Tabtoxin (Figure 2) is produced by P. syringae pvs. tabaci, coronafaciens and garcae, and it is the precursor of tabtoxinine-β-lactam (TβL), the biologically active form. The structure of tabtoxin consists of the active form, tabtoxinine-β-lactam (TβL) and serine or threonine [2,23] (Figure 3). The hydrolysis of the toxic moiety, TβL, linked via an amide bond to threonine is carried out by a zinc-activated aminopeptidase present in the periplasm [31]. The physiological target in plants, glutamine synthetase (GS) (EC 6.3.1.2), is irreversibly inhibited by TβL [29] (Figure 4). GS is the main enzyme in nitrogen assimilation in plants, fungi and bacteria. Nitrate, which is the major source of inorganic nitrogen available for plant is, after uptake from soil, either stored in the vacuole or converted into nitrite by nitrate reductase (NR). After conversion, nitrite enters the chloroplast (or cytosol in the root) and is reduced by nitrite reductase (NiR) into ammonia, which is subsequently converted to glutamine by glutamine synthetase (GS) using glutamate as a substrate (Figure 4) [16,32]. The inactivation by TβL is time-dependent, and the rate of inactivation is slowed by the addition of glutamate. Glutamate and TβL compete for the same site on the enzyme. The failure to recover any activity after extensive dialysis of the inactivated enzyme supports the conclusion that the enzyme is irreversibly inactivated [29]. Therefore, TβL irreversibly inhibits GS, resulting in ammonia accumulation, which disrupts the thylakoid membrane of the chloroplast and uncouples photorespiration, leading to chlorosis [16,33].

Phaseolotoxin (Figure 5) is produced by P. syringae pv. phaseolicola and actinidiae pathogens, causing halo blight of bean and canker of kiwifruit, respectively, as well as by the P. syringae pv. syringae strain CFBP3388 [35]. Phaseolotoxin is a tripeptide consisting of ornithine, alanine and homoarginine, linked on an inorganic sulphodiaminophosphinyl moiety [3] (Figure 6), and it is an inhibitor of OCT, a key enzyme in the urea cycle that converts ornithine and carbamoyl phosphate to citrulline [6] (Figure 4). The principal symptom produced by phaseolotoxin-producing strains is a chlorotic zone or halo around the necrotic infection site. One of the first studies to determine the relationship between phaseolotoxin synthesis and chlorosis symptoms concluded that chlorosis in the systemically affected leaf was caused by a toxin transported via the phloem [36]. The authors concluded that the downward movement of toxin in the petiole and lower stem may be exclusively through the phloem, while part of the upward movement in the upper stem may occur in the transpiration stream in the xylem as result of subsequent transfer of the toxin from the phloem to the xylem [36]. One decade later, a study about the effects of phaseolotoxin on the synthesis of arginine and protein suggested that one of the characteristic symptoms of plants affected by phaseolotoxin was that growth was stunned when the meristems were affected [37]. This author argued that protein catabolism might be enhanced in phaseolotoxin-treated tissues, as suggested by the findings that net protein accumulation was reduced and that many free amino acids, especially the amides glutamine and asparagine, accumulated in the affected tissue [19,38]. Moreover, the same rate of photosynthesis was found in chlorotic lesions and control tissues. The reason for the reduced chlorophyll synthesis that gives rise to phaseolotoxin-induced chlorosis was not clear, but it appears to involve a reversible block in development, due to a block in de novo arginine synthesis at a stage when the young leaf is synthesizing chlorophyll [37].

