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Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information

The Escherichia coli stringent response, mediated by the alarmone ppGpp, is responsible for the reorganization of cellular transcription upon nutritional starvation and other stresses. These transcriptional changes occur mainly as a result of the direct effects of ppGpp and its partner transcription factor DksA on RNA polymerase. An often overlooked feature of the stringent response is the direct targeting of other proteins by ppGpp. Here we review the literature on proteins that are known to bind ppGpp and, based on sequence homology, X-ray crystal structures and in silico docking, we propose new potential protein binding targets for ppGpp. These proteins were found to fall into five main categories: (i) cellular GTPases, (ii) proteins involved in nucleotide metabolism, (iii) proteins involved in lipid metabolism, (iv) general metabolic proteins and (v) PLP-dependent basic aliphatic amino acid decarboxylases. Bioinformatic rationale is provided for expanding the role of ppGpp in regulating the activities of the cellular GTPases. Proteins involved in nucleotide and lipid metabolism and general metabolic proteins provide an interesting set of structurally varied stringent response targets. While the inhibition of some PLP-dependent decarboxylases by ppGpp suggests the existence of cross-talk between the acid stress and stringent response systems.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information

The Escherichia coli stringent response is a sophisticated and rapidly activated system which is induced in response to a number of nutritional or environmental stresses, and that mediates the transition between exponential and stationary phase growth (Cashel et al., 1996; Nystrom, 2004; Potrykus and Cashel, 2008). The stringent response effects are potentiated primarily through the unusual guanosine nucleotides: guanosine tetraphosphate ppGpp [guanosine 3′, 5′-bis(diphosphate)] and guanosine pentaphosphate pppGpp (guanosine 3′-diphosphate, 5′-triphosphate), collectively known as (p)ppGpp. In the cell, pppGpp is synthesized from GTP and ATP via the action of two paralogous enzymes RelA and SpoT (Cashel et al., 1996), which belong to a widely distributed family of RelA/SpoT homologue proteins (Atkinson et al., 2011). Subsequently, pppGpp is converted to ppGpp through the action of the pppGpp 5′-phosphohydrolase GppA enzyme (Fig. 1) (Hara and Sy, 1983). The protein domain boundaries and X-ray crystal structures of the N-terminal (p)ppGpp binding domains of both the RelA/SpoT homologue protein RelSeq from Streptococcus equisimilis (Hogg et al., 2004) and the GppA paralogue PPXAae from Aquifex aeolicus (Kristensen et al., 2008) are shown in Fig. 1A–D. RelA is associated with ribosomes through its C-terminus and is responsible for (p)ppGpp synthesis in response to amino acid limitation (Fig. 1E) (Wendrich et al., 2002). Recent single molecule studies have shown that alarmone synthesis occurs upon release of RelA from the ribosome during the stringent response (English et al., 2011). Cytoplasmic SpoT is responsible for the basal synthesis of (p)ppGpp during growth and for (p)ppGpp degradation (Gentry and Cashel, 1995). SpoT is also responsible for (p)ppGpp synthesis in response to a number of stress conditions (Cashel et al., 1996). Under fatty acid starvation conditions, and potentially under carbon-source starvation conditions, the acyl carrier protein (ACP) binds to and has been proposed to activate SpoT (Fig. 1E) (Battesti and Bouveret, 2006).

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Figure 1. Proteins involved in (p)ppGpp metabolism. A. Schematic of the domain boundaries of the Streptococcus equisimilis RelSeq (Hogg et al., 2004) and E. coli RelA and SpoT proteins (Metzger et al., 1989). The following domains are indicated: HD-type (p)ppGpp hydrolase domain (brown) (Aravind and Koonin, 1998); nucleotidyltransferase-type (p)ppGpp synthetase domain (green) (Hogg et al., 2004); TGS domain (purple) (a small domain found in threonine aminoacyl-tRNA synthetase ThrRS, GTPases and SpoT) (Wolf et al., 1999); and ACT domain (cyan) (aspartate kinase, chorismate mutase and TyrA domain) (Aravind and Koonin, 1999). B. Cartoon representation of the X-ray crystal structure (PDB ID: 1VJ7) of RelSeq HD and synthetase domains coloured as in (A). The helices coloured red form part of the conserved three-helix bundle that mediates communication between the HD and synthetase domains (Hogg et al., 2004). The bound GDP and ppG2′:3′p (a derivative of ppGpp) are illustrated as sticks with carbon atoms coloured purple, oxygen atoms coloured red, nitrogen atoms coloured blue, and phosphorous atoms coloured cyan. The bound Mn2+ ion is shown as a blue sphere. All X-ray structure images were generated using PyMOL (DeLano, 2002). C. The domain boundaries for the paralogous GppA and the polyphosphatases PPX from E. coli (Kuroda et al., 1997) and PPX from Aquifex aeolicus (PPXAqa) are shown. The PPX and GppA enzymes have a pppGpp 5′-phosphohydrolase activity (Hara and Sy, 1983; Kristensen et al., 2008). The following domains are indicated: ASKHA-I (orange) and ASKHA-II (blue) (acetate and sugar kinase/Hsp70/actin) superfamily domains (Reizer et al., 1993); the HD domain III (brown); and the C-terminal domain IV (DIV) (maroon) (Alvarado et al., 2006; Rangarajan et al., 2006). D. Cartoon representation of the X-ray crystal structure (PDB ID: 2J4R) of the PPXAae is shown with the domains coloured as indicated in (C) and the bound ppGpp shown as a stick figure and coloured as in (B). E. A schematic diagram showing the pathway for (p)ppGpp synthesis. Blue arrows indicate synthetic reactions and red arrows indicate degradative reactions. Activation of the ribosome-bound RelA is via amino acid starvation (Wendrich et al., 2002). SpoT is activated in response to a number of stresses including fatty acid starvation (Battesti and Bouveret, 2006), carbon source starvation (Xiao et al., 1991), diauxic shifts (Harshman and Yamazaki, 1971), phosphorous limitation (Spira et al., 1995; Spira and Yagil, 1998; Bougdour and Gottesman, 2007), iron limitation (Vinella et al., 2005), hyper-osmotic shock (Harshman and Yamazaki, 1972; Cashel et al., 1996), and oxidative stress (Chang et al., 2002). ACP senses fatty acid and potentially carbon source starvation and activates ppGpp production by SpoT.

