Functional Characterization of Small Alarmone Synthetase and Small Alarmone Hydrolase Proteins from Treponema denticola

ABSTRACT

The stringent response enables bacteria to survive nutrient starvation, antibiotic challenge, and other threats to cellular survival. Two alarmone (magic spot) second messengers, guanosine pentaphosphate (pppGpp) and guanosine tetraphosphate (ppGpp), which are synthesized by RelA/SpoT homologue (RSH) proteins, play central roles in the stringent response. The pathogenic oral spirochete bacterium Treponema denticola lacks a long-RSH homologue but encodes putative small alarmone synthetase (Tde-SAS, TDE1711) and small alarmone hydrolase (Tde-SAH, TDE1690) proteins. Here, we characterize the respective in vitro and in vivo activities of Tde-SAS and Tde-SAH, which respectively belong to the previously uncharacterized RSH families DsRel and ActSpo2. The tetrameric 410-amino acid (aa) Tde-SAS protein preferentially synthesizes ppGpp over pppGpp and a third alarmone, pGpp. Unlike RelQ homologues, alarmones do not allosterically stimulate the synthetic activities of Tde-SAS. The ~180 aa C-terminal tetratricopeptide repeat (TPR) domain of Tde-SAS acts as a brake on the alarmone synthesis activities of the ~220-aa N-terminal catalytic domain. Tde-SAS also synthesizes “alarmone-like” nucleotides such as adenosine tetraphosphate (ppApp), albeit at considerably lower rates. The 210-aa Tde-SAH protein efficiently hydrolyzes all guanosine and adenosine-based alarmones in a Mn(II) ion-dependent manner. Using a growth assays with a ΔrelAΔspoT strain of Escherichia coli that is deficient in pppGpp/ppGpp synthesis, we demonstrate that Tde-SAS can synthesize alarmones in vivo to restore growth in minimal media. Taken together, our results add to our holistic understanding of alarmone metabolism across diverse bacterial species.

IMPORTANCE The spirochete bacterium Treponema denticola is a common component of the oral microbiota. However, it may play important pathological roles in multispecies oral infectious diseases such as periodontitis: a severe and destructive form of gum disease, which is a major cause of tooth loss in adults. The operation of the stringent response, a highly conserved survival mechanism, is known to help many bacterial species cause persistent or virulent infections. By characterizing the biochemical functions of the proteins putatively responsible for the stringent response in T. denticola, we may gain molecular insight into how this bacterium can survive within harsh oral environments and promote infection. Our results also expand our general understanding of proteins that synthesize nucleotide-based intracellular signaling molecules in bacteria.

INTRODUCTION

The “stringent response” is a coordinated set of physiological processes that bacterial cells initiate when they encounter adverse conditions such as nutrient depletion, osmotic shock, or pH extremes. During the stringent response, cellular growth and proliferation are inhibited, and physiological processes that promote cellular survival and persistence are upregulated (reviewed in references 1–8). Two phosphorylated guanine nucleotides, guanosine pentaphosphate (GTP, 3′-diphosphate, pppGpp) and guanosine tetraphosphate (guanosine 3′,5′-bisdiphosphate, ppGpp), are synthesized to millimolar levels during the stringent response (9–13). Colloquially known as “alarmones” or “magic spots,” (p)ppGpp nucleotides function as intracellular second messengers that alter cellular activities at the transcriptional and translational levels, as well as by the direct modulation of enzymatic activities (1–8). More recently, a third alarmone molecule, guanosine 3′-diphosphate, 5-phosphate (pGpp), has also been shown to function as an alarmone in cells (14, 15).

Alarmones are synthesized by RelA/SpoT Homologue (RSH) family proteins (16, 17). Bacteria primarily use multidomain long-RSH (long-Rel) proteins ~700 amino acids (aa) in length to synthesize and/or degrade alarmones. These comprise an N-terminal domain containing SYNTH (synthesis) and HD (His-Asp motif; hydrolysis) subdomains and a C-terminal regulatory domain (16–20). The SYNTH domain catalyzes (p)ppGpp synthesis via the transfer of a diphosphate (PPi, pyrophosphate) group from ATP to the ribose 3′-OH group of GTP/GDP, with the concomitant production of AMP. The HD domain catalyzes (p)ppGpp degradation by hydrolytically removing the 3′-diphosphate group to regenerate GTP/GDP (21–27).