Least known because of its relatively recent discovery is the antimetabolite toxin named mangotoxin. This toxin is mostly produced by P. syringae pv. syringae, although it has been detected in pv. avellanae [25]. Mangotoxin inhibits ornithine acetyltransferase (OAT), which catalyses the synthesis of ornithine from N-acetyl ornithine (Figure 4). Its structure remains unknown, but its properties suggest that it is an oligopeptide [24]. In 1983, a short communication by Mitchell and Johnston described a toxin produced by P. syringae pv. syringae that inhibited the growth of E. coli [39]. The large zones of inhibition were prevented by arginine and ornithine but not by triglycine. This toxin mentioned by Mitchell and Johnston [39] could be the same toxin that was later described as mangotoxin by Arrebola et al. This toxin was classified as a virulence factor that contributes to the severity of the disease produced by P. syringae pv. syringae. Therefore, studies to determine the contribution of mangotoxin to the virulence and epiphytic fitness of P. syringae pv. syringae have been carried out. Mangotoxin-producing P. syringae pv. syringae strains and derivative non-producing mutants were inoculated in tomato leaflets maintained in vitro (Figure 7) All of the assayed strains grew at similar rates and reached similar population densities in and on inoculated tomato leaflets. However, the necrotic symptoms produced on tomato leaflets after P. syringae pv. syringae inoculation were clearly reduced when the mangotoxin-defective mutants were used, such that mangotoxin was confirmed as a virulence factor [15].

Additionally, most antimetabolite toxins show antimicrobial activity [22], and they could contribute to bacterial competitive ability and epiphytic fitness [21]. Experiments have revealed similar population densities of mangotoxin-producing strains and mangotoxin-defective mutants when they were inoculated individually. However, when the wild type was co-inoculated with mangotoxin-defective mutants, the mutants reached lower population densities. These results suggested that P. syringae pv. syringae strains producing mangotoxin outcompeted non-producing strains to colonize the phyllosphere [15].

Other antimetabolite toxins act by inhibiting enzymes associated with the urea cycle (Figure 4). However, it has been impossible to determine which target enzyme is inhibited by either of the producer strains, P. syringae pv. tomato and pv. apii. These toxins could also inhibit an earlier and undetermined step of the arginine biosynthesis pathway, such as acetylglutamate kinase, acetyl-glutamyl-phosphate reductase or acetylornithine aminotransferase. The specific target enzyme of this antimetabolite toxin from P. syringae pv. tomato and pv. apii remains to be determined (Figure 4) [22,26]. Additionally, another antimetabolite toxin produced by P. syringae pv. aptata and P. syringae pv. atrofaciens could inhibit enzymes that directly catalyze arginine synthesis because only arginine can reverse the toxin effect [22], but, to date, this enzyme has not been located. Finally, pv. maculicola produces a toxin with antimetabolite characteristics; the metabolic inhibition caused by this toxin was not reversed by any of 24 amino acids tested [22].

From this section, it is possible to conclude that antimetabolite toxins are compounds produced only in P. syringae species; however, their production is not generalized, making it possible to find producing and non-producing strains belonging to the same pathovar of P. syringae [40]. The antimetabolite toxins act as virulence factors that contribute to pathogen virulence, increasing the disease symptoms, although they are not a determinant of the pathogenesis of P. syringae. The known antimetabolite toxins are oligopeptides, and all of them inhibit target enzymes involved in amino acid biosynthesis, interfering with nitrogen metabolism.

