Soil microbiomes and one health. Nat. Microbiol. Rev.; 2023; 21, pp. 6-20.
8. Chagas, F.O.; de Cassia Pessotti, R.; Caraballo-Rodriguez, A.M.; Pupo, M.T. Chemical signalling involved in plant-microbe interactions. Chem. Soc. Rev.; 2018; 47, pp. 1652-1704.
9. Jalmi, S.K.; Sinha, A.K. Ambiguities of PGPR-induced plant signaling and stress management. Front. Microbiol.; 2022; 13, 899563. [DOI: https://dx.doi.org/10.3389/fmicb.2022.899563]
10. Omae, N.; Tsuda, K. Plant-microbiota interactions in abiotic stress environments. Mol. Plant-Microbe Interact.; 2022; 35, pp. 511-526.
11. Soumare, A.; Diedhiou, A.G.; Thuita, M.; Hafidi, M.; Ouhdouch, Y.; Gopalakrishnan, S.; Kouisni, L. Exploiting biological nitrogen fixation: A route towards sustainable agriculture. Plants; 2020; 9, 1011. [DOI: https://dx.doi.org/10.3390/plants9081011]
12. Vacheron, J.; Desbrosses, G.; Bouffaud, M.-L.; Touraine, G.; Moënne-Loccoz, Y.; Muller, D.; Legendre, L.; Wisniewski-Dyé, F.; Prigent-Combaret, C. Plant growth-promoting rhizobacteria and root system functioning. Front. Plant Sci.; 2013; 4, 356. [DOI: https://dx.doi.org/10.3389/fpls.2013.00356] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24062756]
13. Mowafy, A.M.; El-ftouh, E.A.A.; Sdick, M.Y.; Abdelshafi, S.A.; Sallam, A.A.; Agha, M.S.; Aeid, W.R.A. Nitrogen-fixing archaea and sustainable agriculture. Nitrogen Fixing Bacteria: Sustainable Growth of Non-Legumes; Maheshwari, D.K.; Dobhal, R.; Dheeman, S. Springer Nature: Singapore, 2022; pp. 115-126.
14. Pal, G.; Kumar, K.; Verma, A.; Verma, S.K. From rhizosphere to endosphere: Bacterial-plant symbiosis and its impact on sus-tainable agriculture. Re-Visiting the Rhizosphere Eco-System for Agricultural Sustainability; Singh, U.B.; Rai, J.P.; Sharma, A.K. Springer Nature: Singapore, 2022; pp. 89-103.
15. Poole, P.; Ramachandran, V.; Terpolilli, J. Rhizobia: From saprophytes to endosymbionts. Nat. Rev. Microbiol.; 2018; 16, 291.
16. Puri, A.; Polyaminedda, K.P.; Chanway, C.P. Nitrogen-fixation by endophytic bacteria in agricultural crops: Recent advances. Nitrogen in Agriculture—Updates; Fahad, A.; Fahad, S. IntechOpen Limited: London, UK, 2018; pp. 73-94.
17. Steenhoudt, O.; Vanderleyden, J. Azospirillum, a free-living nitrogen-fixing bacterium closely associated with grasses: Genetic, biochemical and ecological aspects. FEMS Microbiol. Rev.; 2000; 24, pp. 487-506. [DOI: https://dx.doi.org/10.1111/j.1574-6976.2000.tb00552.x]
18. Yadav, A.J.; Verma, P.; Kaushik, R.; Dhaliwal, H.S.; Saxena, A.K. Archaea endowed with plant growth promoting attributes. EC Microbiol.; 2017; 8, pp. 294-298.
19. Hsu, S.-H.; Shen, M.-W.; Chen, J.-C.; Lur, H.-S.; Liu, C.-T. The photosynthetic bacterium Rhodopseudomonas palustris strain PS3 exerts plant growth-promoting effects by stimulating nitrogen uptake and elevating auxin levels in expanding leaves. Front. Plant Sci.; 2021; 12, 573634. [DOI: https://dx.doi.org/10.3389/fpls.2021.573634]
20. Smercina, D.N.; Evans, S.E.; Friesen, M.L.; Tiemann, L.K. To fix or not to fix: Controls on free-living nitrogen fixation in the rhizosphere. Appl. Environ. Microbiol.; 2019; 85, e02546-18.
21. De la Vega-Camarillo, E.; Sotelo-Aguilar, J.; Rios-Galicia, B.; Mercado-Flores, Y.; Arteaga-Garibay, R.; Villa-Tanaca, L.; Hernández-Rodríguez, C. Promotion of the growth and yield of Zea mays by synthetic microbial communities from Jala maize. Front. Microbiol.; 2023; 14, 1167839. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/37275168]
22. Shah, A.; Nazari, M.; Antar, M.; Msimbira, L.A.; Naamala, J.; Lyu, D.; Rabileh, M.; Zajonc, J.; Smith, D.L. PGPR in agriculture: A sustainable approach to increasing climate change resilience. Front. Sustain. Food Syst.; 2021; 5, 667546.
23. Wen, A.; Havens, K.L.; Bloch, S.E.; Shah, N.; Higgins, D.A.; Davis-Richardson, A.G.; Sharon, J.; Rezaei, F.; Mohiti-Asli, M.; Johnson, A. et al. Enabling biological nitrogen fixation for cereal crops in fertilized fields. ACS Synth. Biol.; 2021; 10, pp. 3264-3277. [DOI: https://dx.doi.org/10.1021/acssynbio.1c00049]
24. Aroney, S.T.J.N.; Poole, P.S.; Sánchez-Cañizares, C. Rhizobial chemotaxis and motility systems at work in the soil. Front. Plant Sci.; 2021; 12, 725338.
25. Hidalgo-Castellanos, J.; Marín-Peña, A.J.; Herrera-Cervera, J.A.; López-Gómez, M. Polyamines: Key elements in the rhizobia-legume symbiosis?. Phytochem. Rev.; 2022; 21, pp. 127-140.
26. Jung, J.; Kim, J.-S.; Taffner, J.; Berg, G.; Ryu, C.-M. Archaea, tiny helpers of land plants. Comput. Struct. Biotechnol. J.; 2020; 18, pp. 2494-2500. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33005311]
27. Lucke, M.; Correa, M.G.; Levy, A. The role of secretion systems, effectors, and secondary metabolites of beneficial rhizobacteria in interactions with plants and microbes. Front. Plant Sci.; 2020; 11, 589416. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33240304]
28. Krysenko, S.; Wohlleben, W. Polyamine and ethanolamine metabolism in bacteria as an important component of nitrogen assimilation for survival and pathogenicity. Med. Sci.; 2022; 10, 40. [DOI: https://dx.doi.org/10.3390/medsci10030040] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35997332]
29. Igarashi, K.; Kashiwagi, K. Effects of polyamines on protein synthesis and growth of Escherichia coli. J. Biol. Chem.; 2018; 293, pp. 18702-18709. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30108177]
30. Michael, A.J. Polyamine function in archaea and bacteria. J. Biol. Chem.; 2018; 293, pp. 18693-18701.
31. Rhee, H.J.; Kim, E.-J.; Lee, J.K. Physiological polyamines: Simple primordial stress molecules. J. Cell. Mol. Med.; 2007; 11, pp. 685-703. [DOI: https://dx.doi.org/10.1111/j.1582-4934.2007.00077.x]
32. Shah, P.; Swiatlo, E. A multifaceted role for polyamines in bacterial pathogens. Mol. Microbiol.; 2008; 68, pp. 4-16. [DOI: https://dx.doi.org/10.1111/j.1365-2958.2008.06126.x]
33. Michael, A.J. Biosynthesis of polyamines and polyamine-containing molecules. Biochem. J.; 2016; 473, pp. 2315-2329.
34. Michael, A.J. Polyamines in Eukaryotes, Bacteria, and Archaea. J. Biol. Chem.; 2016; 291, pp. 14896-14903.
35. Di Martino, M.L.; Campilongo, R.; Casalino, M.; Micheli, G.; Colonna, B.; Prosseda, G. Polyamines: Emerging players in bacte-ria–host interactions. Int. J. Med. Microbiol.; 2013; 303, pp. 484-491. [DOI: https://dx.doi.org/10.1016/j.ijmm.2013.06.008] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23871215]
36. Becerra-Rivera, V.; Dunn, M.F. Polyamine biosynthesis and biological roles in rhizobia. FEMS Microbiol. Lett.; 2019; 366, fnz084. [DOI: https://dx.doi.org/10.1093/femsle/fnz084] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31062028]
37. Dunn, M.F. Rhizobial amino acid metabolism: Polyamine biosynthesis and functions. Handbook of Microbial Metabolism of Amino Acids; D’Mello, F.J. CAB International: Oxfordshire, UK, 2017; pp. 352-370.