In addition to long-RSH, many bacteria encode monofunctional small alarmone synthetase (SAS) or small alarmone hydrolase (SAH) proteins (16). SASs contain a SYNTH domain, catalyze only alarmone synthesis, and are typically ~200 to 230 aa in length (28–36). SAHs contain an HD domain, are typically 180 to 210 aa in length, and catalyze alarmone hydrolysis (37–39).

The best studied SASs are the respective RelQ and RelP proteins from Staphylococcus aureus, Bacillus subtilis, Streptococcus mutans, and Enterococcus faecalis (RelQ only) (28, 29, 34, 40–42). Other identified SAS lineages include RelV from Vibrio cholerae (30, 43), a dual function RNase HII-(p)ppGpp synthetase from Mycobacterium smegmatis (MS_RHII-RSD) (44, 45), and RelS (Cg2324) from Corynebacterium glutamicum (36). Certain RelQ, RelP, and RelS protein homologues are capable of synthesizing pGpp, albeit to different extents (1, 14, 15, 35, 40, 45, 46).

Recently, certain SAS homologues were shown to synthesize the “alarmone-like” molecules adenosine pentaphosphate (ATP, 3′-diphosphate; pppApp) and adenosine tetraphosphate (adenosine 3′,5′-bisdiphosphate; ppApp). These “toxigenic” SAS (Tox-SAS) proteins and their (p)ppApp products elicited lethal or highly cytotoxic effects within affected cells (1, 31, 47, 48). These Tox-SAS homologues were commonly encoded in toxin-antitoxin (TA) modules alongside a cognate “antitoxin” SAH protein, which had the ability to hydrolyze (p)ppApp to ATP/ADP (31, 38). Certain long-RSH homologues also possess the ability to directly synthesize and/or hydrolyze (p)ppApp (49, 50).

The anaerobic spirochete bacterium Treponema denticola is strongly associated with the development of periodontitis, a chronic infectious-inflammatory disease that affects the gum tissues and underlying tooth-supporting structures (51–55). Periodontitis affects hundreds of millions of people worldwide, and it is the leading cause of tooth loss in adult populations (56). T. denticola is one of ~70 species/species-level phylotypes of Treponema taxa resident in the oral cavity (57, 58).

T. denticola is highly unusual in the bacterial kingdom, as it lacks a long-RSH homologue but encodes a putative SAS homologue (Tde-SAS, TDE1711; 410 aa) and a putative SAH homologue (Tde-SAH, TDE1690; 205 aa) (16, 59). Here, we show that Tde-SAS optimally catalyzes the synthesis of ppGpp over pppGpp and pGpp but also has lower-level ppApp synthesizing activities. We reveal that the presence of the C-terminal TPR motif domain of Tde-SAS represses the synthetic activities of the N-terminal catalytic domain. We further demonstrate that Tde-SAH efficiently hydrolyzes (pp)pGpp and (p)ppApp to the corresponding guanosine- or adenosine-based nucleotides, thus putatively functioning to remove the alarmone (alarmone-like) nucleotides synthesized by Tde-SAS.

RESULTS

Domain structure of Tde-SAS.

The 410-aa Tde-SAS (TDE1711) protein has a distinctive two-domain structure. The N-terminal ~220-aa region shares notable sequence similarity with RelP, RelQ, and RelS (ActRel) homologues and contains all the conserved amino acid residues and structural motifs (Syn1 to Syn4) predicted to be required for alarmone synthesis (Fig. 1A) (27, 41, 42). The C-terminal ~180-aa region contains four tetratricopeptide repeat (TPR) motifs, each of which are predicted to adopt a typical helix-turn-helix conformation (60). Analysis using the TPRpred webserver (61) indicates that these four TPR motifs (TPR1 to TPR4) correspond to residues 255 to 288 (TPR1), 292 to 325 (TPR2), 326 to 359 (TPR3), and 360 to 393 (TPR4), respectively (Fig. 1B and 2). The predicted structural arrangement of the four TPR motifs is shown in Fig. 1C. From here on, we refer to the N-terminal ~220-aa region as the “catalytic domain” and the C-terminal ~180-aa region as the “TPR domain.”

FIG 1.