The first studies on antimetabolite toxins were focused on tabtoxin, which has a chemical structure consisting of tabtoxinine-β-lactam [2-amino-4-(3-hydroxy-2-oxo-azacyclobutan-3-yl)-butanoic acid] linked to threonine (and, less often, to serine) (Figure 2 and Figure 3). Interest in this toxin produced by P. syringae dates back more than three decades. In 1975, Lee and Rapoport reported the synthesis of tabtoxinine-δ-lactam, an amino acid produced by various Pseudomonas species that appears to be one of the compounds found in the hydrolysis of tabtoxin or isotabtoxin. The other hydrolysis products were tabtoxinine and threonine [41]. These authors described tabtoxin as a relatively unstable molecule. At room temperature (25 °C) and pH 7, the biological activity of a solution of the toxin decreases with a half-life of approximately one day, as translactamization occurred to form the more stable and nontoxic δ-lactam isomer, isotabtoxin [41]. Twelve years later, Müller et al. published a study on the biosynthesis of tabtoxin, the toxin responsible for wildfire disease, using radioactive precursors. They showed that the molecular structure of tabtoxin consists of L-threonine and the unusual amino acid tabtoxinine-β-lactam, which proved to be difficult to synthesize, due to the toxin’s instability [13]. The biologically active β-lactam tabtoxin readily undergoes intramolecular transacylation on the stable but inactive δ-lactam isotabtoxin. The authors concluded that isotabtoxin appeared as an artifact of the workup, and no effort was made to isolate tabtoxin in its native form because the analysis of isotabtoxin was representative of the biosynthesis investigations [42]. This work established L-threonine as a direct precursor of the threonine moiety of tabtoxin. Therefore, and because the biosynthesis of threonine was well known, the corresponding moiety of tabtoxin could serve as an internal standard for the interpretation of the incorporation of labeled aspartate, glycerol, and acetate into the tabtoxinine moiety. Thus, L-aspartic acid was established as the biogenetic origin of the side chain of tabtoxinine-β-lactam. The question of how β-lactam ring formation might proceed was subsequently raised [42]. A hypothetical mechanism for β-lactam ring closure in the biosynthesis of tabtoxin was formulated according to the mechanism of a similar photoprocess [43]. However, the pathways proposed at that time were based on a chemical reaction point of view. The synthesis pathway of this toxin and its relationship to the lysine pathway was not determined until tabtoxin biosynthesis had been genetically characterized [6].

Phaseolotoxin, like tabtoxin, is composed of several amino acids linked to an inorganic moiety. Phaseolotoxin is a tripeptide consisting of ornithine, alanine and homoarginine linked to an inorganic sulphodiaminophosphinyl moiety (Figure 5 and Figure 6A). Patil reported in 1974 that acid hydrolysis of “phaseotoxin” yielded glycine, serine, valine and two unidentified amino acids [44]. The fraction corresponding to the two unknown amino acids produced chlorosis in bean leaves but, apparently, did not cause the characteristic accumulation of ornithine. In contrast, two years later, Mitchell analyzed a new antimetabolite toxin to which a new trivial name, phaseolotoxin, was given because it was produced by Pseudomonas phaseolicola [3]. Phaseolotoxin contained only three amino acids, none of which was apparently contained in “phaseotoxin”, produced symptoms resembling halo blight when applied to bean leaves and was completely characterized as the compound (N^δ-phosphosulphamyl)-ornithyl-alanyl-homoarginine. For these reasons, Mitchell could not assume that the properties of phaseolotoxin were the same as those described in the literature for “phaseotoxin” [3]. This author identified the N^δ-substituted ornithine group as the effective part of phaseolotoxin. However, he observed that there was an additional feature of phaseolotoxin that made it a potential competitive inhibitor of OCT. Moreover, N^δ-phosphosulphamylornithine (PSorn or octicidine [30]) (Figure 6A) was the major product related to detected phaseolotoxin, and it showed the same toxicity as phaseolotoxin in bioassays. This product had a chemical structure in which an analogue of carbamoyl phosphate [3] was attached to ornithine (Figure 6B), potentially accounting for the known effect of bean halo blight toxin on ornithine accumulation and OCT inactivation [45]. OCT condenses ornithine with carbamoyl phosphate to produce citrulline (Figure 3). It might, therefore, be of significance that the substituent group in phaseolotoxin, sulphamoyl phosphate, could be regarded as a simple analogue of carbamoyl phosphate in which >C=O was replaced by >SO₂ [3] (Figure 6C). More than one decade later, the biosynthetic precursors for the N^δ-(N'-sulphodiaminophosphinyl) moiety of phaseolotoxin have still not been identified. Märkisch and Reuter published in 1990 that the homoarginine and ornithine residues of phaseolotoxin were synthesized by a transamination reaction from arginine and lysine. The amidinotransferase showed high substrate specificities for arginine and lysine in phaseolotoxin-producing strains [46].