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The stringent response was historically identified by the rapid downregulation of stable RNA (rRNA and tRNA) genes when cells grown in rich media encountered amino acid starvation (Stent and Brenner, 1961; Dennis et al., 2004). Subsequently, it was shown that the stringent response results in global genetic and physiological changes to cellular metabolism (Cashel et al., 1996; Nystrom, 2004; Magnusson et al., 2005; Potrykus et al., 2011; Traxler et al., 2011), listed in Table 1. As the master regulator of the stringent response, (p)ppGpp has two major categories of effects (i) modification of gene transcription and (ii) direct interaction with target proteins. The effects of (p)ppGpp on gene transcription has been extensively studied and reviewed (Cashel et al., 1996; Dennis et al., 2004; Nystrom, 2004; Magnusson et al., 2005; Potrykus and Cashel, 2008). In E. coli, alterations in gene expression profiles during the stringent response are the result of interactions between the RNA polymerase (RNAP), ppGpp, and a specific transcription factor DksA. During the stringent response, ppGpp and DksA are able to facilitate opposing effects on transcription: downregulation of highly expressed stable RNA (rRNA and tRNA) and cell proliferation genes and simultaneous upregulation of stress and starvation genes (Magnusson et al., 2005). A binding site for ppGpp was observed in a Thermus thermophilus RNAP-ppGpp co-crystal structure (Artsimovitch et al., 2004), but subsequent analysis of the E. coli RNAP has shown that this site is probably not responsible for mediating RNAP regulation by ppGpp (Kasai et al., 2006; Vrentas et al., 2008).

Table 1.  Summary of processes affected by the stringent response.
ProcessReferences
  1. Cellular processes that are downregulated or upregulated during the stringent response are shown.

Downregulated proliferative processes  
 Cell division Schreiber et al. (1991); Ferullo and Lovett (2008); Traxler et al. (2008)
 Cell motility (fimbriae and flagellar) Aberg et al. (2006); Magnusson et al. (2007)
 DNA replication Hernandez and Bremer (1993); Schreiber et al. (1995); Wang et al. (2007); Ferullo and Lovett (2008); Traxler et al. (2008)
 rRNA and tRNA synthesis Hernandez and Bremer (1993); Cashel et al. (1996); Traxler et al. (2008)
 Ribosome synthesis Cashel et al. (1996); Zhang et al. (2006); Lemke et al. (2011)
 Protein synthesis Svitil et al. (1993)
 Translation initiation and elongation Rojas et al. (1984); Cashel et al. (1996); Milon et al. (2006); Bremer and Dennis (2008)
 Nucleotide biosynthesis Hochstadt-Ozer and Cashel (1972); Fast and Skold (1977); Morton and Parsons (1977); Pao and Dyess (1981); Cashel et al. (1996); Traxler et al. (2008)
 Metabolite transport Hochstadt-Ozer and Cashel (1972); Hochstadt (1978)
 Phospholipid synthesis Merlie and Pizer (1973); Polakis et al. (1973); Lueking and Goldfine (1975); Heath et al. (1994)
 Oxidative metabolism Chang et al. (2002)
Upregulated stress response processes
 Amino acid biosynthesis Cashel et al. (1996); Tedin and Norel (2001); Barker et al. (2001b); Magnusson et al. (2005); Paul et al. (2005)
 σS synthesis Gentry et al. (1993); Chang et al. (2002)
 Universal stress protein synthesis Kvint et al. (2000); Gustavsson et al. (2002); Trautinger et al. (2005)
 Carbohydrate metabolism Dietzler and Leckie (1977); Traxler et al. (2006); 2008)
 Virulence gene expression Magnusson et al. (2005); Nakanishi et al. (2006)
 Toxin/antitoxin systems Chang et al. (2002)
 Antibiotic resistance Rodionov and Ishiguro (1995); Greenway and England (1999); Korch et al. (2003)
 Cyclopropane fatty acid synthesis Eichel et al. (1999)
 Chaperones and proteolysis systems Cashel et al. (1996); Chang et al. (2002); Yang and Ishiguro (2003)

