Guanosine pentaphosphate - review (2023)

7.4 Alarm indication: RelA, RelQ and CodY

Accumulationguanosine pentaphosphate(ppGpp) often serves as an "alarm", a stress response signal (Irving, Choudhury i Corrigan, 2020). The production of ppGpp by RelA is triggered in response to stress, such as amino acid or carbon deficiency (Magnusson, Goodbye and Nystrom, 2005). Two ppGpp synthetases, RelA and RelQ, have been identifiedThe same.RelA mutants differentially express 294 genes, including genes related to metabolism, cell division and growth (Zhang i in., 2016). Both RelA and RelQ are involved in virulence because the expression of many genes associated with virulence factors is under the control of RelA/Q or ppGpp. RelA and RelQ expression is associated with human epithelial cell adhesion and immune evasion, and contributes to virulence in a mouse model (Zhu i sur., 2016). There also seems to be a link between RelA and RelQ with the DNA-binding repressor protein CodY. The additional deletion of CodY in RelA/Q deficient mutants significantly reduces both adherence and survival in whole blood, although this mutant produces a much thicker capsule. The virulence in the mouse model was also reduced, and it has been suggested that many virulence factors influenced by RelA/Q may also be co-regulated by CodY, suggesting a link between these global regulators (Zhu i sur., 2019).

B.2.4. The relA gene encoding the (p)ppGpp alarm

Another well-known global regulatory system is the stringent response that allows microorganisms to adapt to suboptimal growth conditions (Chatterji i Ojha, 2001). The reaction is mediated by synthesisguanosine pentaphosphateand guanosine tetraphosphate, collectively called (p)ppGpp, which act as alarm signals to initiate an intense stress response and coordinate the entry of the bacterial cell into the stationary phase. atAgrobacterium,The bifunctional enzyme synthetase and hydrolase (p)ppGpp is involved in the production of (p)ppGpp, which has been experimentally investigatedA. tumefaciensA6, where, as expected, the gene encoding (relA) is transcribed only during the stationary growth phase (Zhang i in., 2004). The corresponding gene in LTU50 with which it is 90% identicalrelA_A6and 96% identity withrelA_C58, was discovered by random Tn mutagenesis, as a defect resulted in curdlan deficiency (Taner i on., 2008). The colony morphology of the mutant on aniline blue agar was characterized by separation of the bacteria into a white colony line that was stable and a light blue colony line that was unstable and maintained both the white and light blue forms after subculturing. Consistent with the staining reaction, both lines produced little recoverable curdlan (6% vs. 2% for light blue and white lines, respectively).

In complementation studies, the mutant restored curdlan production when the clone was reintroducedrelAgene, confirming that the defect inrelAis responsible for the curdlan-deficient phenotype and not for the polarity of downstream genes that are transcribed in the same direction asrelA.ParticipationrelAin polymer production is consistent with the formation of curdlate as a secondary metabolite after cell growth is complete. Use of a constitutive expressionrelAthe complementation study also found that low expression of the gene resulted in colonies that were evenly stained with aniline blue, while increased expression of P_gumilakaThe vector plasmid promoter restored curdlan production in some colonies (about 50% stained dark blue) but not in others (which stained light blue). This heterogeneity likely reflects a significant readjustment of the complex and finely tuned cell physiology to unnaturally elevated levels of a key global regulator. Not surprisingly, RelA plays multiple roles in a variety of bacteria, including control of biofilm formation, cell division, and long-term viability (Braeken in sur., 2006), appeared in pleiotropic sequencesrelAmutation in LTU50. In addition to reducing curdlan production, the mutant showed a reduced growth rate with minimal amounts of saltThe medium formed abnormally large cell clumps and produced a brown colored metabolite after prolonged incubation. Also cells fromrelAThe mutants were elongated compared to the LTU50 mutants (~3.6 Pm vs. 2.5 Pm). All these characteristics were restored by feeding the wild typerelAgenes, with the exception of flocculation, which remained at a reduced level in the complemented strain (Taner i on., 2008).