Mangotoxin was detected in the 1990s by the Escherichia coli growth inhibition assay, testing Pseudomonas syringae pv. syringae isolated from mango trees affected by bacterial apical necrosis [26,47]. Preliminary data about the chemical nature of mangotoxin was obtained using toxic cell-free filtrates subjected to different treatments. The toxic activity resisted high temperatures (100 °C) and extreme pH (2 and 12). In contrast, treatments with proteinase K and pronase eliminated the toxic activity. Ultrafiltration tests revealed that mangotoxin had a molecular size of less than 3 kDa and showed a hydrophilic nature after treatment with an organic solvent. These results were compatible with mangotoxin being an oligopeptide. By TLC, the toxic activity from P. syringae pv. syringae, corresponding to mangotoxin, was associated with a single growth-inhibiting spot with an Rf-value of 0.62. This spot was not detected in filtrates from the non-producer mutant. The mangotoxin spot was specifically detected because E. coli growth inhibition was reversed with ornithine, whereas the toxicity of tabtoxin and phaseolotoxin remained. Further chromatography analysis by HPLC showed that toxic activity eluted after 10 min was associated with a single peak, which was detected only in filtrates from the wild-type strains and was not present in filtrates from non-producing mutant derivatives [24]. The most recent knowledge about the chemical structure of mangotoxin, obtained using mass spectrometry, suggests that mangotoxin consists of two amino acids linked to a sugar residue [48].

In summary, antimetabolite toxins are small oligopeptide molecules of two or three amino acids linked to organic moieties that are directly involved in the inhibition of target enzymes. These molecules or derivatives interfere with the enzyme-substrate bond by forming an irreversible link with the catabolic domain of the target enzyme.

Tabtoxinine-β-lactam (TβL) is an irreversible inhibitor of GS; however, GS is also present in P. syringae cells. Purified GS from P. syringae pv. tabaci is irreversibly inhibited in vitro by TβL, and the inactivation is slower in the presence of glutamate, therefore suggesting that the inactivation is active-site directed [31]. This enzyme is regulated by an adenylylation system in which the GS is unadenylylated when the culture is growing with nitrate as the sole nitrogen source and is adenylylated when the culture is supplied with ammonia as a nitrogen source. Thus, GS from P. syringae pathovars is regulated in a similar manner as other gram-negative bacteria [31,49]. However, the GS of pathovar tabaci becomes partially adenylylated when TβL production is initiated. Knight et al. conducted a study about the self-protection of P. syringae pv. tabaci from its TβL. Their results showed that GS from pathovar tabaci is only partially adenylylated during TβL production, and a significant amount of GS activity remains available to the bacterium for continued assimilation of ammonia and glutamine production. Investigation of this adenylylation showed that zinc and TβL did not affect the adenylylation state of GS. However, the addition of serine was immediately followed by partial adenylylation of GS in the Tox⁺ strains growing in the tabtoxin-producing mode. These results strongly suggested that the release of serine and, probably, also threonine upon the zinc-activated hydrolysis of tabtoxin causes the partial adenylylation of GS in TβL-producing cultures [31]. Knight also observed that increasing the adenylylation state of GS slowed the rate of the inactivation, such that fully adenylylated GS was completely protected from TβL, and partially adenylylated GS was inactivated very slowly. These finding suggested that adenylylation of GS during toxin production provides partial protection of GS, and the presence of the well-established end-product effectors of GS can increase this protection [31]. However, GS adenylylation is not the only means by which P. syringae pv. tabaci is protected. Characterisation studies of β-lactamases from P. syringae using the strain BR2 identified three different β-lactamases. This kind of enzyme detoxifies TβL by cleaving the β-lactam ring to produce tabtoxinine, an inactive product. Mutants defective in tabtoxin production and tabtoxin-sensitive mutants still produced β-lactamase [50].