38. Gerlin, L.; Baroukh, C.; Genin, S. Polyamines: Double agents in disease and plant immunity. Trends Plant Sci.; 2021; 26, pp. 1061-1071. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34127368]
39. Jiménez-Bremont, J.F.; Marina, M.; Guerrero-González, M.d.l.L.; Rossi, F.R.; Sánchez-Rangel, D.; Rodríguez-Kessler, M.; Ruiz, O.A.; Gárriz, A. Physiological and molecular implications of polyamine metabolism during biotic interactions. Front. Plant Sci.; 2014; 5, 95. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24672533]
40. Navakoudis, E.; Kotzabasis, K. Polyamines: A bioenergetic smart switch for plant protection and development. J. Plant Physiol.; 2022; 270, 153618.
41. Recalde, L.; Gómez Mansur, N.M.; Cabrera, A.V.; Matayoshi, C.L.; Gallego, S.M.; Groppolyamine, M.D.; Benavides, M.P. Unravelling ties in the nitrogen network: Polyamines and nitric oxide emerging as essential players in signalling roadway. Ann. Appl. Biol.; 2020; 178, pp. 192-208. [DOI: https://dx.doi.org/10.1111/aab.12642]
42. Tilgram, J.; Sreeshma, N.; Priyadarshini, P.; Bhavyasree, R.K.; Choudhury, S.; Bharati, A.; Ashajyothi, M. Rhizosphere engineering for systemic resistance/tolerance to biotic and abiotic stress. Re-Visiting the Rhizosphere Eco-System for Agricultural Sustainability; Singh, U.B.; Rai, J.P.; Sharma, A.K. Springer Nature: Singapore, 2022; pp. 271-300.
43. Vandana, U.K.; Rajkumari, J.; Singha, L.P.; Satish, L.; Alavilli, H.; Sudheer, P.D.V.N.; Chauhan, S.; Ratnala, R.; Satturu, V.; Mazumder, P.B. et al. The endophytic microbiome as a hotspot of synergistic interactions, with prospects of plant growth promotion. Biology; 2021; 10, 101.
44. Corral-Lugo, A.; De la Torre, J.; Matilla, M.A.; Marnández, M.; Morel, B.; Espinosa-Urgel, M.; Krell, T. Assessment of the con-tribution of chemoreceptor-based signaling to biofilm formation. Environ. Microbiol.; 2016; 18, pp. 3355-3372. [DOI: https://dx.doi.org/10.1111/1462-2920.13170]
45. Vessey, J.K. Plant growth promoting rhizobacteria as biofertilizers. Plant Soil; 2003; 255, pp. 571-586. [DOI: https://dx.doi.org/10.1023/A:1026037216893]
46. Mus, F.; Crook, M.B.; Garcia, K.; Garcia Costas, A.; Geddes, B.A.; Kouri, E.D.; Paramasivan, P.; Ryu, M.-H.; Oldroyd, G.E.D.; Poole, P.S. et al. Symbiotic nitrogen fixation and the challenges to its extension to nonlegumes. Appl. Environ. Microbiol.; 2016; 82, pp. 3698-3710. [DOI: https://dx.doi.org/10.1128/AEM.01055-16]
47. Liu, W.; Wang, Q.; Hou, J.; Tu, C.; Luo, T.; Christie, P. Whole genome analysis of halotolerant and alkalotolerant plant growth-promoting rhizobacterium Klebsiella Sp. D5A. Sci. Rep.; 2016; 6, 26710. [DOI: https://dx.doi.org/10.1038/srep26710] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27216548]
48. Duran-Bedolla, J.; Garza-Ramos, U.; Rodríguez-Medina, N.; Aguilar Vera, A.; Barrios-Camacho, H. Exploring the environmenttal traits and applications of Klebsiella variicola. Braz. J. Microbiol.; 2021; 52, pp. 2233-2245. [DOI: https://dx.doi.org/10.1007/s42770-021-00630-z] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34626346]
49. Ardley, J.; Sprent, J. Evolution and biogeography of actinorhizal plants and legumes: A comparison. J. Ecol.; 2020; 109, pp. 1098-1121.
50. Op den Camp, R.H.M.; Polone, E.; Fedorova, E.; Roelofsen, W.; Squartini, A.; Op den Camp, H.J.M.; Bisseling, T.; Guerts, R. Nonlegume Parasponia andersonii deploys a broad Rhizobium host range strategy resulting in largely variable symbiotic effectiveness. Mol. Plant-Microbe Interact.; 2012; 25, pp. 954-963. [DOI: https://dx.doi.org/10.1094/MPMI-11-11-0304]
51. Gyaneshwar, P.; Hirsch, A.M.; Moulin, L.; Chen, W.-M.; Elliott, G.N.; Bontemps, C.; Estrada-de los Santos, P.; Gross, E.; Bueno dos Reis, F., Jr.; Sprent, J.I. et al. Legume-nodulating betaproteobacteria: Diversity, host range, and future prospects. Mol. Plant-Microbe Interact.; 2011; 24, pp. 1276-1288. [DOI: https://dx.doi.org/10.1094/MPMI-06-11-0172]
52. Doty, S.L.; Dosher, M.R.; Singleton, G.L.; Moore, A.L.; Van Aken, B.; Stettler, R.F.; Strand, S.E.; Gordon, M.P. Identification of an endophytic Rhizobium in stems of Populus. Symbiosis; 2005; 39, pp. 27-35.
53. Knoth, J.L.; Kim, S.-H.; Ettl, G.J.; Doty, S.L. Effects of cross host species inoculation of nitrogen-fixing endophytes on growth and leaf physiology of maize. GCB Bioenergy; 2013; 5, pp. 408-418. [DOI: https://dx.doi.org/10.1111/gcbb.12006]
54. Antoun, H.; Beauchamp, C.J.; Goussard, N.; Chabot, R.; Lelande, R. Potential of Rhizobium and Bradyrhizobium species as plant growth promoting rhizobacteria on non-legumes: Effect on radishes (Raphanus sativus L.). Plant Soil; 1998; 204, pp. 57-67. [DOI: https://dx.doi.org/10.1023/A:1004326910584]
55. Padukkage, D.; Geekiyanage, S.; Reparaz, J.M.; Bezus, R.; Balatti, P.A.; Degrassi, G. Bradyrhizobium japonicum, B. elkanii and B. diazoefficiens interact with rice (Oryza sativa), promote growth and increase yield. Curr. Microbiol.; 2021; 78, pp. 417-428.
56. Alemneh, A.A.; Zhou, Y.; Ryder, M.H.; Denton, M.D. Mechanisms in plant growth-promoting rhizobacteria that enhance legume-rhizobia symbioses. J. Appl. Microbiol.; 2020; 129, pp. 1133-1156. [DOI: https://dx.doi.org/10.1111/jam.14754] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32592603]
57. Xu, T.; Vo, Q.A.T.; Barnett, S.J.; Ballard, R.A.; Zhu, Y.; Franco, C.M.M. Revealing the underlying mechanisms mediated by endophytic actinobacteria to enhance the rhizobia-chickpea (Cicer arietinum L.) symbiosis. Plant Soil; 2022; 474, pp. 319-320. [DOI: https://dx.doi.org/10.1007/s11104-022-05402-8]
58. Bae, H.-S.; Morrison, E.; Chanton, J.P.; Ogram, A. Methanogens are major contributors to nitrogen fixation in soils of the Florida Everglades. Appl. Environ. Microbiol.; 2018; 84, e02222-17. [DOI: https://dx.doi.org/10.1128/AEM.02222-17]
59. Li, B.; Liang, J.; Hanfrey, C.C.; Phillips, M.A.; Michael, A.J. Discovery of ancestral L-ornithine and L-lysine decarboxylases reveals parallel, pseudoconvergent evolution of polyamine biosynthesis. J. Biol. Chem.; 2021; 297, 101219. [DOI: https://dx.doi.org/10.1016/j.jbc.2021.101219] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34560100]
60. Becerra-Rivera, V.A.; Bergström, E.; Thomas-Oates, J.; Dunn, M.F. Polyamines are required for normal growth in Sinorhizobium meliloti. Microbiology; 2018; 164, pp. 600-613. [DOI: https://dx.doi.org/10.1099/mic.0.000615]
61. Bertalan, M.; Albano, R.; de Pádua, V.; Rouws, L.; Rojas, C.; Hemerly, A.; Teixeira, K.; Schwab, S.; Araujo, J.; Oliveira, A. et al. Complete genome sequence of the sugarcane nitrogen-fixing endophyte Gluconacetobacter diazotrophicus Pal5. BMC Genom.; 2009; 10, 450.