While the global effects of the stringent response are mediated via changes in the transcription profile of the cell, there are a number of specific proteins that are directly targeted by ppGpp. Apart from the proteins that are involved in (p)ppGpp synthesis and degradation (RelA, SpoT, GppA) (Fig. 1) and the main target of (p)ppGpp regulation (RNAP), we identified five major categories of E. coli (p)ppGpp targets based on literature reports, bioinformatics, and in silico docking analysis: (i) cellular GTPases, (ii) proteins involved in nucleotide metabolism, (iii) proteins involved in lipid metabolism, (iv) general metabolic proteins and (v) the basic aliphatic amino acid decarboxylases.

GTPases

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information

The GTPase superfamily of proteins are found in all kingdoms of life, and in prokaryotes they function in translation, cell cycle regulation, protein translocation, and other essential but poorly characterized cellular functions (Caldon and March, 2003; Brown, 2005; Margus et al., 2007). The GTPases share a number of common GTP binding motifs that include: (G1) P-loop [GX4GK(S/T)] involved in binding 5′α- and 5′β-phosphates of GTP; (G2) conserved T involved in Mg2+ binding; (G3) Walker B [DX2G] involved in binding Mg2+ and the 5′γ-phosphate of GTP; (G4) [(N/T)(K/Q)XD] involved in binding the guanosine ring; and (G5) [poor consensus] involved in stabilizing G4 residues (Bourne et al., 1991; Brown, 2005). In many (but not all) cases, the GTPase activity cycle is regulated by two types of proteins: a GTP-bound protein is stimulated to hydrolyse GTP to GDP upon the binding of a GTPase activating protein (GAP); subsequently, the GDP is released upon interaction with a guanine nucleotide release protein (GNRP)/guanine nucleotide exchange factor (GEF) (Bourne et al., 1991; Caldon and March, 2003).

The E. coli GTPases can be divided into three major categories: the translation elongation-factor group (CysN, EFG, TypA/BipA, LepA, EF-Tu, RF3, SelB, IF2) (Margus et al., 2007), the cell-signalling and cell division Era/Obg group (Der/EngA, EngB, EngD, Era, HflX, MnmE/TrmE, Obg, YfjP, YkfA, RsgA) (Caldon and March, 2003), and the protein translocation FtsY/Ffh group (FtsY, Ffh) (Caldon and March, 2003). A number of other GTPases (FeoB, PurA) are also present but have not been categorized. Figure 2A shows the domain organization of the E. coli GTPases and Fig. 2B shows a sequence alignment of the GTP binding motifs and highlights the residue conservation. The strong amino acid conservation at the GTP binding site and the similarities in structure between GTP and ppGpp suggest that these proteins could bind ppGpp; there are reports of such interactions for the five GTPases discussed below.

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Figure 2. E. coli GTPases as direct targets of ppGpp action. A. Domain organization of the various E. coli GTPases is shown. Each protein contains a GTPase domain (coloured yellow) that has the conserved GTP binding motifs. Proteins that have been shown to interact with ppGpp are highlighted in purple. The following is a list of the UniProt (UniProt Consortium, 2011) accession numbers in parentheses for each protein followed where applicable by the full domain name(s). For the translation/elongation GTPases (CysN, EFG, TypA/BipA, LepA, EF-Tu, RF3, SelB), the GTPase domain is followed by two conserved domains: DII – domain II and DIII – domain III. CysN (P23845); EFG (P0A6M8) G′– GTPase prime insertion domain, DIV – domain IV, DV – domain V; TypA/BipA (P32132) DV – domain V, CTD – C-terminal domain; LepA (P60785) DV – domain V, CTD – C-terminal domain; EF-Tu (P0CE47); RF3 (P0A7I4), SelB (P14081); IF2 (P0A705) N1 – N-terminal domain 1, N2 – N-terminal domain 2, G1 – pre-GTPase domain, DII – domain II, C1 – penultimate C-terminal domain, C2 – C-terminal domain; Der/EngA (P0A6P5); EngB/YihA (P0A6P7); EngD (P0ABU2) TGS – ThrRS, GTPases, and SpoT domain; Era (P06616) KH – K-homology RNA binding domain; FeoB (P33650) HD – His-Asp metal binding domain; HflX (P25519); MnmE/TrmE (P25522) TrmE-N–N-terminal TrmE domain; Obg/CgtA (P42641) Obg-N – Obg-fold N-terminal domain; YfjP (P52131); YkfA (P75678); RsgA (P39286) RBD – OB-fold RNA binding domain, CTD – C-terminal domain; FtsY (P10121) NTD – N-terminal domain; Ffh (P0AGD7) NTD – N-terminal domain, CTD – C-terminal domain, and PurA (P0A7D4) ID – insertion domain. B. A multiple sequence alignment of the GTPase domains was determined using MUSCLE (Edgar, 2004) and the results were manually verified using JALVIEW (Clamp et al., 2004). The conserved GTPase-features including G1/P-loop, G3, G4 and G5 signature sequences and residues that make up these signature sequences are shown in bold. Highly conserved residues are coloured red and residues that vary between one of two predominant residues in a position are shown as green and blue. The protein names are abbreviated as in (A) except for Obg-Ec (E. coli Obg – P42641) and Obg-Bs (Bacillus subtilis Obg – P20964). C–F. Cartoon representation of the X-ray crystal structures of (C) EF-Tu bound to GDP (PDB ID: 1DG1) (Abel et al., 1996), (D) Obg from B. subtilis (ObgBs) bound to ppGpp (PDB ID: 1LNZ) (Buglino et al., 2002), (E) PurA bound to GDP (PDB ID: 1CIB) or (F) ppG2′:3′p (PDB ID: 1CH8) (Hou et al., 1999). In all cases, the GTPase domain is coloured yellow and the specific GTP binding motifs are indicated as follows: G1/P-loop – orange, G3– light blue, G4– pink, and G5– light green. The guanosine nucleotide is shown as a stick figure with oxygen atoms coloured red, nitrogen atoms coloured blue, phosphorous atoms coloured cyan, and carbon atoms coloured purple. Where applicable, the Mg2+ ion is shown as a green sphere.