The mode of action of phaseolotoxin on ornithine catabolism was determined by varying the concentration of phaseolotoxin at different concentration of ornithine [51]. Phaseolotoxin appeared to be competing for the carbamoyl phosphate binding site of OCT. The toxin also appeared to be able to bind to the enzyme-carbamoyl phosphate complex. The presence of a transient phase of enzyme activity before a steady state was reached showed that phaseolotoxin shares a characteristic of tight-binding inhibitors in that it had low rates of association with and dissociation from the enzyme [51]. However, co-migration of labeled [³⁵S]PSorn and OCT during gel-filtration chromatography provided further evidence of a covalent link between [³⁵S]PSorn and OCT, such that 1 mol of PSorn is necessary to inactive 1 mol of OCT [52]. This enzyme is also common in P. syringae pv. phaseolicola; however, it is insensitive to its own toxin. In 1987, Peet et al. published the presence of genes in P. syringae pv. phaseolicola encoding the two OCTs that differ in their nucleotide sequences, conferring different sensitivities to purified phaseolotoxin. These authors concluded that the genes encoding the phaseolotoxin-insensitive OCT are part of a gene cluster involved in phaseolotoxin production. The close linkage between these genes suggested an explanation for the simultaneous loss of the ability to produce phaseolotoxin and the loss of phaseolotoxin resistance observed in naturally occurring isolates of the pathogen [53].

Mangotoxin is the most recent antimetabolite toxin to be detected, and its mode of action is still not well characterized. In 2003, Arrebola et al. published the preliminary characterization of an antimetabolite toxin produced by P. syringae pv. syringae strains that they named mangotoxin because almost all producing strains were isolated from mango trees. The antimetabolite toxin activity that was produced by many P. syringae pv. syringae strains was not reversed when N-acetyl glutamate or N-acetyl ornithine were added (as determined with the E. coli inhibition assay), but it was completely reversed after ornithine was added, suggesting that mangotoxin targets the catalysis of ornithine from N-acetyl ornithine. Therefore, the target enzyme could be either N-acetylornithinase (AO, EC: 3.5.1.16) or ornithine N-acetyltransferase (OAT, EC: 2.3.1.35) (Figure 4), neither of which had previously been described as target of an antimetabolite toxin. To clarify which enzyme, OA or OAT, was the specific target of mangotoxin, a known inhibitor of OAT, p-chloromercuribenzoic acid (PCMB), was used, and its effects were compared with the inhibition obtained by toxic filtrates from mangotoxin-producing P. syringae pv. syringae strains. A strong decrease of OAT activity was observed when cell-free filtrates from P. syringae pv. syringae strains that produce mangotoxin were assayed. The OAT activity decrease was similar to that observed with PCMB. Thus, OAT was established as the target of mangotoxin [24]. OAT is also present in bacteria (Figure 4), but mangotoxin does not seem to affect the bacterial versions of the enzyme. It is possible that mangotoxin-producing bacteria synthesize a mangotoxin-resistant OAT, although the gene that encodes this resistant OAT is still unknown. Moreover, it is also possible that mangotoxin-producing strains perform the OAT catabolic process using alternatives pathways in which OAT is not involved (Figure 4).

In summary, antimetabolite toxins act as analogues to natural substrates of their respective target enzymes, competing with the catalytic moiety of the enzyme and blocking the enzyme activity. Almost all of the target enzymes except GS, are involved in the urea cycle, and all of them act in the biosynthesis of essential amino acids for protein biosynthesis and nitrogen assimilation.