In conclusion, a relatively large number of genes have been shown to influence curdlan production, and others undoubtedly remain to be discovered. Studies to optimize curdlan yield in batch and commercial cultures have shown that curdlan production occurs in the post-stationary phase of growth and depends on many factors, including C and N sources, phosphate, sulfate and cationic composition of the culture medium, as well as pH and degree of permeability substrate (Lee, 2002 (monograph).;McIntosh i sur., 2005). It is likely that the RelA-mediated stress response initiates a cascade of events leading to curdlan production. However, the initiating event and entry into the stationary phase are not sufficient to initiate curdlan production as this only occurs in a chemically defined medium when the N source is depleted and not at all in a nutrient-rich medium. The involvement of NtrBC and NtrYX as activators in the cascade is consistent with the observed relationship between curdlan production and nitrogen supply, as well as the role of the Ntr protein as a sensor of nitrogen deficiency. CrdR may therefore intervene at an even later stage in the cascadefor CPlasmid carrying mutantscrdRThe gene may produce some curdlan on aniline blue agar (Aračić, 2009. (encyclopedic entry).). The target genes of each activator are unknown, and it seems unlikely that NtrC will directly activate any of themcrdRLubcrdASCbecause the areas above these locations contain no visible σ⁵⁴-similar promoter consensus sequence (Dombrecht et al., 2002 (encyclopedia entry).).

Curdlan production is also strain specific and the required genes appear to be permanently suppressed in many wild strainsAgrobacteriumtribes. Such strains typically produce succinoglycan, an acidic heteropolysaccharide, and produce no or very little curdlan (Dr Nakanishi, 1976).A. tumefaciensAn example is the C58 strain: it is a producer of succinoglycan (Changelosi and sur., 1987) and although it has homologscrdASCGene (and others described above) produce little curdlan. On the other hand, mutants spontaneously producing curdlan, derived from wild-type strains (e.g. LTU50 line) lose the ability to produce succinoglycan (Hisamatsu i dr, 1977), indicating a negative correlation in the production of these polysaccharides. The production and regulation of succinoglycan is poorly understoodAgrobacterium(Ard i sur., 1991;Changelosi and sur., 1987;Tiburtius and Dr., 1996), but are well documented inS. melodywhere the succinoglycan and galactoglucan production pathways are negatively co-regulated (Becker i Puhler, 1998). Hence the superficially similar regulatory interaction between the succinoglycan and curdlan production pathwaysThe replacement of the latter may explain why curdlan production has been overlooked as a feature of agrobacterial biology and raises questions about its role (seeChapter 4.1).

the (p)ppGpp cycle

The hydrolysis reaction performed by SpoT/RSH is located in the N-terminal domain of these proteins (illustration 1). The activity and conformation of this domain is affected by ligand binding to the adjacent synthetase domain and possibly additional interactions originating from the C-terminal domain. The regulation of activity is reciprocal - when one increases, the other decreases. The hydrolysis reaction itself requires Mn²⁺and is specific for the removal of 3'-pyrophosphate from ppGpp and pppGpp with no known other substrates. The TGS domain is largely conserved in RSH enzymes; For SpoT, it has been identified as the binding site for acylated acyl carrier proteins. The SpoT synthetase domain is weakly active compared to most RSH enzymes. The function of the conserved ACT domain is unknown, although mutants in this region, as well as in other parts of the C-terminal domain, may limit dimerization, the physiological consequences of which are unknown.