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The translation elongation and initiation GTPases EFG, EF-Tu, and IF2 are large, multi-domain proteins that have homologous GTPase domains followed by a short β-barrel domain (DII) (Fig. 2A) (Margus et al., 2007). EFG GTPase activity powers the translocation of the ribosome during protein synthesis (al-Karadaghi et al., 1996). A complex between EFG and ppGpp has been postulated and ppGpp has been suggested to inhibit EFG activity (Table 2) (Rojas et al., 1984). The translation elongation factor EF-Tu is one of the most common proteins in the cell. GTP-bound EF-Tu ferries aminoacylated-tRNA to the A-site of the translating ribosome and, upon recognition of the correct codon/anticodon pair, EF-Tu is released from the tRNA by GTP hydrolysis (Song et al., 1999). The tightly bound EF-Tu-GDP complex is recycled via the action of the EF-Ts GNRP. The X-ray crystal structure of GDP-bound EF-Tu (Abel et al., 1996) is shown in Fig. 2C. EF-Tu can bind ppGpp alone (Legault et al., 1972; Miller et al., 1973; Hamel and Cashel, 1974; Rojas et al., 1984), in complex with aa-tRNA (Pingoud et al., 1983) or in complex with the EF-Ts (Rojas et al., 1984). It has been proposed that EF-Tu bound to ppGpp increases the translation fidelity during stress and starvation conditions (Rojas et al., 1984; Dix and Thompson, 1986).

Table 2.  Measured binding affinities of (p)ppGpp and other substrates to E. coli proteins. Thumbnail image of

Initiation factor 2 (IF2) is a GTPase that binds to the initiator fMet-tRNAfMet during assembly of the translating ribosome. IF2 is GTP-bound during active cell growth but its activity is inhibited by ppGpp binding under stress conditions (Legault et al., 1972). A nuclear magnetic resonance (NMR) solution structure of the GTPase domain of Bacillus stearothermophilus IF2 in complex with ppGpp has been reported (Milon et al., 2006) and ppGpp was found to bind at the same site as GTP/GDP. The binding of the 5′α- and 5′β-phosphates and the guanosine base are essentially the same for both GTP and ppGpp, while in the latter case the 3′α- and β-phosphates project away from the binding site and are thought to interfere with IF2-interacting partners (fMet-tRNAfMet or ribosomal proteins).

Obg also known as CgtA/YhbZ is an essential GTPase that may function in DNA replication, in ribosome assembly through interaction with the 50S ribosomal subunit, and in the stringent response by interacting with SpoT (Wout et al., 2004; Persky et al., 2009). Obg has a moderate affinity for GTP/GDP, a high exchange rate and a weak GTPase activity (Wout et al., 2004). In addition, the protein has been shown to bind (p)ppGpp and to influence the balance of pppGpp/ppGpp in the cell, suggesting that Obg functions as a pppGpp 5′-phosphohydrolase (Persky et al., 2009). A crystal structure of residues 1–342 of the Obg homologue from Bacillus subtilis (ObgBs) in complex with ppGpp has been determined (Buglino et al., 2002) and is shown in Fig. 2D. The binding of ppGpp was found to be dependent on the 5′α- and 5′β-phosphates and the guanosine base while the 3′α- and 3′β-phosphates are not directly coordinated and face away from the binding site.

The adenylosuccinate synthetase PurA is involved in de novo ATP biosynthesis and catalyses the following GTP-dependent reaction: inosine monophosphate +l-aspartate + GTP [LEFT RIGHT ARROW] adenylosuccinate + GDP + phosphate (Honzatko and Fromm, 1999). PurA activity can be inhibited by ppGpp (Table 2) (Gallant et al., 1971; Stayton and Fromm, 1979; Pao and Dyess, 1981) and co-crystal structures of GDP- and ppGpp-bound PurA have been determined (Honzatko and Fromm, 1999; Hou et al., 1999) (Fig. 2E and F). The alarmone was bound as, and potentially converted to, a ppG2′:3′p derivative of ppGpp in the GTP binding site, suggestive of a more complex inhibition mechanism (Hou et al., 1999).