Domain structure and conserved motifs within Treponema denticola encoding putative small alarmone synthetase (Tde-SAS). (A) Schematic diagram showing the domain structure and conserved motifs present within Tde-SAS and Tde-SAS_1–246 (N-terminal catalytic domain). (B) Sequence alignment of the four tetratricopeptide repeat (TPR) motifs (TPR1 to TPR4) in the C-terminal TPR domain of Tde-SAS, indicating their respective start and end positions, as identified by TPRpred (61). (C) Predicted structural arrangement of the four TPR motifs within the Tde-SAS TPR domain (prepared using SwissModel [90]). The tandem array of four TPR motifs generates a right-handed superhelical structure, with the region immediately C-terminal to TPR4 (colored dark orange/red) forming an extended α-helix. (D) Multiple sequence alignment of the catalytic domain of Tde-SAS (residues 1 to 228 shown) with diverse SAS proteins: C. glutamicum RelS (RelS_Cg; CgRelS), S. aureus RelP/SAS2 (SaRelP), B. subtilis RelP/Ywac/SAS2 (BsRelP), S. aureus RelQ/SAS1 (SaRelQ), and B. subtilis RelQ/YjbM/SAS1 (BsRelQ). The four respective “synthesis” Syn motifs (Syn1 to Syn4) (27) are indicated with brackets. The figure was prepared using ESPript 3.0 (88).

FIG 2.

Phylogeny of SAS-TPR (DsRel) homologues and alignment of treponeme SAS-TPR protein sequences. (A) Maximum likelihood (ML) phylogenetic tree of small alarmone synthetase (SAS) homologues, with the major families (phylogenetic clades) indicated. The respective phylogenetic positions of Tde-SAS and other SAS-TPR homologues (DsRel2 family members) from the genus Treponema (shaded violet), from other spirochetes (shaded pink), and from nonspirochete taxa (shaded orange) are indicated. (B) Multiple sequence alignment of Tde-SAS with representative SAS-TPR (DsRel) homologues from Treponema vincentii F0403 (TvinSAS, 409 aa, WP_016518062), Treponema socranskii subsp. socranskii ATCC 35536^T (TsocSAS, 413 aa, WP_021330562), and Treponema pectinovorum ATCC 700769^T (TpecSAS, 415 aa, WP_147612650). The locations of the four synthesis motifs (Syn1 to Syn4) and four TPR motifs (TPR1 to TPR4) are indicated with brackets. The putative linker region is highlighted with a blue box. The C terminus of the Tde-SAS_1–246 protein is highlighted with an arrow. The SAS/small alarmone hydrolase (SAH) classification system used here was defined as in reference 31. The alignment figure was prepared using ESPript 3.0 (88). The phylograms were prepared using ITOL (94).

Tde-SAS belongs to the DsRel family of RSH proteins (31), all of which comprise an N-terminal catalytic domain and C-terminal TPR domain. Correspondingly, we refer to these two-domain DsRel-family proteins as “SAS-TPR” homologues (Fig. 2A). SAS-TPR homologues encoded by treponemes (Treponema spp.) share high levels of sequence similarity (Fig. 2B) and form a phylogenetic clade that is distinct from that of SAS-TPR homologues encoded other spirochete taxa or by Deltaproteobacteria (Fig. 2A).

Biophysical characterization of Tde-SAS.

Recombinant Tde-SAS (molecular weight [MW], 51,318 Da), migrated with an apparent MW of ~220 kDa upon size-exclusion chromatography (SEC) analysis, indicating it formed a stable homotetramer in solution (Fig. S2A). A recombinant protein that comprised only the catalytic domain (Tde-SAS_1–246; residues 1 to 246; MW, 32,557 Da) migrated with an apparent MW of ~160 kDa upon SEC analysis, suggesting it similarly adopted a homotetrameric arrangement (Fig. S2B). The purified Tde-SAS and Tde-SAS_1–246 proteins had A_260nm/A_280nm ratios of 0.71 ± 0.12 and 0.53 ± 0.11, respectively. This indicated that neither protein (obtained by heterologous expression in Escherichia coli) was complexed to RNA, as has previously been observed for long-RSH proteins (62).