Global regulation

It is an axiom that the processes of intermediary metabolism between prokaryotes vary greatly depending on the ecological niche occupied by a given organism, while the mechanisms of synthesis and macromolecular regulation remain highly conserved regardless of ecological considerations. This situation largely concerns macromolecular synthesisY. pestisbut not for global regulation.Y. pestisit undergoes traditional DNA repair processes and is stringent (i.e. it produces guanosine tetra- and pentaphosphate after removal of an essential amino acid). As recent genomic analysis shows, plague bacilli develop common key global regulators, including cAMP receptor protein (CRP), leucine regulatory protein (Lrp), DNA double bond transcription regulator PhoP and its sensor protein PhoQ, transcription regulator RovA (MARS family member/ SlyA) and H-NS nucleoid structural protein. However, many of these systems have been significantly redesigned compared to those inE colialso consider recent publicationsY. pestisfrom severe environmental constraints and the need to adapt to unique in vivo signals (although these modifications are sometimes shared with others).Y. pseudotuberkuloza). The classic two-component PhoP/PhoQ regulatory system is required for survivalY. pestisyou Mg²⁺-deficiency of macrophage phagolysosomes[47]; This effect is partly due to the induction of the magnesium translocating P-type ATPase (MgtA). This is due to the high osmolarity of the mammalian intestine (≥300 mOsm).E colito increase OmpC porin (small pore size) and decrease OmpF (pore size), while in soil and water the opposite situation occurs where osmolarity is significantly reduced. In this case, CRP-regulated phosphorylated OmpR gradually binds to the upstream regionsompFIompCas a function of osmolarity until saturation followed by transcriptionompFstops due to the formation of a circular loop. However, inY. pestisCRP directly recognizes the promoterompCIompFbut not its own structural gene and therefore has no effect onompR [48]. The PhoP/PhoQ system normally controls a number of other functions, including the transcription of the RstA/RstB binary regulatory system (controlling acid-fast genes), its own autoregulator (MgrB), initiator of hemin synthesis (Heml), and surface lipoproteins (e.g., B., SlyB), which are also reduced in Mg²⁺Surplus. TheThe regulated PhoP and RovA are integratedY. pestisfound that PhoP downregulates RovA by recognizing its promoter, while RovA directly activates the transcription of its own structural gene[49]. In addition, PhoPY. pestisis capable of both positive and negative autoregulation and acts as an activator of adenylyl cyclase transcriptioncrpThis shows that the PhoP and CRP regulators of this organism have evolved into a single regulatory cascade[50].

2.3 PNPaze structure

Pure PNPase was isolatedE colistrong subunit of the homotrimer with a mass of 78 kDa.⁸⁴X-ray crystal structuresE coli,Streptomyces antibioticIC is increasingPNPase reveals the organization of a homotrimeric subunit with ring architecture (Thread. 3).^11,51,85,86Each monomer has an assembly of five domains: at the N-terminus, the two RNase PH domains (first and second core domains) are connected by an α-helical domain⁸⁷; At the C-terminus are two RNA binding domains called KH and S1⁸⁸(Thread. 3). In the quaternary structure, the KH and S1 domains are on one side of the trimer, while the active site (RNase PH domain) is on the opposite side. The three subunits associate through the trimerization sites of the core domains and form a central channel where catalysis takes place. The presence of conserved basic residues in the gate region as well as adequate channel constriction are critical for RNA capture for process degradation.⁸⁵Two constriction points in the channel were identified, and the structure of the PNPase in complex with RNA clearly indicates that the path followed by the RNA molecule is along the central pore towards the active site.^11,51,85The dynamic translocation of RNA by the enzyme depends on the conformational changes that occur in the opening of the central channel and adjacent regions.⁵¹Kinetic studies have shown that the presence of distinct and different RNA binding sites is partly responsible for the processivity of the PNPase. The trimeric channel probably contributes to the processivity and regulation of PNPase activity through RNA structural elements.¹¹

The first core RNase PH domain has a conserved FFRR loop that interacts with the RNA about 20 Å from the putative catalytic site in the second core.⁸⁹TwoE coliPNPase mutants in the FFRR loop (Arg79Ala and Arg80Ala) showed a strong increasek_Mfor ADP/Pi binding. In addition, it has been suggested that the first primary domain is involvedguanosine pentaphosphate(pppGpp) synthetase activityS.antibioticPNPase, absent in most PNPases.¹¹The second core domain of RNase PH binds tungstate, a phosphate analog, suggesting that a catalytic center is located here. This region may also have been separately adapted as a second active site in PNPases overall.¹¹Most of the conserved residues are concentrated in the second core domain (which contains the putative catalytic site) and a small portion of the first core domain. Few of them are in the KH or S1 RNA-binding domains. Mutations in the amino acids around the tungstate binding site abolish or significantly reduce all catalytic activities of the enzyme, suggesting that these mutations affect the catalytic site directly.⁹⁰ E coliPNPase crystals obtained in the presence of Mn²⁺showed that the cofactor is coordinated by the conserved residues Asp486, Asp492 and Lys494.⁵¹In addition, the Asp492 mutation eliminates both phosphorolytic and polymeric activity.⁹⁰

The α-helical domains connecting the first and second core domains are involved in the catalytic activityE coliPNPase,⁹¹although they are the least maintained domains.^25,90uE coliPNPase mutating adjacent α-helical domain residues appears to affect catalytic activity.⁹¹AnalyzeS.antibioticThe structure revealed that the α1 helix faces the putative phosphate binding site and thus may be part of the catalytic center.^11,91These results were combined with studies of spinach chloroplasts and human PNPases^92,93suggest a catalytic siteE coliThe PNPase likely consists of structural elements in the first and second core domains (Thread. 3).