In the early 1990s, a chromosomal region from Pseudomonas syringae pv. tabaci strain BR2 that contained genes for tabtoxin biosynthesis and resistance was cloned [54]. The authors described the biosynthetic region as physically unstable, and spontaneous deletions in this region resulted in both tabtoxin-deficient and tabtoxin-sensitive bacteria and eliminated disease symptoms on bean. Therefore, Kinscherf demonstrated that tabtoxin was required by BR2 for both chlorosis and lesion formation on beans; however, toxin production was not required for growth in plants [54]. Investigations of the genes in this region and their putative functions have shed some light on the synthesis of tabtoxin. Several genes in the cluster showed high sequence identity to genes of the bacterial lysine biosynthesis pathway. Thorough studies have demonstrated that tabtoxin biosynthesis proceeds along the lysine pathway, branching off after tetrahydropicolineate formation and before diaminopimelate formation (Figure 3). Specifically, pyruvate and aspartate semialdehyde are linked by dihydropicolineate synthase, producing dihydropicolineate. Next, the enzyme dihydropicolineate reductase produces tetrahydropicolineate. From this branch point, tabtoxin is assembled by the action of TblS (a putative β-lactam synthetase), TblC (a putative clavaminic acid synthase), TblD (a putative GNAT acyltransferase), and TblE in conjunction with TblF (a putative membrane protein, forming a functional pair with a D-Ala-D-Ala ligase). Upon assembly, the completed tabtoxin can be converted by the metallopeptidase TabP into the toxin TβL (Figure 3), which is subsequently exported by the transporter TblR. The cleavage of tabtoxin occurs only in the presence of zinc, which is required for peptidase activity [6]. Outside of the tabtoxin gene cluster, the gene ttr (tabtoxin resistance gene), encodes an acetyltransferase that confer resistance to TβL. Transgene studies indicated that the overexpression of Ttr abolished the chlorotic symptoms caused by either TβL challenge or infection with P. syringae pv. tabaci. The crystal structure revealed that Ttr is a member of the GCN5-related N-acetyltransferase (GNAT) superfamily [55], members of which have been identified as bacterial aminoglycoside acetyltransferases that became resistant to the action of the antibiotics gentamicin and kanamycin [56,57]. The characterization of the Ttr structure suggested that this protein might utilize a single-step catalysis mode to transfer the acetyl group of acetyl-coenzyme A (AcCoA) directly to TβL. Furthermore, histone acetyltransferase (HAT) activity was found, implying an evolutionary link between Ttr and the other members of the GNAT superfamily. He et al. proposed a single-step catalytic mechanism as follows: first, despite the large, hydrophobic nature of the putative TβL-binding pocket, the polar environment formed by residues Glu92, Asp130 and Lys95 could assist TβL docking at the active site by adding a tightly bound water molecule. The build-up of negative charge on the oxygen facilitates a nucleophilic attack on the imide carbon of the lactam, disclosing the lone pair on the imide nitrogen of the lactam, which subsequently makes a direct nucleophilic attack on the carbonyl carbon of AcCoA. This carbonyl carbon is polarized by Tyr141, Leu96 and Lys95 through one strong and two alternate hydrogen bonds, respectively. As a result, the C-S bond on AcCoA and C-N bond on the lactam are broken, and TβL is acetylated on the imide nitrogen of the lactam. The negatively charged sulphur of CoA could subsequently be protonated and thus stabilized. Finally, both CoA and the acetylated product leave the catalytic site [55]. Thus, the toxic TβL can be transformed into a non-toxic product.