62. Flores-Tinoco, C.E.; Tschan, F.; Fuhrer, T.; Margot, C.; Sauer, U.; Christen, M.; Christen, B. Co-catabolism of arginine and succinate drives symbiotic nitrogen fixation. Mol. Syst. Biol.; 2020; 16, e9419. [DOI: https://dx.doi.org/10.15252/msb.20199419]
63. Burnat, M.; Li, B.; Kim, S.H.; Michael, A.J.; Flores, E. Homospermidine biosynthesis in the cyanobacterium Anabaena requires a deoxyhypusine synthase homologue and is essential for normal diazotrophic growth. Mol. Microbiol.; 2018; 109, pp. 763-780. [DOI: https://dx.doi.org/10.1111/mmi.14006]
64. Zhang, L.; Zhong, J.; Liu, J.H.; Xin, K.; Chen, C.; Li, Q.; Wei, Y.; Wang, Y.; Chen, F.; Shen, X. Complete genome sequence of the drought resistance-promoting endophyte Klebsiella sp. LTGF-6F. J. Biotechnol.; 2017; 246, pp. 36-39. [DOI: https://dx.doi.org/10.1016/j.jbiotec.2017.02.008]
65. del Barrio-Duque, A.; Samad, A.; Nybroe, O.; Antonielli, L.; Sessitsch, A.; Compant, S. Interaction between endophytic Proteobacteria strains and Serendipita in-dica enhances biocontrol activity against fungal pathogens. Plant Soil; 2020; 451, pp. 277-305. [DOI: https://dx.doi.org/10.1007/s11104-020-04512-5]
66. Fouts, D.E.; Tyler, H.L.; DeBoy, R.T.; Daugherty, S.; Ren, Q.; Badger, J.H.; Durkin, A.S.; Huot, H.; Shrivastava, S.; Kothari, S. et al. Complete genome sequence of the N2-fixing broad host range endophyte Klebsiella pneumoniae 342 and virulence predictions verified in mice. PLoS Genet.; 2008; 4, e1000141. [DOI: https://dx.doi.org/10.1371/journal.pgen.1000141] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18654632]
67. Arteaga Ide, A.; Hernández, V.M.; Medina-Aparicio, L.; Carcamo-Noriega, E.; Girard, L.; Hernández-Lucas, I.; Dunn, M.F. Genetic regulation, biochemical properties and physiological importance of arginase from Sinorhizobium meliloti. Microbiology; 2020; 166, pp. 484-497.
68. Hernández, V.M.; Girard, L.; Hernández-Lucas, I.; Vázquez, A.; Ortíz-Ortíz, C.; Díaz, R.; Dunn, M.F. Genetic and biochemical characterization of arginine biosynthesis in Sinorhizobium meliloti 1021. Microbiology; 2015; 161, pp. 1671-1682. [DOI: https://dx.doi.org/10.1099/mic.0.000122] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26271664]
69. Hernández, V.M.; Arteaga, A.; Dunn, M.F. Diversity, properties and functions of bacterial arginases. FEMS Microbiol. Rev.; 2021; 45, fuab034. [DOI: https://dx.doi.org/10.1093/femsre/fuab034]
70. Burrell, M.; Hanfrey, C.C.; Murray, E.J.; Stanley-Wall, N.R.; Michael, A.J. Evolution and multiplicity of arginine decarboxylases in polyamine biosynthesis and essential role in Bacillus subtilis biofilm formation. J. Biol. Chem.; 2010; 285, pp. 39224-39238. [DOI: https://dx.doi.org/10.1074/jbc.M110.163154]
71. Lee, J.; Michael, A.J.; Martynowski, D.; Goldsmith, E.J.; Phillips, M.A. Phylogenetic diversity and the structural basis of sub-strate specificity in the β/α-barrel fold basic amino acid decarboxylases. J. Biol. Chem.; 2007; 282, pp. 27115-27125. [DOI: https://dx.doi.org/10.1074/jbc.M704066200]
72. Carriel, D. Structure-Function Relationships of the Lysine Decarboxylase LdcA from Pseudomonas aeruginosa. Ph.D. Thesis; Université Grenoble Alpes: Grenoble, France, 2017; 220.
73. Auling, G.; Busse, H.-J.; Pilz, F.; Webb, L.; Kneifel, H.; Claus, D. Rapid differentiation, by polyamine analysis, of Xanthomonas strains from phytopathogenic Pseudomonads and other members of the Class Proteobacteria interacting with plants. Int. J. Syst. Bacteriol.; 1991; 41, pp. 223-228. [DOI: https://dx.doi.org/10.1099/00207713-41-2-223]
74. Goris, J.; Kersters, K.; De Vos, P. Polyamine distribution among authentic Pseudomonads and Azotobacteraceae. Syst. Appl. Microbiol.; 1998; 21, pp. 285-290. [DOI: https://dx.doi.org/10.1016/S0723-2020(98)80035-0]
75. Cassán, F.; Maiale, S.; Macciarelli, O.; Vidal, A.; Luna, V.; Ruiz, O. Cadaverine production by Azospirillum brasilense and its possible role in plant growth promotion and osmotic stress mitigation. Eur. J. Soil Biol.; 2009; 45, pp. 12-19. [DOI: https://dx.doi.org/10.1016/j.ejsobi.2008.08.003]
76. Perrig, D.; Boiero, M.L.; Masciarelli, O.A.; Penna, C.; Ruiz, O.A.; Cassán, F.D.; Luna, M.V. Plant-growth-promoting compounds produced by two agronomically important strains of Azosprillum brasilense, and implications for inoculant formation. Appl. Microbiol. Biotechnol.; 2007; 75, pp. 1143-1150. [DOI: https://dx.doi.org/10.1007/s00253-007-0909-9]
77. Lin, S.-Y.; Hameed, A.; Tsai, C.-F.; Huang, G.-H.; Hus, Y.-H.; Young, C.-C. Description of Azoarcus nasutitermitis Sp. Nov. and Azoarcus rhizosphaerae Sp. Nov., two nitrogen-fixing species isolated from termite nest and rhizosphere of Ficus religiosa. Antonie Van Leeuwenhoek; 2020; 113, pp. 933-946. [DOI: https://dx.doi.org/10.1007/s10482-020-01401-w]
78. El-Tarabily, K.A.; Ramadan, G.A.; Elbadawi, A.A.; Hassan, A.H.; Tariq, S.; Ghazal, E.W.; Gamar, M.I.A.; AbuQamar, S.F. The marine endophytic polyamine-producing Streptomyces mutabilis UAE1 isolated in the Arabian Gulf promotes performance of mangrove (Avicennia manina) seedlings under greenhouse conditions. Front. Mar. Sci.; 2021; 8, 710200. [DOI: https://dx.doi.org/10.3389/fmars.2021.710200]
79. Kakehi, J.-I.; Kuwashiro, Y.; Motose, H.; Igarashi, K.; Takahashi, T. Norspermine substitutes for thermospermine in the control of stem elongation in Arabidopsis thaliana. FEBS Lett.; 2010; 584, pp. 3042-3046. [DOI: https://dx.doi.org/10.1016/j.febslet.2010.05.035] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20580714]
80. Kim, S.H.; Wang, Y.; Khomutov, M.; Khomutov, A.; Fuqua, C.; Michael, A.J. The essential role of spermidine in growth of Agrobacterium tumefaciens is determined by the 1,3-diaminopropolyaminene moiety. ACS Chem. Biol.; 2016; 11, pp. 491-499. [DOI: https://dx.doi.org/10.1021/acschembio.5b00893]
81. Hamana, K.; Takeuchi, M. Polyamine profiles as chemotaxonomic marker within alpha, beta, gamma, delta and epsilon subclasses of class Proteobacteria: Distribution of 2-hydroxyputrescine and homospermidine. Microbiol. Cult. Coll.; 1998; 14, pp. 1-14.
82. Shaw, F.L.; Elliott, K.A.; Kinch, L.N.; Fuell, C.; Phillips, M.A.; Michael, A.J. Evolution and multifarious horizontal transfer of an alternative biosynthetic polyamine pathway for the alternative polyamine sym-homospermidine. J. Biol. Chem.; 2010; 285, pp. 14711-14723. [DOI: https://dx.doi.org/10.1074/jbc.M110.107219] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20194510]
83. Shaw, F.L. From Prediction to Function: Polyamine Biosynthesis and Formate Metabolism in the α- and ε-Proteobacteria. Ph.D. Thesis; University of East Anglia: Norwich, UK, 2011; 311.