The domain structure of Tde-SAS was probed using limited proteolysis with the nonspecific protease subtilisin (18, 63). Two major protein fragments were formed that had apparent MWs of ~26 kDa and ~21 kDa (Fig. S2D). Peptide mass fingerprint (PMF) analysis revealed that these protein fragments respectively corresponded to the catalytic domain and the TPR domain (data not shown). This indicated that subtilisin had primarily cleaved the Tde-SAS protein within the interdomain “linker” region comprising residues ~230 to 250 (Fig. 2B).

Tde-SAS preferentially synthesizes ppGpp, and its catalytic activities are not notably modulated by alarmones.

The results from initial sets of biochemical assays revealed that Tde-SAS catalyzed the synthesis of pppGpp, ppGpp, and pGpp from ATP + GTP, ATP + GDP, and ATP + GMP, respectively, with the concomitant production of AMP. Representative chromatograms of enzymatic product mixtures are shown in Fig. 3A. The optimal pH for ppGpp synthesis activities was approximately 8.8 to 9.2 (Fig. S3A). The specific molar activities of Tde-SAS were determined under standardized conditions to quantify the respective rates of pppGpp, ppGpp, and pGpp synthesis. This was defined in units of micromoles of (pp)pGpp product synthesized per minute per micromole of Tde-SAS protein (μmol · min⁻¹ · μmol⁻¹) based on monomeric protein concentrations, as previously described (40). The rate of ppGpp synthesis was 689 ± 87 μmol · min⁻¹ · μmol⁻¹, which was ~22-fold faster than the rate of pGpp synthesis (30.9 ± 3.6 μmol · min⁻¹ · μmol⁻¹) and ~88-fold faster than the rate of pppGpp synthesis (7.8 ± 1.6 μmol · min⁻¹ · μmol⁻¹) (Fig. 3B).

FIG 3.

Synthesis of (pp)pGpp by Tde-SAS and Tde-SAS_1–246. (A) Representative Mono Q anion-exchange chromatograms of enzymatic product mixtures showing the respective (pp)pGpp synthesis activities of Tde-SAS (blue) and Tde-SAS_1–246 (red) upon incubation with ATP + GMP (left), ATP + GDP (center) or ATP + GTP (right). All sets of assays were performed under analogous conditions. (B) Specific molar activities of the Tde-SAS and Tde-SAS_1–246 proteins for the synthesis of pGpp (ATP + GMP), ppGpp (ATP + GDP) and pppGpp (ATP + GTP). These rates are reported in the following units: micromoles of (pp)pGpp synthesized per minute per micromole of protein (μmol · min⁻¹ · μmol⁻¹). All reactions were performed in triplicate, reporting the means ± standard deviation. See Materials and Methods for experimental details.

Analogous sets of assays were performed for the Tde-SAS_1–246 protein under identical conditions. Chromatographic analysis of enzymatic product mixtures indicated that Tde-SAS_1–246 had catalytic activities that were equivalent to those of Tde-SAS but were more potent (Fig. 3A). The respective specific molar rates of pppGpp, ppGpp, and pGpp synthesis by Tde SAS_1–246 were subsequently determined to be ~3- to 10-fold faster than those of Tde-SAS (Fig. 3B). Both Tde-SAS_1–246 and Tde-SAS synthesized ppGpp considerably faster than pGpp or pppGpp, suggesting that their substrate utilization patterns were equivalent (i.e., GDP ≫ GMP > GTP).

Micromolar concentrations of Zn²⁺ ions were previously shown to enhance the alarmone-synthesizing activities of Sa-RelP, while Fe²⁺/Fe³⁺ ions played a putative structural role (42). Therefore, we assayed the ppGpp-synthesizing activities of Tde-SAS_1–246 in the presence of supplementary Fe³⁺ or Zn²⁺ ions (over the concentration range 0 to 32 μM). The results clearly indicated that neither of these metal ions had significant effects on the rate of ppGpp synthesis by Tde-SAS_1–246 (Fig. S4).