Phaseolotoxin biosynthesis is not as clear as that of tabtoxin. From very early in phaseolotoxin study, temperature was determined to regulate phaseolotoxin production. The chlorosis associated with P. syringae pv. phaseolicola infection on beans was induced at cooler temperatures (18–20 °C) but not at warmer temperatures (28-32 °C) [58]. Subsequent studies showed that phaseolotoxin production decreased progressively at temperatures above 18 °C [59]. Rowley et al. observed the production of a repressor when P. syringae pv. phaseolicola was grown at 28 °C, which is a temperature unfavorable for phaseolotoxin production [60]. Production of this repressor at the nonpermissive temperature could explain the thermoregulation of phaseolotoxin biosynthesis [4]. On the other hand, Zhang and Patil suggested in 1997 that the ORF3 product of the phtE locus might catalyze the formation of the ornithine needed for phaseolotoxin production, but biochemical evidence for this function was lacking [4,61]. Several years later, Hernández-Guzman and Álvarez-Morales isolated and characterized the gene encoding an amidinotransferase involved in the biosynthesis of phaseolotoxin [62]. These authors identified the gene amtA from P. syringae pv. phaseolicola, which showed sequence similarity to L-arginine:inosamine-phosphate amidinotransferase. The cysteine, histidine and aspartic acid residues involved in substrate binding were conserved. When Hernández-Guzman and Álvarez-Morales studied amtA expression in P. syringae pv. phaseolicola grown at 18 °C and 28 °C, they observed that the gene expression was thermoregulated, showing significant expression only at the lower temperature and following a pattern similar to that obtained for argK, the gene encoding the phaseolotoxin-resistant OCT (ROCT). This data suggested that they had obtained a fragment from an amidinotransferase gene involved in homoarginine biosynthesis [62]. Actually, the cluster encoding phaseolotoxin production genes (argK-tox) contains the gene argK, which encodes the phaseolotoxin-resistant OCT [62,63]. Templeton et al. suggested that, although the toxin-insensitive enzyme has significantly lower affinity for carbamoyl phosphate and binds ornithine more slowly than the sensitive enzyme, it should still efficiently catalyze the synthesis of citrulline, provided that the concentration of ornithine is sufficiently high [64]. Peet et al. further showed that the OCT-resistant enzyme was not thermosensitive in vivo and could presumably support maximal growth rates of P. syringae pv. phaseolicola at 30 °C. The two enzymes could be maintained by the pathogen to deal with the regulatory constraints imposed by the coordination of phaseolotoxin and arginine biosynthesis [53]. Subsequent studies of OCT-resistant determined that the STRTR binding motif in the carbamoyl phosphate domain was changed to SGRTS in ROCT. In addition, the sequences following the SMG residues in the SMG loop were different. Of the many residues whose side chains form hydrogen bonds with PSorn in the complex with OCT, Gln82 was altered to arginine in ROCT. While the side chain of this arginine, Arg84 in ROCT, could also bind the phosphate group of carbamoyl phosphate, this and other changes to the binding site were sufficient to predict a small but significant difference in the physical and electrostatic topology of the carbamoyl phosphate binding domain. ROCT retained a largely ordered mechanism with carbamoyl phosphate being the first substrate to bind, although there was a random element to the mechanism that was not observed in other OCTs. The higher binding constant for carbamoyl phosphate and conventional inhibitors indicated that the carbamoyl phosphate binding site was altered. Residues thought to be involved in stabilizing the tetrahedral carbon in the transition state were observed in ROCT, however, and this finding suggested that the chemical mechanism of ROCT is also involved in the formation of a tetrahedral intermediate. Structural analysis of the OCT/PSorn complex indicated that PSorn is a strong mimic of the transition state of OCT [65].

The synthesis of mangotoxin is currently under study. To detect putative genes involved in mangotoxin production, mangotoxin-defective mutants have been obtained by transposon mutagenesis. Important information has been obtained from several mutants. One of them helped to characterize a gene (mgoA) that encodes a non-ribosomal peptide synthetase (NRPS) and is involved in mangotoxin production. Homologues of mgoA are present in other P. syringae strains, such as pv. syringae B728a, pv. tomato DC3000 or pv. phaseolicola 144A, that fail to produce mangotoxin. This finding suggests the presence of specific genes that are involved in mangotoxin production. In fact, analysis of two Tn5 mutants showed evidence of at least one other gene cluster involved in mangotoxin production and not present in the P. syringae pathovars mentioned. One of these two Tn5-mutants is disrupted in a gene encoding a putative carboxylase that does not show significant identity with any gene in the sequenced strains of P. syringae and seems to be directly involved in mangotoxin biosynthesis [14]. The analysis of the other Tn5-mutants, which were unable to produce mangotoxin, showed disruptions in the gacA or gacS genes. The GacA/GacS two-component system controls the production of several pathogenicity and virulence factors, ecological fitness, quorum-sensing systems and synthesis of secondary metabolites in different species of Pseudomonas [66], and available data suggest that GacA/GacS also regulates antimetabolite toxin production [14,66,67].

Therefore, antimetabolite toxins are biosynthesized by non-ribosomal pathways; these toxins are assembled by enzymatic reactions using amino acids and inorganic residues as substrates. The enzymes responsible for antimetabolite toxin biosynthesis are generally encoded in gene clusters that contain genes encoding biosynthetic enzymes that confer self-protection to toxin-producing bacteria.