To examine if the ppGpp binding interactions that have been observed for ObgBs and PurA (Fig. 2D and F) are structurally conserved, we performed in silico docking experiments with ppGpp and the X-ray crystal structures of the E. coli or homologous GTPases listed in Fig. 2A. For each model, ppGpp was docked onto the position of the natural substrate GTP/GDP or a substrate analogue (depending on their availability in the PDB file) and molecular dynamics simulations were performed to relax the system (Fig. S1). Additionally a measure of the thermodynamic favourability of the interaction between ppGpp and the target protein was calculated and in all cases a negative ΔG was obtained, indicating a favourable interaction (Tables S1 and S2). While the results of the docking experiments should be treated with caution as they are based on an in silico simulation, we can infer several useful trends related to the interaction between ppGpp and the target proteins. When ppGpp was docked into the GTPase active site, the guanosine base, ribose ring and 5′-phosphates occupy very similar positions to the natural substrate/product (GTP/GDP), while the 3′-phosphates tend to point away from the active site and, hence, away from the surface of the protein (Fig. S1). The 3′-phosphates generally do not make significant contacts with the protein, but where they do, they tend to be coordinated most often by residues from the X3 position of the G1 motif (mostly acidic or polar residues) (see Fig. 2B) and/or from G3 loop residues, as well as, from residues outside of the conserved GTPase motifs (see Table S3). The position of the 3′-phosphates directed away from the protein surface may sterically interfere with the binding of GEF and GNRF proteins that help to regulate GTPase activity.

ppGpp binding may also result in competitive inhibition with GTP/GDP. A comparison of the reported Ki values for ppGpp and Kd or KM values for GTP/GDP for the various GTPases (Table 2) indicates that all of the guanosine nucleotides bind with a similar range of affinities, mostly in the low micromolar to nanomolar range. It is significant that, where data are available for the same enzyme, ppGpp does not bind comparatively better than GTP/GDP. This suggests that any inhibitory effect on GTPase activity would be easily reversible and would likely only be significant when the intracellular concentration of ppGpp is very high, such as during the peak of the stringent response where ppGpp concentrations may reach millimolar amounts (Cashel et al., 1996). This is important as many of the GTPases are essential for cell growth (Caldon and March, 2003; Brown, 2005; Margus et al., 2007) and a reversible inhibition by ppGpp, resulting in a transient reduction in protein translation and translocation, increased translation fidelity and reduced cell division rates, would serve to complement the transcriptional effects of the stringent response that contribute to the shift from exponential to stationary phase growth.

Nucleotide and lipid metabolism

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information

During the stringent response one of the major effects is the downregulation of genes involved in nucleotide and lipid biosynthesis (Table 1) (Cashel et al., 1996; Traxler et al., 2008). In addition, there are direct effects of ppGpp on a number of enzymes involved in nucleotide and lipid metabolism, which are listed schematically in Fig. 3. These enzymes belong to different functional classes and are, therefore, likely to interact with ppGpp via different mechanisms. At present there are no co-crystal structures available for any of these enzymes bound to ppGpp, so a docking approach was used in order to gain some insights into these interactions. A common theme revealed by the docking experiments is the potential for ppGpp to bind at a nucleotide/nucleotide analogue binding site and to potentially act as a competitive inhibitor.

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Figure 3. Known and potential targets of direct (p)ppGpp action are involved in nucleotide and lipid metabolism. Domain organization of proteins involved in nucleotide and lipid metabolism that are either known to interact with ppGpp (labelled purple) or based on sequence homology are potential targets of ppGpp. For each protein, two columns of additional information are provided: (i) the general cellular function and (ii) the effect of (p)ppGpp on its activity. Proteins with demonstrated inhibition and activation in the presence of (p)ppGpp are indicated in bold and those with inferred inhibition are shown in brackets. The following is a list of the UniProt (UniProt Consortium, 2011) accession numbers in parentheses for each protein followed where applicable by the full domain name(s). DnaG (P0ABS5): ZBD – zinc binding domain, α/β subdomain, TOPRIM – topoisomerase/primase, 3HB – 3-helix bundle, DBD – DnaB-interacting domain; MazG (P0AEY3) NTD – N-terminal domain, CTD – C-terminal domain; GuaB (P0ADG7) TIM-barrel catalytic domain, CBS – tandem cystathione β-synthase domain, GuaC (P60560) TIM-barrel catalytic domain. The following proteins contain a type I phosphoribosyltransferase domain (PRT-I): Gpt (P0A9M5), Apt (P69503), Upp (P0A8F0), Hpt (P0A9M2), PyrE (P0A7E3), and PurF (P0AG16). PurF also has glutamine phosphoribosylpyrophosphate amidotransferase (GPATase) domain. HisG (P60757) contains a type IV phosphoribosyltransferase domain that consists of PBP-a, PBP-b – periplasmic binding protein domains and FDX – ferredoxin-like domain; PlsB (P0A7A7) LPLAT – lysophospholipid acyltransferase of glycerophospholipid biosynthesis; PgsA (P0ABF8) CDP-OH – cytosine diphosphate-alcohol phosphatidyltransferase; YnjF (P76226) CDP-OH – cytosine diphosphate-alcohol phosphatidyltransferase; AccA (P0ABD5) α/β spiral domain; AccD (P0A9Q5) ZBD – zinc binding domain, α/β spiral domain; FabA (P0A6Q3) α + β hot-dog domain; FabZ (P0A6Q6) α + β hot-dog domain; GdhA (P00370) DI – domain I, DII – domain II; GlgC (P0A6V1) ADP-G-PP – ADP-glucose-pyrophosphorylase domain, AT – glucose-1-phosphate adenylyltransferase; Ppc (P00864) PEPC – phosphoenolpyruvate carboxylase domain.