The addition of alarmones have previously been shown to modulate the synthetic activities of RelQ- and RelP-family SAS proteins (14, 40–42, 64). Therefore, the Michaelis-Menten kinetic parameters for ppGpp synthesis were determined for Tde-SAS in the absence of alarmone, as well as in the presence of 100 μM pppGpp or ppGpp (Fig. 4B). In the absence of (p)ppGpp, the K_m(GDP) value was 5.29 ± 2.61 mM, with a corresponding k_cat value (turnover number) of 13.77 ± 4.13 s⁻¹, giving a catalytic efficiency (k_cat/K_m) value of 2.60 ± 0.51 mM⁻¹ s⁻¹. The Hill coefficient (h) was 1.35 ± 0.37, indicating that there may be low levels of positive cooperativity. (Note that h > 1 indicates product-mediated stimulation of enzymatic activities, also known as positive cooperativity or feed-forward control.) When 100 μM ppGpp or pppGpp was added, the corresponding K_m(GDP) values increased slightly to 6.59 ± 2.80 and 6.03 ± 2.28 mM, respectively. There was little effect on the corresponding turnover numbers, catalytic efficiencies, or h coefficients. These data indicated that added (p)ppGpp had slight stimulatory effects on the catalytic activities of Tde-SAS at a concentration of 100 μM. Additional sets of assays containing 200 μM (allosteric) ppGpp or pppGpp led to only ~10% enhancements in rates of alarmone synthesis (Fig. S5).

FIG 4.

Enzymatic kinetic parameters for (pp)pGpp synthesis by Tde-SAS and Tde-SAS_1–246 with/without addition of (p)ppGpp. (A) Plots of initial reaction velocity (V₀) versus (GMP, GDP, and GTP) substrate concentration ([S]) used to calculate the kinetic parameters for pGpp (left), ppGpp (center), and pppGpp (right) synthesis by Tde-SAS. The y axes show the reaction velocity in units of micromoles of AMP by-product formed per minute per micromole of protein, which is equivalent (equimolar) to the rate of (pp)pGpp synthesis. The x axes show substrate concentration (GMP, GDP, or GTP) in millimolar units. (B) Plots of V₀ versus [GDP] used to calculate the kinetic parameters for ppGpp synthesis by Tde-SAS without added ppGpp (control, blue filled circles), in the presence of 100 μM ppGpp (red filled squares), or 100 μM pppGpp (green filled triangles). (C) Plots of V₀ versus [S] used to calculate the kinetic parameters for pGpp (left), ppGpp (center), and pppGpp (right) synthesis by Tde-SAS_1–246. (D) Table summarizing the kinetic parameters obtained for (pp)pGpp production by Tde-SAS and Tde-SAS_1–246 from the respective sets of assays shown in panels A to C: maximum reaction velocity (V_max), Michaelis constants for GTP (K_m(GTP)), GDP (K_m(GDP)), or GMP (K_m(GMP)) in millimolar units; turnover number (k_cat) in units of AMP (= (pp)pGpp) molecules formed per second, enzymatic catalytic efficiency (k_cat/K_m). All reactions were performed in triplicate, reporting the means ± standard deviation. See Materials and Methods for experimental details.

As supplementary (p)ppGpp had minor effects on the ppGpp-synthesizing activities of Tde-SAS, the kinetic parameters for pGpp and pppGpp synthesis were determined in the absence of (p)ppGpp (Fig. 4). While the K_m value for GTP (6.09 ± 1.01 mM) was reasonably similar to that for GDP, the K_m(GMP) value was ~2-fold higher (11.68 ± 1.69 mM). The k_cat values for ppppGpp and pGpp synthesis by Tde-SAS were 25-fold and 10-fold lower than that for ppGpp, respectively. Thus, the catalytic efficiencies for pGpp or pppGpp synthesis by Tde-SAS were ~20- to 30-fold lower than that for ppGpp synthesis. The h coefficients for pGpp and pppGpp synthesis were almost identical to that for ppGpp synthesis, indicative of very low-level product-mediated stimulation of enzymatic activities.

The C-terminal TPR domain inhibits alarmone production.

The respective Michaelis-Menten kinetic parameters for alarmone synthesis by Tde-SAS_1–246 were determined in the absence of supplementary (p)ppGpp (Fig. 4), as preliminary assays indicated that the addition of 100 μM pppGpp or ppGpp had very minor effects on the rate of ppGpp synthesis (data not shown). The k_cat value for ppGpp synthesis by Tde-SAS_1–246 (45.62 ± 5.65 s⁻¹) was ~4.5-fold higher than for pGpp synthesis (10.80 ± 1.10 s⁻¹) and ~14.5-fold higher than for pppGpp synthesis (3.11 ± 0.45 s⁻¹). The K_m(GTP) and K_m(GMP) values were both ~3-fold higher than the K_m(GDP) value. Correspondingly, ppGpp was synthesized with the highest catalytic efficiency (44.10 ± 6.20 mM⁻¹ s⁻¹), which was ~14-fold higher than that of pppGpp synthesis (3.14 ± 0.25 mM⁻¹ s⁻¹) and 41-fold higher than that of pGpp synthesis (1.07 ± 0.13 mM⁻¹ s⁻¹).