84. Wheeler, C.T.; Tonin, G.S.; Sutcliffe, A. Polyamines of Frankia in relation to nitrogen nutrition. Soil Biol. Biochem.; 1994; 26, pp. 577-581. [DOI: https://dx.doi.org/10.1016/0038-0717(94)90245-3]
85. Hamana, K.; Minamisawa, K.; Matsuzaki, S. Polyamines in Rhizobium, Bradyrhizobium, Azorhizobium and Agrobacterium. FEMS Microbiol. Lett.; 1990; 71, pp. 71-76. [DOI: https://dx.doi.org/10.1111/j.1574-6968.1990.tb03800.x]
86. Hamana, K.; Soto, W.; Guma, K.; Yu, J.H.; Ino, Y.; Umemura, Y.; Mochizuki, C.; Takatsuka, K.; Kigure, Y.; Tanaka, N. et al. Cellular polyamine catalogues of the five classes of the phylum Proteobacteria: Distributions of homospermidine with the class alphaproteobacteria, hydroxyputrescine with the class proteobacteria, norspermidine within the class gammaproteobac-teria and spermine in the classes deltaproteobacteria and epsilonproteobacteria. Ann. Gunma Health Sci.; 2006; 27, pp. 1-16.
87. Li, B.; Lowe-Power, T.; Kurihara, S.; Gonzalez, S.; Naidoo, J.; MacMillian, J.B.; Allen, C.; Michael, A.J. Functional identification of putrescine C- and N-hydroxylases. ACS Chem. Biol.; 2016; 11, pp. 2782-2789. [DOI: https://dx.doi.org/10.1021/acschembio.6b00629]
88. Hamana, K.; Hayashi, H.; Niitsu, M.; Itoh, T. Polyamine analysis of thermophilic, acidophilic, alkaliphilic and radio-tolerant bacteria belonging to the domain Bacteria and methanogens, thermophiles and extreme halophiles belonging to the domain Archaea. J. Jpn. Soc. Extrem.; 2009; 8, pp. 59-68.
89. Hanfrey, C.C.; Pearson, B.M.; Hazeldine, S.; Lee, J.; Gaskin, D.J.; Woster, P.M.; Phillips, M.A.; Michael, A.J. Alternative sper-midine biosynthetic route is critical for growth of Campylobacter jejuni and is the dominant polyamine pathway in human gut microbiota. J. Biol. Chem.; 2011; 286, pp. 43301-43312. [DOI: https://dx.doi.org/10.1074/jbc.M111.307835] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22025614]
90. Mauchline, T.H.; Fowler, J.E.; East, A.K.; Sartor, A.L.; Zaheer, R.; Hosie, A.H.F.; Poole, P.S.; Finan, T.M. Mapping the Sinorhi-zobium meliloti 1021 solute-binding protein-dependent transportome. Proc. Natl. Acad. Sci. USA; 2006; 103, pp. 17933-17938. [DOI: https://dx.doi.org/10.1073/pnas.0606673103]
91. Wang, Y.; Kim, S.H.; Natarajan, R.; Heindl, J.E.; Bruger, E.L.; Waters, C.M.; Michael, A.J.; Fuqua, C. Spermidine inversely influences surface interactions and planktonic growth in Agrobacterium tumefaciens. J. Bacteriol.; 2016; 198, 2682. [DOI: https://dx.doi.org/10.1128/JB.00265-16]
92. Duprey, A.; Groisman, E.A. DNA supercoiling differences in bacteria result from disparate DNA gyrase activation by polyamines. PLoS Genet.; 2020; 16, e1009085. [DOI: https://dx.doi.org/10.1371/journal.pgen.1009085]
93. Michael, A.J. Biosynthesis of polyamines in Eukaryotes, Archaea, and bacteria. Polyamines; Kusano, T.; Suzuki, H. Springer: Tokyo, Japan, 2015; pp. 3-14.
94. Baraquet, C.; Dai, W.; Mendiola, J.; Pechter, K.; Harwood, C.S. Transposon sequencing analysis of Bradyrhizobium diazoefficiens 110spc4. Sci. Rep.; 2021; 11, 13211. [DOI: https://dx.doi.org/10.1038/s41598-021-92534-z]
95. Busse, J.; Auling, G. Polyamine pattern as a chemotaxonomic marker within the Proteobacteria. Syst. Appl. Microbiol.; 1988; 11, pp. 1-8. [DOI: https://dx.doi.org/10.1016/S0723-2020(88)80040-7]
96. DiCenzo, G.C.; Benedict, A.B.; Fondi, M.; Walker, G.C.; Finan, T.M.; Mengoni, A.; Griffitts, J.S. Robustness encoded across essential and accessory replicons of the ecologically versatile bacterium Sinorhizobium meliloti. PLoS Genet.; 2018; 14, e1007357. [DOI: https://dx.doi.org/10.1371/journal.pgen.1007357]
97. Perry, B.J.; Akter, M.S.; Yost, C.K. The use of transposon insertion sequencing to interrogate the core functional genome of the legume symbiont Rhizobium leguminosarum. Front. Microbiol.; 2016; 7, 1873. [DOI: https://dx.doi.org/10.3389/fmicb.2016.01873]
98. Alexandre, A.; Oliveira, S. Response to temperature stress in rhizobia. Crit. Rev. Microbiol.; 2013; 39, pp. 219-228. [DOI: https://dx.doi.org/10.3109/1040841X.2012.702097]
99. Chang, C.; Damiani, I.; Puppo, A.; Frendo, P. Redox changes during the legume–Rhizobium symbiosis. Mol. Plant; 2009; 2, pp. 370-377. [DOI: https://dx.doi.org/10.1093/mp/ssn090] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19825622]
100. Ferguson, B.J.; Lin, M.-H.; Gresshoff, P.M. Regulation of legume nodulation by acidic growth conditions. Plant Signal. Behav.; 2013; 8, e23426. [DOI: https://dx.doi.org/10.4161/psb.23426] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23333963]
101. Hall, H.K.; Karem, K.L.; Foster, J.W. Molecular responses of microbes to environmental pH stress. Adv. Microb. Physiol.; 1995; 37, pp. 229-272.
102. Hawkins, J.P.; Oresnik, I.J. The Rhizobium-legume symbiosis: Co-opting successful stress management. Front. Plant Sci.; 2022; 12, 796045. [DOI: https://dx.doi.org/10.3389/fpls.2021.796045] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35046982]
103. Kanjee, U.; Houry, W.A. Mechanisms of acid resistance in Escherichia coli. Annu. Rev. Microbiol.; 2013; 67, pp. 65-81. [DOI: https://dx.doi.org/10.1146/annurev-micro-092412-155708]
104. Mandon, K.; Nazaret, F.; Farajzadeh, D.; Allong, G.; Frendo, P. Redox regulation in diazotrophic bacteria in interaction with plants. Antioxidants; 2021; 10, 880. [DOI: https://dx.doi.org/10.3390/antiox10060880]
105. Menéndez, A.B.; Calzadilla, P.I.; Sansberro, P.A.; Espasandin, F.D.; Gazquez, A.; Bordenave, C.D.; Maiale, S.J.; Rodríguez, A.A.; Maguire, V.G.; Campestre, M.P. et al. Polyamines and legumes: Joint stories of stress, nitrogen fixation and evironment. Front. Plant Sci.; 2019; 10, 1415. [DOI: https://dx.doi.org/10.3389/fpls.2019.01415]
106. Zahran, H.H. Rhizobium-legume symbiosis and nitrogen fixation under severe conditions and in an arid climate. Microbiol. Molec. Biol. Rev.; 1999; 63, pp. 968-989. [DOI: https://dx.doi.org/10.1128/MMBR.63.4.968-989.1999]
107. Imlay, J.A. Diagnosing oxidative stress in bacteria: Not as easy as you might think. Curr. Opin. Microbiol.; 2015; 24, pp. 124-131. [DOI: https://dx.doi.org/10.1016/j.mib.2015.01.004]
108. Andrio, E.; Marino, D.; Marmeys, A.; Dunoyer de Segonzac, M.; Damiani, I.; Genre, A.; Huguet, S.; Frendo, P.; Puppo, A.; Pauly, N. Hydrogen peroxide-regulated genes in the Medicago truncatula-Sinorhizobium meliloti symbiosis. New Phytol.; 2013; 198, pp. 190-202. [DOI: https://dx.doi.org/10.1111/nph.12120]
109. Jamet, A.; Mandon, K.; Puppo, A.; Hérouart, D. H2O2 is required for optimal establishment of the Medicago sativa/Sinorhizobium meliloti symbiosis. J. Bacteriol.; 2007; 187, pp. 8741-8745. [DOI: https://dx.doi.org/10.1128/JB.01130-07] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17921312]
110. Dombrecht, B.; Heusdens, C.; Beullens, S.; Verreth, C.; Mulkers, E.; Proost, P.; Vanderleyden, J.; Michiels, J. Defense of Rhizobium etli bacteroids against oxidative stress involves a complexly regulated atypical 2-Cys peroxiredoxin. Mol. Microbiol.; 2005; 55, pp. 1207-1221. [DOI: https://dx.doi.org/10.1111/j.1365-2958.2005.04457.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/15686565]