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DNA primase (DnaG) is a multi-domain enzyme that is a component of the replisome and interacts with the DnaB helicase, single stranded binding protein, and DNA polymerase III. Direct inhibition of DnaG by pppGpp was first observed in B. subtilis where pppGpp was found to be more inhibitory than ppGpp (Table 2) (Wang et al., 2007). A subsequent study of E. coli DnaG indicated that, in the presence of DnaB, ppGpp was a more potent inhibitor than pppGpp (Table 2) (Maciag et al., 2010). DnaG was inhibited by concentrations of (p)ppGpp in the 0.2–1.0 mM range, which are readily reached during the stringent response (Cashel et al., 1996; Buckstein et al., 2007; Traxler et al., 2008). ppGpp was successfully docked into the proposed nucleotide binding site (Keck et al., 2000) (Fig. S2A) at the interface between the α/β and topoisomerase/primase domains (Fig. 3) and, thus, may function by competitive inhibition. Inhibition of DNA primase would serve to strongly reduce the rate of de novo DNA synthesis and, hence, of cell division.

The nucleotide pyrophosphohydrolase MazG functions in regulating programmed cell death in E. coli and is negatively regulated by the MazEF toxin-antitoxin system (Lee et al., 2008). MazG has low levels of ppGpp pyrophosphohydrolase activity in vitro, but this activity is insufficient to complement a deletion of spoT, which causes a toxic accumulation of ppGpp (Xiao et al., 1991). ppGpp was docked at the ATP binding pocket of MazG (Fig. S2B).

The first step in the guanosine nucleotide de novo biosynthesis pathway is catalysed by inosine monophosphate dehydrogenase (GuaB). GuaB activity is inhibited by ppGpp with a Ki of ∼ 50 µM, indicating that GuaB will be efficiently inhibited during the stringent response (Table 2) (Gallant et al., 1971; Gilbert et al., 1979; Pao and Dyess, 1981). Inhibition of GuaB results in decreased pools of GTP and this has been implicated in mediating effects of the stringent response including induction of sporulation in B. subtilis (Ochi et al., 1982) and modulating RNAP activity in T. thermophilus (Kasai et al., 2006). A homologous enzyme guanosine-5′-monophosphate dehydrogenase (GuaC), which is involved in the purine salvage pathway, may also be a target of (p)ppGpp as the enzymes share a common TIM-barrel fold (Andrews and Guest, 1988). (p)ppGpp may act as a competitive inhibitor by binding to the guanosine binding site present in these enzymes, which is observed in the docked models of ppGpp with GuaB (Fig. S2C) and GuaC (Fig. S2D).

A number of phosphoribosyltransferases (PRTases) have been implicated in the stringent response. There are four major categories of PRTases: class I to IV (Lohkamp et al., 2004). Class I PRTases (PRT-I) contain a unique conserved motif [VL(IVL)VDDX4G] that is involved in binding to phosphoribosylpyrophosphate (PRPP). Four PRT-I enzymes involved in purine and pyrimidine salvage pathways: xanthine-guanine PRTase (Gpt), adenine PRTase (Apt), uracil PRTase (Upp) and hypoxanthine PRTase (Hpt) are inhibited by (p)ppGpp (Hochstadt-Ozer and Cashel, 1972; Fast and Skold, 1977; Morton and Parsons, 1977). There are two other E. coli PRT-I enzymes that may also be inhibited by (p)ppGpp: orotate PRTase (PyrE), and glutamine phosphoribosylpyrophosphate amidotransferase (PurF). We have docked ppGpp at the active sites of the various PRT-I enzymes (Fig. S2E–J) and the alarmone seems to adopt a conformation similar to the expected transition state intermediate formed between PRPP and the purine/pyrimidine base (see Fig. S3). The KM values for the various substrates and PRPP for these enzymes are generally within the low to moderate micromolar range and where available, measures of ppGpp binding are either of similar or lower affinity to the bona fide substrates (Table 2). The class IV PRTase HisG (ATP PRTase), which catalyses the first step in histidine biosynthesis, is also inhibited by the activity of (p)ppGpp (Morton and Parsons, 1977). This protein has a different domain architecture from the PRT-I enzymes (Fig. 3) but does contain PRPP and ATP binding sites either of which may be possible targets for (p)ppGpp binding and inhibition (Fig. S2K). There is both in vivo (Barker et al., 2001a) and in vitro (Paul et al., 2005) evidence that the PhisG promoter is upregulated by (p)ppGpp and DksA during the stringent response, so it is important to determine the extent of (p)ppGpp-based inhibition of this enzyme.