The K_m(GDP) value for Tde-SAS_1–246 (1.03 ± 0.30 mM) was ~5-fold lower than the corresponding value obtained for Tde-SAS (5.29 ± 2.61 mM). The catalytic efficiency for ppGpp synthesis by Tde-SAS_1–246 was ~18-fold higher than for Tde-SAS (in the absence of added alarmone) (Fig. 4D). Analogously, the K_m values for GMP (3.44 ± 0.6 mM) and GTP (2.91 ± 0.75 mM) for Tde-SAS_1–246 were, respectively, ~3- and 2-fold lower than the values determined for Tde-SAS. The resultant catalytic efficiencies for pGpp and pppGpp synthesis by Tde-SAS_1–246 were, respectively, ~26- and 12-fold higher than those of Tde-SAS. Taken together, these results indicated that the presence of the C-terminal TPR domain greatly reduced the overall rates and catalytic efficiencies of (pp)pGpp synthesis but did not greatly affect the substrate preference; GDP was by far the most efficiently utilized substrate.

Tde-SAS (Tde SAS_1–246) synthesizes alarmone-like nucleotides (pp)pApp and ppIpp.

Tde-SAS and Tde-SAS_1–246 were incubated with ATP, ATP + ADP, or ATP + AMP, respectively, under standardized conditions to determine their respective abilities to synthesize pppApp, ppApp, and pApp (adenosine 3′-diphosphate, 5′-phosphate). Representative chromatograms of product mixtures are shown in Fig. 5A.

FIG 5.

Synthesis of (pp)pApp by Tde-SAS and Tde-SAS_1–246. (A) Representative Mono-Q anion-exchange chromatograms of enzymatic product mixtures showing the respective (pp)pGpp synthesis activities of Tde-SAS and Tde-SAS_1–246 upon incubation with ATP + AMP (left), ATP + ADP (center), or ATP (2 equivalents) (right). Blue chromatograms: Tde-SAS, red chromatograms: Tde-SAS_1–246. All sets of assays were performed under analogous conditions. The corresponding identities of the smaller peaks are indicated with arrows. (B) Specific molar activities of the Tde-SAS and Tde-SAS_1–246 proteins for the synthesis of pApp (ATP + AMP), ppApp (ATP + ADP), and pppApp (ATP, 2 equivalents). These rates are reported in the following units: micromoles (μmol) of (pp)pGpp or (pp)pApp synthesized per minute per micromole of protein (μmol · min⁻¹ · μmol⁻¹). All reactions were performed in triplicate, reporting the means ± standard deviation. See Materials and Methods for experimental details.

Tde-SAS_1–246 synthesized ppApp most effectively, with a specific molar activity (15.7 ± 1.8 μmol · min⁻¹ · μmol⁻¹) that was ~120- and 70-fold higher than that for pppApp and pApp synthesis, respectively (Fig. 5B). Tde-SAS synthesized ppApp ~8-fold slower than Tde-SAS_1–246, while the rates of pppApp and pApp synthesis were very low under these conditions. The respective rates of ppApp synthesis by Tde-SAS and Tde-SAS_1–246 were ~160- and 360-fold lower than those of ppGpp synthesis (Fig. 3B). (Note that the additional small (pp)pApp peaks labeled in the chromatograms correspond to the products of competing side reactions, with ppApp primarily formed from the low levels of ADP present in ATP solutions.) Tde-SAS_1–246 also catalyzed the synthesis of inosine-3′,5′-bis(diphosphate) (inosine tetraphosphate, ppIpp) from IDP and ATP (Fig. S6), as has previously been found for other long-RSH and SAS proteins (18, 40). Tde-SAS exhibited low-level ppIpp synthesis activities under comparable conditions. ITP could not function as a pyrophosphate donor in place of ATP (data not shown).

Tde-SAH hydrolyzes (pp)pGpp and (p)ppApp in a Mn²⁺ ion-dependent manner.