111. Ampe, F.; Kiss, E.; Sabourdy, F.; Batut, J. Transcriptome analysis of Sinorhizobium meliloti during symbiosis. Genome Biol.; 2003; 4, R15. [DOI: https://dx.doi.org/10.1186/gb-2003-4-2-r15]
112. Crockford, A.J.; Davis, G.A.; Williams, H.D. Evidence for cell-density-dependent regulation of catalase activity in Rhizobium leguminosarum bv. phaseoli. Microbiology; 1995; 141, pp. 843-851. [DOI: https://dx.doi.org/10.1099/13500872-141-4-843]
113. Lemire, J.; Alhasawi, A.; Appanna, V.P.; Tharmalingam, S.; Appanna, V.D. Metabolic defence against oxidative stress: The road less travelled so far. J. Appl. Microbiol.; 2017; 123, pp. 798-809. [DOI: https://dx.doi.org/10.1111/jam.13509] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28609580]
114. Santos, M.R.; Cosme, A.M.; Becker, J.D.; Medeiros, J.M.C.; Mata, M.F.; Moreira, L.M. Absence of functional TolC protein causes increased stress response gene expression in Sinorhizobium meliloti. BMC Microbiol.; 2010; 10, 180. [DOI: https://dx.doi.org/10.1186/1471-2180-10-180]
115. Becerra-Rivera, V.A.; Arteaga, A.; Leija, A.; Hernández, G.; Dunn, M.F. Polyamines produced by Sinorhizobium meliloti Rm8530 contribute to symbiotically relevant phenotypes ex planta and to nodulation efficiency on alfalfa. Microbiology; 2020; 166, pp. 278-287. [DOI: https://dx.doi.org/10.1099/mic.0.000886]
116. Lehman, A.P.; Long, S.R. Exopolysaccharides from Sinorhizobium meliloti can protect against H2O2-dependent damage. J. Bacteriol.; 2013; 195, pp. 5362-5369. [DOI: https://dx.doi.org/10.1128/JB.00681-13]
117. Bhat, S.V.; Booth, S.C.; McGrath, S.G.; Dahms, T.E. Rhizobium leguminosarum bv. viciae 3841 adapts to 2,4-dichlorophenoxya-cetic acid with “auxin-like” morphological changes, cell envelope remodeling and upregulation of central metabolic pathways. PLoS ONE; 2015; 10, e0123813. [DOI: https://dx.doi.org/10.1371/journal.pone.0123813]
118. Neira, G.; Vergara, E.; Cortez, D.; Holmes, D.S. A large-scale multiple genome comparison of acidophilic archaea (pH ≤ 5.0) extends our understanding of oxidative stress responses in polyextreme environments. Antioxidants; 2022; 11, 59. [DOI: https://dx.doi.org/10.3390/antiox11010059]
119. Tonner, P.D.; Pittman, A.M.C.; Gulli, J.G.; Sharma, K.; Schmid, A.K. A regulatory hierarchy controls the dynamic transcript-ional response to extreme oxidative stress in Archaea. PLoS Genet.; 2015; 11, e1004912. [DOI: https://dx.doi.org/10.1371/journal.pgen.1004912]
120. Fujihara, S. Biogenic amines in rhizobia and legume root nodules. Microbes Environ.; 2009; 24, pp. 1-13. [DOI: https://dx.doi.org/10.1264/jsme2.ME08557]
121. Lebrazi, S.; Fikri Benbrahim, K. Environmental stress conditions affecting the N2 fixing Rhizobium-legume symbiosis and adaptation mechanisms. Afr. J. Microbiol. Res.; 2014; 8, pp. 4053-4061.
122. López-Gómez, M.; Hidalgo-Castellanos, J.; Marín-Peña, A.J.; Herrera-Cervera, J.A. Relationship between polyamines and osmoprotectants in the response to salinity of the Legume–Rhizobia Symbiosis. Osmoprotectant-Mediated Abiotic Stress Tolerance in Plants; Hossain, M.A.; Kumar, V.; Burritt, D.J.; Fujita, M.; Mäkelä, P.S.A. Springer International Publishing: Cham, Switzerland, 2019; pp. 269-285.
123. López-Gómez, M.; Cobos-Porras, L.; Prell, J.; Lluch, C. Homospermidine synthase contributes to salt tolerance in free-living Rhizobium tropici and in symbiosis with Phaseolus vulgaris. Plant Soil; 2016; 404, pp. 413-425. [DOI: https://dx.doi.org/10.1007/s11104-016-2848-7]
124. Schiller, D.; Kruse, D.; Kneifel, H.; Krämer, R.; Burkovski, A. Polyamine transport and role of potE in response to osmotic stress in Escherichia coli. J. Bacteriol.; 2000; 182, pp. 6247-6249. [DOI: https://dx.doi.org/10.1128/JB.182.21.6247-6249.2000] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11029450]
125. Domínguez-Ferreras, A.; Pérez-Arnedo, R.; Becker, A.; Oliveres, J.; Soto, M.J.; Sanjuán, J. Transcriptome profiling reveals the importance of plasmid pSymB for osmoadaptation of Sinorhizobium meliloti. J. Bacteriol.; 2006; 188, pp. 7617-7625. [DOI: https://dx.doi.org/10.1128/JB.00719-06] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16916894]
126. López-Leal, G.; Tabche, M.L.; Castillo-Ramírez, S.; Mendoza-Vargas, A.; Ramírez-Romero, M.A.; Dávila, G. RNA-Seq analysis of the multipartite genome of Rhizobium etli CE3 shows different replicon contributions under heat and saline shock. BMC Genom.; 2014; 15, 770. [DOI: https://dx.doi.org/10.1186/1471-2164-15-770]
127. Fujihara, S.; Yoneyama, T. Effects of pH and osmotic stress on cellular polyamine contents in the soybean rhizobia Rhizobium fredii P220 and Bradyrhizobium japonicum A1017. Appl. Environ. Microbiol.; 1993; 59, 1104. [DOI: https://dx.doi.org/10.1128/aem.59.4.1104-1109.1993] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16348911]
128. Jantaro, S.; Mäenpää, P.; Mulo, P. Ischaroensakdi, A. Content and biosynthesis of polyamines in salt and osmotically stressed cells of Synechocystis sp. PCC 6803. FEMS Microbiol. Lett.; 2003; 228, pp. 129-135. [DOI: https://dx.doi.org/10.1016/S0378-1097(03)00747-X]
129. Tullio, L.D.; Gomes, D.F.; Silva, L.P.; Hungria, M.; da Silva Batista, J.S. Proteomic analysis of Rhizobium freirei PRF 81T reveals the key role of central metabolic pathways in acid tolerance. Appl. Soil Ecol.; 2019; 135, pp. 98-103. [DOI: https://dx.doi.org/10.1016/j.apsoil.2018.11.014]
130. Foster, J.W. Escherichia coli acid resistance: Tales of an amateur acidophile. Nat. Rev. Microbiol.; 2004; 2, pp. 898-907. [DOI: https://dx.doi.org/10.1038/nrmicro1021] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/15494746]
131. Hellweg, C.; Pühler, A.; Weidner, S. The time course of the transcriptomic response of Sinorhizobium meliloti 1021 following a shift to acidic pH. BMC Microbiol.; 2009; 9, 37. [DOI: https://dx.doi.org/10.1186/1471-2180-9-37] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19216801]
132. Draghi, W.O.; Del Papa, M.F.; Hellweg, C.; Watt, S.A.; Watt, T.F.; Barsch, A.; Lozano, M.J.; Lagares, A., Jr.; Salas, M.E.; López, J.L. et al. A consolidated analysis of the physiologic and molecular responses induced under acid stress in the legume-symbiont model-soil bacterium Sinorhizobium meliloti. Sci. Rep.; 2016; 6, 29278. [DOI: https://dx.doi.org/10.1038/srep29278] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27404346]
133. Wais, R.J.; Wells, D.H.; Long, S.R. Analysis of differences between Sinorhizobium meliloti 1021 and 2011 strains using the host calcium spiking response. Mol. Plant-Microbe Interact.; 2002; 15, pp. 1245-1252. [DOI: https://dx.doi.org/10.1094/MPMI.2002.15.12.1245]