The enzyme responsible for the first step of lipid biosynthesis is the membrane-bound glycerol-3-phosphate acyltransfersae (PlsB) (Fig. 3). PlsB activity is directly inhibited by low millimolar quantities of (p)ppGpp (Merlie and Pizer, 1973; Heath et al., 1994). As there is no current structure for this integral membrane protein, predicting the site of action of (p)ppGpp is not currently feasible. A down-stream enzyme that catalyses the first step of phospholipid biosynthesis is phosphatidylglycerophosphate synthase (PgsA) (Fig. 3) and this membrane protein is similarly inhibited by (p)ppGpp (Merlie and Pizer, 1973). A homologous predicted membrane protein YnjF (Fig. 3) is also present in E. coli and while the function of this protein is currently not known with certainty, it is expected to function as a phosphatidyl transferase and may also be inhibited by (p)ppGpp.

A key enzyme in the bacterial type-II fatty acid biosynthesis (FAS-II) pathway is the acetyl-CoA carboxylase complex that consists of three separate protein complexes: the dimeric biotinoyl carboxyl carrier protein (BCCP) (AccB)2; dimeric biotin carboxylase (BC) (AccC)2 and the heterotetrameric acetyl-CoA carboxytransferase (CT) (AccA)2(AccD)2 (Bilder et al., 2006). BC catalyses the ATP-dependent addition of HCO3- to BCCP-biotin, generating BCCP-biotin-CO2. The carbonyl group is subsequently transferred to acetyl-CoA to generate malonyl-CoA via CT. The AccA and AccD proteins that make up the CT complex are homologous and the CT complex is inhibited by low millimolar concentrations of (p)ppGpp (Polakis et al., 1973). Two other FAS-II enzymes involved in the formation of unsaturated fatty acids are also inhibited by low millimolar concentrations (p)ppGpp (Stein and Bloch, 1976): the homologous FabA and FabZ proteins that contain a unique α + β hot-dog topology and catalyse β-hydroxyacyl-ACP dehydratase reactions (Leesong et al., 1996).

A number of other metabolic enzymes are also regulated by (p)ppGpp and include the NADP+-dependent glutamate dehydrogenase (GdhA) that is inhibited by (p)ppGpp (Maurizi and Rasulova, 2002), the first enzyme in the glycogen biosynthesis pathway glucose-1-phosphate adenylyltransferase (GlgC) that is inhibited by low millimolar concentrations of (p)ppGpp (Dietzler and Leckie, 1977), and the metabolic enzyme phosphoenolpyruvate carboxylase (Ppc) that, uniquely among the enzymes considered, is activated by (p)ppGpp (Pao and Dyess, 1981). Interestingly, GlgC transcription is activated by ppGpp (Romeo and Preiss, 1989) and translation is blocked through the action of the carbon storage regulator CsrA (Romeo et al., 1993), suggesting a complex regulatory network and potentially opposing effects of ppGpp for this enzyme.

Docking of ppGpp to AccA, FabA, GdhA, GlgC and Ppc was performed (Fig. S4A–E) but the docking procedure was more challenging in several of the cases (AccA, FabA, Ppc) where a suitable substrate on which to model ppGpp was unavailable. In order to deal with this, the DOCK6 (Lang et al., 2009) software package was used to obtain suitable docking conformations. In addition, all the enzymes are from different structural classes, thus further complicating a search for a common mechanism of action of ppGpp. More accurate measurements of the actual affinity of ppGpp for these enzymes would be necessary to determine the extent of direct inhibition during the stringent response.

Basic aliphatic amino acid decarboxylases

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information

Escherichia coli possesses five enzymes belonging to the prokaryotic ornithine decarboxylase (pODC) subclass of Fold Type I pyridoxal-5′-phosphate (PLP)-dependent decarboxylases: inducible lysine decarboxylase (LdcI), constitutive lysine decarboxylase (LdcC), inducible arginine decarboxylase (AdiA), inducible ornithine decarboxylase (SpeF) and constitutive ornithine decarboxylase (SpeC) (Fig. 4A) (Kanjee et al., 2011a). These multidomain enzymes form large oligomeric complexes consisting of dimers (SpeF, SpeC) or decamers (LdcI, LdcC, AdiA). The inducible enzymes are involved in the acid stress response while the constitutive enzymes, particularly SpeC, are important in polyamine production.