On the other hand, RNA binding domains have a limited effect on phosphorolysis because PNPase mutants lacking either domain are catalytically active. Electrophoretic RNA mobility shift assays showed that both KH and S1 domains are required for proper binding⁹⁴and for PNPase autoregulation.⁹⁵Interestingly, the PNPase truncated in these two domains can still bind RNA, albeit with lower affinity, confirming that the catalytic core has intrinsic RNA-binding activity. In addition, the S1 and KH domains are required to release the substrate from the catalytic site.⁹⁶To explain this, a two-step model has been proposed based on the indirect facilitation of PNPase activity by the KH and S1 domains, allowing substrate binding and product release. The model assumes that there are two RNA-binding surfaces on each PNPase monomer: one in the catalytic core domain and the other formed by the KH and S1 domains. First, the ssRNA molecules are thought to weakly interact with one of the KH-S1 tandem domains. This bound ssRNA then interacts strongly with the catalytic site where it undergoes phosphorolysis until the stem loop is formed and the enzyme stops as the substrate remains bound to the binding surface of the RNA core. Finally, another molecule could bind to another empty KH-S1 tandem domain, migrate to the nuclear RNA binding site and allow the locked molecule to move. Therefore, the truncated ΔKH-S1 proteins are much less efficient in releasing products after catalysis.⁹⁶

fatty acid biosynthesis

Fatty acid synthases can be broadly divided into type I and type II enzymes. Type I synthases are multifunctional proteins in which the proteins catalyze individual partial reactions in separate domains and the protein acyl carrier is covalently bound to the protein. This type includes synthases from higher bacteria (Mycobacterium smegmatisICornybacteriumspp.) and yeast (S. cerevisiae). Type I synthases are proteins with a characteristic high molecular weight (0.4 × 10).⁶–2,5 × 10⁶), which consists of two or more large multifunctional polypeptide chains (molecular weight 1.8 × 10).⁵–2,7 × 10⁵). Typ IM. smegmatykis unusual in several respects; The two reductases have different specificities for reduced pyridine nucleotides, the β-ketoacyl-ACP reductase requiring NADPH and the enoyl-ACP reductase requiring NADH (other type I enzymes use only NADPH). In animals, the two chains are generally assumed to be identical. Recent genetic analyzes of yeast fatty acid synthase mutants have revealed that there are two unrelated polycistronic genes, designatedfas1 andfas2.Thefas1 gene encodes the enzymes acetyltransferase, malonyl (palmitoyl) transacylase, dehydratase and enoyl reductasefas2 encodes the binding region for phospho-pantothene and the enzymes β-ketoacyl synthase and reductase. The yeast synthase is probably A₆B₆A complex of two different multifunctional proteins (A and B). Type I yeast inhibits palmitoyl-CoA and this zAspergillus fumigatusinhibit malonyl-CoA.

Type II synthases contain enzymes that can be separated, purified, and studied separately, and are present in most lower bacteria (E coli) and the acyl carrier protein are easily separated from the enzyme. The type II synthase is the best studiedE coliseven proteins were isolated. It is assumed that in the cell, individual enzymes combine into a loosely bound multienzyme complex. The site of association may be the cell membrane, because inE coli,ACP is located in the membrane. In these reactions, substrates bind to ACP. The type II enzyme synthesizes both saturated and unsaturated fatty acids. This is due to the presence of releasing β-hydroxydecanoyl-ACP-β,γ-dehydrasecis-3-decenoyl-ACP (precursor of unsaturated fatty acids).trans-2-decenoyl-ACP, which is converted into a saturated fatty acid.

In yeast and fungi, the saturated fatty acids are palmitic acid (C₁₆) and stearic acid (C₁₈) serve as precursors to the monounsaturated fatty acids palmitoleic acid and oleic acid. The double bond is formed by the action of the fatty acyl-CoA oxygenase enzyme in an oxidation reaction. In the reaction, NADPH is oxidized to NADP⁺.