134. Ghobakhlou, A.F.; Johnston, A.; Harris, L.; Antoun, H.; Legerge, S. Microarray transcriptional profiling of artic Mesorhizobium strain N33 at low temperature provides insights into cold adaptation strategies. BMC Genom.; 2015; 16, 383. [DOI: https://dx.doi.org/10.1186/s12864-015-1611-4] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25975821]
135. Webb, B.A.; Hildreth, S.; Helm, R.F.; Scharf, B.E. Sinorhizobium meliloti chemoreceptor McpU mediates chemotaxis toward host plant exudates through direct proline sensing. Appl. Environ. Microbiol.; 2014; 80, pp. 3404-3415. [DOI: https://dx.doi.org/10.1128/AEM.00115-14]
136. Chávez-Jacobo, V.M.; Becerra-Rivera, V.A.; Guerrero, G.; Dunn, M.F. The Sinorhizobium meliloti NspS-MbaA system affects biofilm formation, exopolysaccharide production and motility in response to specific polyamines. Microbiology; 2023; 169, 001293. [DOI: https://dx.doi.org/10.1099/mic.0.001293]
137. Braeken, K.; Daniels, R.; Vos, K.; Fauvart, M.; Bachaspatimayum, D.; Vanderleyden, J.; Michiels, J. Genetic determinants of swarm-ing in Rhizobium etli. Microbial. Ecol.; 2007; 55, pp. 54-64. [DOI: https://dx.doi.org/10.1007/s00248-007-9250-1]
138. Kusano, T.; Berberich, T.; Tateda, C.; Takahashi, Y. Polyamines: Essential factors for growth and survival. Planta; 2008; 228, pp. 367-381. [DOI: https://dx.doi.org/10.1007/s00425-008-0772-7]
139. Takahashi, T.; Kakehi, J.-I. Polyamines: Ubiquitous polycations with unique roles in growth and stress responses. Ann. Bot.; 2010; 105, pp. 1-6. [DOI: https://dx.doi.org/10.1093/aob/mcp259]
140. Rodríguez-Garay, B.; Phillips, G.C.; Kuehn, G.D. Detection of norspermidine and norspermine in Medicago sativa L. (alfalfa). Plant Physiol.; 1989; 89, pp. 525-529. [DOI: https://dx.doi.org/10.1104/pp.89.2.525]
141. Kuehn, G.D.; Rodríguez-Garay, B.; Bagga, S.; Phillips, G.C. Novel occurrence of uncommon polyamines in higher plants. Plant Physiol.; 1990; 94, pp. 855-857. [DOI: https://dx.doi.org/10.1104/pp.94.3.855]
142. Sagor, G.H.M.; Inoue, M.; Kim, D.W.; Kojima, S.; Niitsu, M.; Berberich, T.; Kusano, T. The polyamine oxidase from lycophyte Selaginella lepidophylla (SelpaO5), unlike that of angiosperms, back-converts thermospermine to norspermidine. FEBS Lett.; 2015; 589, pp. 3071-3078. [DOI: https://dx.doi.org/10.1016/j.febslet.2015.08.045] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26348400]
143. Takahashi, T.; Tong, W. Regulation and diversity of polyamine biosynthesis in plants. Plant Polyamines; Takahashi, T. Springer: Tokyo, Japan, 2015; pp. 27-44.
144. Tanaka, T.; Koyama, D.; Saraumi, M.; Motose, H.; Takahashi, T. RNA processing/modifying enzymes play key roles in the response to thermospermine in Arabidopsis thaliana. bioRxiv; 2022; 2022.09.19.508594. [DOI: https://dx.doi.org/10.1101/2022.09.19.508594]
145. von Arnim, A.G.; Jia, Q.; Vaughn, J.N. Regulation of plant translation by upstream open reading frames. Plant Sci.; 2014; 214, pp. 1-12. [DOI: https://dx.doi.org/10.1016/j.plantsci.2013.09.006] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24268158]
146. Chen, D.; Shao, Q.; Yin, L.; Younis, A.; Zheng, B. Polyamine function in plants: Metabolism, regulation on development, and roles in abiotic stress responses. Front. Plant Sci.; 2019; 9, 1945. [DOI: https://dx.doi.org/10.3389/fpls.2018.01945] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30687350]
147. Tiburcio, A.F.; Altabella, T.; Bitrián, M.; Alcázar, R. The roles of polyamines during the lifespan of plants: From development to stress. Planta; 2014; 240, pp. 1-18. [DOI: https://dx.doi.org/10.1007/s00425-014-2055-9]
148. Bunsupa, S.; Katayama, K.; Ikeura, E.; Oikawa, A.; Toyooka, K.; Saito, K.; Yamazaki, M. Lysine decarboxylase catalyzes the first step of quinolizidine alkaloid biosynthesis and coevolved with alkaloid production in Leguminosae. Plant Cell; 2012; 24, pp. 1202-1216. [DOI: https://dx.doi.org/10.1105/tpc.112.095885]
149. Irmer, S.; Podzun, N.; Langel, D.; Heidemann, F.; Kaltenegger, E.; Schemmerling, B.; Geilfus, C.M.; Zörb, C.; Ober, D. New aspect of plant–rhizobia interaction: Alkaloid biosynthesis in Crotalaria depends on nodulation. Proc. Natl. Acad. Sci. USA; 2015; 112, pp. 4164-4169. [DOI: https://dx.doi.org/10.1073/pnas.1423457112]
150. Gill, S.S.; Tuteja, N. Polyamines and abiotic stress tolerance in plants. Plant Signal. Behav.; 2010; 5, pp. 26-33. [DOI: https://dx.doi.org/10.4161/psb.5.1.10291]
151. Pottosin, I.; Olivas-Aguirre, M.; Dobrovinskaya, O.; Zepeda-Jazo, I.; Shabala, S. Modulation of ion transport across plant membranes by polyamines: Understanding specific modes of action under stress. Front. Plant Sci.; 2021; 11, 616077. [DOI: https://dx.doi.org/10.3389/fpls.2020.616077] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33574826]
152. Du, H.; Chen, B.; Li, Q.; Liu, H.; Kurtenbach, R. Conjugated polyamines in root plasma membrane enhanced the tolerance of plum seedling to osmotic stress by stabilizing membrane structure and therefore elevating H+-ATPase activity. Front. Plant Sci.; 2022; 12, 812360. [DOI: https://dx.doi.org/10.3389/fpls.2021.812360] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35095985]
153. Pál, M.; Szalai, G.; Janda, T. Speculation: Polyamines are important in abiotic stress signaling. Plant Sci.; 2015; 237, pp. 16-23. [DOI: https://dx.doi.org/10.1016/j.plantsci.2015.05.003] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26089148]
154. Alok, A.; Nag, A.; Kumar, J.; Jogam, P.; Singh, K.; Singh, S.P. Improvement of abiotic stress tolerance by modulating polyamine pathway in crop plants. Compatible Solutes Engineering for Crop Plants Facing Climate Change; Wani, S.H.; Gangola, M.P.; Ramadoss, B.R. Springer International Publishing: Cham, Switzerland, 2021; pp. 129-143.
155. Groppolyamine, M.D.; Benavides, M.P. Polyamines and abiotic stress: Recent advances. Amino Acids; 2008; 34, pp. 35-45.
156. Zepeda-Jazo, I.; Valarde-Buendía, A.M.; Enríquez-Figeroa, R.; Bose, J.; Shabala, S.; Muñiz-Murguía, J.; Pottosin, I.I. Polyamines interact with hydroxyl radicals in activating Ca2+ and K+ transport across the root epidermal plasma membranes. Plant Physiol.; 2011; 157, pp. 2167-2180. [DOI: https://dx.doi.org/10.1104/pp.111.179671] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21980172]
157. Juzoń, K.; Czyczyło-Mysza, I.; Marcińska, I.; Dziurka, M.; Waligórski, P.; Skrzypek, E. Polyamines in yellow lupin (Lupinus luteus L.) tolerance to soil drought. Acta Physiol. Plant.; 2017; 39, 202. [DOI: https://dx.doi.org/10.1007/s11738-017-2500-z]
158. Antoniou, C.; Zarza, X.; Gohari, G.; Panahirad, S.; Filippou, P.; Tiburcio, A.F.; Fotopoulos, V. Involvement of polyamine met-abolism in the response of Medicago truncatula genotypes to salt stress. Plants; 2021; 10, 269. [DOI: https://dx.doi.org/10.3390/plants10020269]
159. Zhang, G.W.; Hu, Q.Z.; Xu, S.C.; Gong, Y.M. Polyamines play a positive role in salt tolerant mechanisms by activating antioxidant enzymes in roots of vegetable soybean. Legumes Res.; 2013; 36, pp. 234-240.