image

Figure 4. Regulation of lysine and ornithine decarboxylases by (p)ppGpp. A. The domain organization of the five related PLP-dependent basic aliphatic amino acid decarboxylases is shown along with the effects of GTP/GPD and (p)ppGpp on enzyme activities (Kanjee et al., 2011a). Each of the decarboxylases shares a common domain architecture consisting of: an N-terminal Wing domain; a Core domain made up of a short α-helical linker, a PLP binding subdomain (PLP-SD) and a subdomain four/aspartate aminotransferase small domain (SD4); and a C-terminal domain (CTD). The following is a list of the UniProt (UniProt Consortium, 2011) accession numbers in parentheses for each protein: LdcI (P0A9H3), LdcC (P52095), AdiA (P28629), SpeF (P24169) and SpeC (P21169). B. X-ray crystal structure of the E. coli LdcI decamer (PDB ID: 3N75) (Kanjee et al., 2011b) with each monomer in the top ring highlighted in a different colour and shown as a cartoon. The bottom ring monomers are shown in surface representation. The five ppGpp molecules that interact with the top ring are indicated. The insert shows a close-up of one of the ppGpp binding sites. The guanosine nucleotide is shown as a stick figure with oxygen atoms coloured red, nitrogen atoms coloured blue, phosphorous atoms coloured cyan, and carbon atoms coloured purple.

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In the recently determined X-ray crystal structure of LdcI by our group, the enzyme was found to bind ppGpp with high affinity (Table 2) at specific sites between neighbouring monomers in the LdcI decamer (Fig. 4B) (Kanjee et al., 2011b). Furthermore, it was found that LdcI activity was specifically inhibited by ppGpp and pppGpp over a range of pH values in vitro and in vivo. Of the related E. coli decarboxylases, it was found that LdcC was similarly inhibited by (p)ppGpp (Kanjee et al., 2011a). SpeF and SpeC were both activated by GTP and GDP, while SpeC was inhibited by (p)ppGpp. The arginine decarboxylase AdiA was unaffected by any of the guanosine nucleotides (Fig. 4A). Inhibition of these decarboxylases by ppGpp likely results in the conservation of amino acids when the stringent response is activated under acid stress conditions and, thus, serves as an additional means of regulating decarboxylation activity (Kanjee et al., 2011a). Further experimentation is required to elucidate the exact mechanism of inhibition by (p)ppGpp and to elucidate the wider effects this inhibition has on cellular adaptation to acid stress.

Conclusion

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information

DNA microarray experiments comparing wild-type and either ΔrelA or ppGpp0 strains have extended our understanding of the global transcriptional changes that take place upon the induction of the stringent response (Chang et al., 2002; Traxler et al., 2006; 2008; 2011; Durfee et al., 2008). Direct interaction of proteins with ppGpp provides a central regulatory framework for many different types of processes, and this is exemplified by the transcriptional effects of ppGpp (and DksA) on RNAP. These transcriptional effects serve to manage a core set of genes that are involved in the shift between exponential phase growth conditions and stationary phase stress response conditions (Nystrom, 2004).

Based on the reported inhibition constants for ppGpp (Table 2) it is likely that, in the majority of cases, inhibition by ppGpp is transient, reversible and dependent on the high concentrations of ppGpp reached during the peak of the stringent response. The potential consequences on cell physiology of inhibition of these proteins by ppGpp are likely to be complex but may serve to complement the transcriptional effects of ppGpp on RNAP. Inhibition of the cellular GTPases may result in an overall decrease in protein translation and cell growth rates. Inhibition of enzymes involved in nucleotide and lipid metabolism is also consistent with reduction in the cell division rates as there is a reduced demand for producing nucleotides for DNA replication and stable RNA transcription and lower need for lipids to form new membranes. Similarly inhibition by ppGpp of certain metabolic enzymes and the amino acid decarboxylases would serve to conserve nutrients and amino acids during conditions of nutrient deprivation. Direct targeting of enzymes by (p)ppGpp may have evolved as a mechanism to specifically extend stringent control to and enable a rapid and reversible control of metabolic and stress response processes and help to fine-tune the effects of the stringent response.

In order to identify the direct targets of ppGpp, we have used reports from the existing literature as well as bioinformatic approaches (sequence alignments and in silico docking) in order to compile a list of proteins that are known or speculated to be regulated by the alarmone. While we have made every attempt to be rigorous in our analysis and selection of protein targets, we cannot be certain that all of these proteins are bona fide targets. As such, further structural and biochemical investigations into the known and proposed enzymes targeted by (p)ppGpp are essential to more completely define the role of this unusual nucleotide in regulating the stringent response.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information

The docking calculations were performed using the RIKEN Integrated Cluster of Clusters (RICC) facility. U. K. is the recipient of a National Sciences and Engineering Research Council of Canada (NSERC) Postgraduate Scholarship, a Fellowship of the Canadian Institutes of Health Research (CIHR) Strategic Training Program in the Structural Biology of Membrane Proteins Linked to Disease, and a University of Toronto Open Fellowship. This work was supported by a grant from CIHR (MOP-67210) to W. A. H.

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  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. GTPases
  5. Nucleotide and lipid metabolism
  6. Basic aliphatic amino acid decarboxylases
  7. Conclusion
  8. Acknowledgements
  9. References
  10. Supporting Information
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MMI_8177_sm_SuppInfor.pdf62082KSupporting info item
MMI_8177_sm_TS3.xlsx34KSupporting info item

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