160. Li, L.; Dou, N.; Zhang, H.; Wu, C. The versatile GABA in plants. Plant Signal. Behav.; 2021; 16, e1862565. [DOI: https://dx.doi.org/10.1080/15592324.2020.1862565] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33404284]
161. Podlešáková, K.; Ugena, L.; Spíchal, L.; Doležal, K.; De Diego, N. Phytohormones and polyamines regulate plant stress responses by altering GABA pathway. New Biotechnol.; 2019; 48, pp. 53-65. [DOI: https://dx.doi.org/10.1016/j.nbt.2018.07.003]
162. Xing, S.G.; Jun, Y.B.; Hau, Z.W.; Liang, L.Y. Higher accumulation of γ-aminobutyric acid induced by salt stress through stim-ulating the activity of diamine oxidases in Glycine max (L.) Merr. roots. Plant Physiol. Biochem.; 2007; 45, pp. 560-566. [DOI: https://dx.doi.org/10.1016/j.plaphy.2007.05.007] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17624796]
163. Fang, W.; Qi, F.; Yin, Y.; Yang, Z. Exogenous spermidine promotes γ-aminobutyric acid accumulation and alleviates the negative effect of NaCl stress in germinating soybean (Glycine max L.). Foods; 2020; 9, 267. [DOI: https://dx.doi.org/10.3390/foods9030267] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32131397]
164. Zhang, G.W.; Xu, S.C.; Hu, Q.Z.; Mao, W.H.; Gong, Y.M. Putrescine plays a positive role in salt-tolerance mechanisms by reducing oxidative damage in roots of vegetable soybean. J. Integr. Agric.; 2014; 13, pp. 349-357. [DOI: https://dx.doi.org/10.1016/S2095-3119(13)60405-0]
165. Seifi, H.S.; Shelp, B.J. Spermine differentially refines plant defense responses against biotic and abiotic stresses. Front. Plant Sci.; 2019; 10, 117. [DOI: https://dx.doi.org/10.3389/fpls.2019.00117]
166. Shi, Z.; Wang, Q.; Li, Y.; Liang, Z.; Xu, L.; Zhou, J.; Cui, Z.; Zhang, L.H. Putrescine is an intraspecies and interkingdom cell-cell communication signal modulating the virulence of Dickeya zeae. Front. Microbiol.; 2019; 10, 1950. [DOI: https://dx.doi.org/10.3389/fmicb.2019.01950] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31497009]
167. Solmi, L.; Rossi, F.R.; Romero, F.M.; Bach-Pages, M.; Preston, G.M.; Ruiz, O.A.; Gárriz, A. Polyamine-mediated mechanisms contribute to oxidative stress tolerance in Pseudomonas syringae. Sci. Rep.; 2023; 13, 4279. [DOI: https://dx.doi.org/10.1038/s41598-023-31239-x]
168. Wu, D.; von Roepenack-Lahaye, E.; Bantru, M.; de Lange, O.; Schandry, N.; Pérez-Qunitero, A.L.; Weinberg, Z.; Lowe-Power, T.M.; Szurek, B.; Michael, A.J. et al. A plant pathogen type III effector protein subverts translation regulation to boost host polyamine levels. Cell Host Microbe; 2019; 26, pp. 638-649. [DOI: https://dx.doi.org/10.1016/j.chom.2019.09.014]
169. Ober, D.; Hartmann, T. Homospermidine synthase, the first pathway-specific enzyme of pyrrolizidine alkaloid biosynthesis, evolved from deoxyhypusine synthase. Proc. Natl. Acad. Sci. USA; 1999; 96, pp. 14777-14782. [DOI: https://dx.doi.org/10.1073/pnas.96.26.14777]
170. Ivanova, K.A.; Tsyganov, V.E. Analysis of the expression of polyamine biosynthesis genes in nodules of garden pea (Pisum sativum L.) and the effect of exogenous treatment with polyamines on their development. Ecol. Genet.; 2021; 19, pp. 197-208. [DOI: https://dx.doi.org/10.17816/ecogen53771]
171. Fujihara, S.; Abe, H.; Yoneyama, T. A New polyamine 4-aminobutylcadaverine: Occurrence and its biosynthesis in root nodules of adzuki bean plant Vigna angularis. J. Biol. Chem.; 1995; 270, pp. 9932-9938. [DOI: https://dx.doi.org/10.1074/jbc.270.17.9932]
172. López-Gómez, M.; Cobos-Porras, L.; Hidalgo-Castellanos, J.; Lluch, C. Occurrence of polyamines in root nodules of Phaseolus vulgaris in symbiosis with Rhizobium tropici in response to salt stress. Phytochemistry; 2014; 107, pp. 32-41. [DOI: https://dx.doi.org/10.1016/j.phytochem.2014.08.017]
173. López-Gómez, M.; Hidalgo-Castellanos, J.; Iribarne, C.; Lluch, C. Proline accumulation has prevalence over polyamines in nodules of Medicago sativa in symbiosis with Sinorhizobium meliloti during the initial response to salinity. Plant Soil; 2014; 374, pp. 149-159. [DOI: https://dx.doi.org/10.1007/s11104-013-1871-1]
174. Fujihara, S.; Abe, S.; Minakawa, Y.; Akao, S.; Yoneyama, T. Polyamines in nodules from various plant-microbe symbiotic associations. Plant Cell Physiol.; 1994; 35, pp. 1127-1134. [DOI: https://dx.doi.org/10.1093/oxfordjournals.pcp.a078705]
175. Green, R.T.; East, A.K.; Karunakaran, R.; Downie, J.A.; Poole, P.S. Transcriptomic analysis of Rhizobium leguminosarum bacter-oids in determinate and indeterminate nodules. Microb. Genom.; 2019; 5, e000254. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30777812]
176. Durán, D.; Albareda, M.; García, C.; Marina, A.-I.; Ruiz-Argüeso, T.; Palacios, J.-M. Proteome analysis reveals a significant host-specific response in Rhizobium leguminosarum bv. viciae endosymbiotic cells. Mol. Cell. Proteom.; 2021; 20, 100009. [DOI: https://dx.doi.org/10.1074/mcp.RA120.002276] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33214187]
177. Ballesteros-Gutiérrez, M.; Albareda, M.; Barbas, C.; López-Gonzálvez, Á.; Dunn, M.F.; Palacios, J.M. A host-specific diaminobutyrate aminotransferase contributes to symbiotic performance, homoserine metabolism, and competitiveness in the Rhizobium leguminosarum/Pisum sativum system. Front. Microbiol.; 2023; 14, 1182563. [DOI: https://dx.doi.org/10.3389/fmicb.2023.1182563]
178. Hungria, M.; Eaglesham, A.R.J.; Hardy, R.W.F. Physiological comparisons of root and stem nodules of Aschynomene scabra and Sesbania rostrata. Plant Soil; 1992; 139, pp. 7-13. [DOI: https://dx.doi.org/10.1007/BF00012836]
179. Nahar, K.; Hasanuzzaman, M.; Rahman, A.; Alam, M.M.; Mahmud, J.-A.; Suzukin, T.; Fujita, M. Polyamines confer salt toler-ance in mung bean (Vigna radiata L.) by reducing sodium uptake, improving nutrient homeostasis, antioxidant defense and methylglyocal detoxification systems. Front. Plant Sci.; 2016; 7, 1104. [DOI: https://dx.doi.org/10.3389/fpls.2016.01104]
180. Atici, O.; Ögütcü, H.; Algur, O.F. Effect of putrescine on inducing symbiosis in chickpea and vetch inoculated with commercial or indigenous strains of Rhizobium. Symbiosis; 2005; 38, pp. 163-174.
181. Vassileva, V.; Ignatov, G. Polyamine-induced changes in symbiotic parameters of the Galega orientalis–Rhizobium galegae nitrogen-fixing system. Plant Soil; 1999; 210, pp. 83-91. [DOI: https://dx.doi.org/10.1023/A:1004636800869]
182. Vassileva, V.; Ignatov, G.; Gigova, L. Exogenous polyamines affect the attachment of Rhizobium galegae to the root surface of Galega orientalis. Cytobios; 1998; 96, pp. 141-150.
183. Wisniewski, J.-P.; Brewin, N.J. Construction of transgenic pea lines with modified expression of diamine oxidase and modified nodulation responses with exogenous putrescine. Mol. Plant-Microbe Interact.; 2000; 13, pp. 922-928. [DOI: https://dx.doi.org/10.1094/MPMI.2000.13.9.922] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/10975649]
184. Pereyra, C.M.; Dal Lago, C.C.; Creus, C.M.; Pereyra, M.A. Azospirillum baldaniorum Sp 245 inoculation affects cell wall and pol-yamines metabolism in cucumber seedling roots. FEMS Microbiol. Lett.; 2023; 370, fnad005. [DOI: https://dx.doi.org/10.1093/femsle/fnad005] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36690345]
185. Méndez-Gómez, M.; Castro-Mercado, E.; Alexandre, G.; García-Pineda, E. Oxidative and antioxidative responses in the wheat-Azosprillum brasilense interaction. Protoplasma; 2016; 253, pp. 477-486. [DOI: https://dx.doi.org/10.1007/s00709-015-0826-1] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25952083]
186. Ozawa, T.; Tsuji, T. Inhibition of growth of Bradyrhizobium japonicum bacteroid by spermidine and spermine in yeast extract. Soil Sci. Plant Nutr.; 1992; 38, pp. 375-379. [DOI: https://dx.doi.org/10.1080/00380768.1992.10416503]
187. Terakado, J.; Yoneyama, T.; Fujihara, S. Shoot-applied polyamines suppress nodule formation in soybean (Glycine max). J. Plant Physiol.; 2006; 163, pp. 497-505. [DOI: https://dx.doi.org/10.1016/j.jplph.2005.05.007]
188. Terakado-Tonooka, J.; Fujihara, S. Involvement of polyamines in the root nodule regulation of soybeans (Glycine max). Plant Root; 2008; 2, pp. 46-53. [DOI: https://dx.doi.org/10.3117/plantroot.2.46]
189. Chaulagain, D.; Frugoli, J. The regulation of nodule number in legumes is a balance of three signal transduction pathways. Int. J. Mol. Sci.; 2021; 22, 1117. [DOI: https://dx.doi.org/10.3390/ijms22031117]
190. Wheatley, R.M.; Ford, B.L.; Li, L.; Aroney, S.T.; Knights, H.E.; Ledermann, R.; East, A.K.; Ramachandran, V.K.; Poole, P.S. Lifestyle adaptations of Rhizobium from rhizosphere to symbiosis. Proc. Natl. Acad. Sci. USA; 2020; 117, pp. 23823-23834. [DOI: https://dx.doi.org/10.1073/pnas.2009094117]
191. Xie, S.S.; Wu, H.J.; Zang, H.Y.; Wu, L.M.; Zhu, Q.Q.; Gao, X.W. Plant growth promotion by spermidine-producing Bacillus subtilis OKB105. Mol. Plant-Microbe Interact.; 2014; 27, pp. 655-663. [DOI: https://dx.doi.org/10.1094/MPMI-01-14-0010-R]
192. Zhou, C.; Ma, Z.; Zhu, L.; Xiao, X.; Xie, Y.; Zhu, J.; Wang, J. Rhizobacterial strain Bacillus megaterium BOFC15 induces cellular polyamine changes that improve plant growth and drought resistance. Int. J. Mol. Sci.; 2016; 17, 976. [DOI: https://dx.doi.org/10.3390/ijms17060976] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27338359]
193. Alavi, P.; Starcher, M.R.; Zachow, C.; Müller, H.; Berg, G. Root-microbe systems: The effect and mode of interaction of stress protecting agent (SPA) Stenotrophomonas rhizophilia DSM14405T. Front. Plant. Sci.; 2013; 4, 141. [DOI: https://dx.doi.org/10.3389/fpls.2013.00141] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23717321]
194. Higashi, K.; Ishigure, H.; Demizu, R.; Uemura, T.; Nishino, K.; Yamaguchi, A.; Kashiwagi, K.; Igarashi, K. Identification of a spermidine excretion protein complex (MdtJI) in Escherichia coli. J. Bacteriol.; 2008; 190, pp. 872-878. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18039771]
195. Egamberdieva, D.; Jabborova, D.; Berg, G. Synergistic interactions between Bradyrhizobium japonicum and the endophyte Stenotrophomonas rhizophila and their effects on growth, and nodulation of soybean under salt stress. Plant Soil; 2016; 105, pp. 35-45.
196. López-Gómez, M.; Hidalgo-Castellanos, J.; Muñoz-Sánchez, J.R.; Marín-Peña, A.J.; LLuch, C.; Herrera-Cervera, J.A. Polyam-ines contribute to salinity tolerance in the symbiosis Medicago truncatula-Sinorhizobium meliloti by preventing oxidative damage. Plant Physiol. Biochem.; 2017; 116, pp. 9-17.
197. López-Gómez, M.; Hidalgo-Castellanos, J.; Lluch, C.; Herrera-Cervera, J.A. 24-epibrassinolide ameliorates salt stress effects in the symbiosis Medicago truncatula-Sinorhizobium meliloti and regulates the nodulation in cross-talk with polyamines. Plant Physiol. Biochem.; 2016; 108, pp. 212-221. [DOI: https://dx.doi.org/10.1016/j.plaphy.2016.07.017]
198. Palma, F.; López-Gómez, M.; Tejera, N.A.; Lluch, C. Involvement of abscisic acid in the response of Medicago sativa plants in symbiosis with Sinorhizobium meliloti to salinity. Plant Sci.; 2014; 223, pp. 16-24. [DOI: https://dx.doi.org/10.1016/j.plantsci.2014.02.005]
199. Camilios-Neto, D.; Bonato, P.; Wassem, R.; Tadra-Sfeir, M.Z.; Brusamarello-Santos, L.C.C.; Valdemeri, G.; Donatti, L.; Faoro, H.; Weiss, V.A.; Chubatsu, L.S. et al. Dual RNA-seq transcriptional análisis of wheat roots colonized by Azosprillum brasilense reveals up-regulation of nutrient acquisition and cell cycle genes. BMC Genom.; 2014; 15, 378.
200. Spaepen, S.; Bossuyt, S.; Engelen, K.; Marchal, K.; Vanderleyden, J. Phenotypical and molecular responses of Arabidopsis thali-ana roots as a result of inoculation with the auxin-producing bacterium Azospirillum brasilense. New Phytol.; 2014; 201, pp. 850-861.
201. Fujita, M.; Shinozaki, K. Identification of polyamine transporters in plants: Paraquat transport provides crucial clues. Plant Cell Physiol.; 2014; 55, pp. 855-861.
202. Burghardt, L.T.; DiCenzo, G.C. The evolutionary ecology of rhizobia: Multiple facets of competition before, during, and after symbiosis with legumes. Curr. Opin. Microbiol.; 2023; 72, 102281. [DOI: https://dx.doi.org/10.1016/j.mib.2023.102281] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36848712]
203. Fagorzi, C.; Mangoin, A. Endophytes: Improving plant performance. Microorganisms; 2022; 10, 1777. [DOI: https://dx.doi.org/10.3390/microorganisms10091777] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36144379]
204. Pueppke, S.G.; Broughton, W.J. Rhizobium sp. strain NGR234 and R. fredii USDA257 share exceptionally broad, nested host ranges. Mol. Plant-Microbe Interact.; 1999; 12, pp. 293-318. [DOI: https://dx.doi.org/10.1094/MPMI.1999.12.4.293] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/10188270]
205. Delamuta, J.R.M.; Ribeiro, R.A.; Ormeno-Orrillo, E.; Melo, I.S.; Martínez-Romero, E.; Hungria, M. Polyphasic evidence supporting the reclassification of Bradyrhizobium japonicum group Ia strains as Bradyrhizobium diazoefficiens sp. nov. Int. J. Syst. Evol. Microbiol.; 2013; 63, pp. 3342-3351.
206. Nelson, M.; Guhlin, J.; Epstein, B.; Tiffin, P.; Sadowsky, M.J. The complete replicons of 16 Ensifer meliloti strains offer insights into intra- and inter-replicon gene transfer, transposon-associated loci, and repeat elements. Microb. Genom.; 2018; 4, e000174. [DOI: https://dx.doi.org/10.1099/mgen.0.000174]
207. Ponappa, T.; Lawson, D.R.; Miller, R.A. Inhibition of ornithine decarboxylase activity reduces polyamine levels and growth of Agrobacterium tumefaciens. Curr. Microbiol.; 1992; 24, pp. 269-273. [DOI: https://dx.doi.org/10.1007/BF01577331]
208. Tower, P.A. Homospermidine, Spermidine, and Putrescine: The Biosynthesis and Metabolism of Polyamines in Rhizobium meliloti. Ph.D. Thesis; Oregon State University: Corvallis, OR, USA, 1987; 114.
© 2023 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/). Notwithstanding the ProQuest Terms and Conditions, you may use this content in accordance with the terms of the License.
Details
; Becerra-Rivera, Víctor A