Tde-SAH (TDE1690) belongs to the ActSpo2 family of RSH proteins (31). Tde-SAH and the other treponeme SAH homologues cluster separately from nontreponeme homologues in the ActSpo2 phylogenetic clade (Fig. 6A), the majority of which belong to taxa from the phylum Actinobacteria (31). A multiple sequence alignment of the amino acid sequences of Tde-SAH and seven previously described SAH homologues is shown in Fig. 6B. This includes the C. glutamicum RelH (CgSAH; RelH_Cg; Protein Data Bank [PDB] entry 7QOD) (36, 39), Listeria monocytogenes Lmo0812 (LmSAH; PDB entry 4YF1), Pseudomonas aeruginosa PA0431 (PaSAH; PDB entry 6YVC) (38), Methylorubrum (Methylobacterium) extorquens SAH_Mex (MexSAH) (50), Cellulomonas marina ATFaRel (CmarSAH, WP_090034991) (31), human MESH1 (HsMESH1) (65), and the recently described Leptospira levettii SAH (LlevSAH; RelH_Ll; PDB entry 7QOE) (39) proteins. The locations of six highly conserved “hydrolase domain motifs” (HD1 to HD6) are indicated (27).

FIG 6.

Catalytically important amino acid motifs present within Tde-SAH and representative SAH/Mesh1 homologues. (A) ML phylogenetic tree of SAH homologues, with the major families indicated. The respective phylogenetic positions of Tde-SAH and other SAH homologues belonging to the ActSpo2 family from the genus Treponema (shaded violet) and from nonspirochete taxa (shaded orange) are indicated. (B) Alignment of amino acid sequences of Tde-SAH and representative SAH homologues from diverse families: P. aeruginosa PA0431 (PaSAH), L. levettii RelH (RelH_Ll; LlevSAH), C. glutamicum RelH (RelH_Cg; CgSAH), L. monocytogenes Lmo0812 (LmSAH), human MESH1 (HsMESH1) M. extorquens SAH (SAH_Mex; MexSAH), and C. marina ATFaRel (CmarSAH) proteins. The locations of motifs HD1 to HD6 are indicated with brackets. The location of the Asp74 residue within the eponymous HD domain motif (HD3) is indicated with an asterisk. This residue was mutated to Ala in the Tde-SAH_D74A mutant. The figure was prepared using ESPript 3.0 (88).

Recombinant Tde-SAH (MW, 27,091 Da) migrated with an apparent MW of ~44 kDa upon SEC analysis (Fig. S2C), indicating that it was homodimeric, analogous to the RelH_Cg and RelH_Ll proteins (36, 39). Tde-SAH specifically hydrolyzed pppGpp, ppGpp, and pGpp to form GTP, GDP, and GMP, respectively (Fig. 7A). Tde-SAH hydrolyzed (pp)pGpp in a Mn²⁺ ion-dependent manner. Other (divalent) metal ions were tested as potential cofactors, including Mg²⁺, Co²⁺, Zn²⁺, Fe³⁺, Ca²⁺, and Ni²⁺ (Fig. 7D). Tde-SAH had low-level ppGpp-hydrolyzing activities in the presence of Mg²⁺ ions and slightly higher activities in the presence of Co²⁺ ions but negligible activities when Zn²⁺, Fe³⁺, Ca²⁺, or Ni²⁺ ions were added (Fig. 7D). The hydrolytic activities of Tde-SAH were slightly higher in the presence of 1 mM Mn²⁺ compared to a combination of 1 mM Mn²⁺ and 10 mM Mg²⁺, suggesting that Mn²⁺ and Mg²⁺ ions compete for the same protein-binding sites. The hydrolysis of ppGpp was optimal at pH 8.4 to 8.8 (Fig. S3B). Correspondingly, all subsequent biochemical assays were performed using pH 8.4 buffer containing 1 mM Mn²⁺ ions.

FIG 7.

Hydrolysis of (pp)pGpp and (p)ppApp by Tde-SAH. (A) Representative Mono-Q anion-exchange chromatograms of enzymatic product mixtures showing the respective hydrolytic activities of Tde-SAH against pppGpp (left), ppGpp (center), or pGpp (right); 5 min incubation time. (B) Analogous chromatograms showing the respective hydrolytic activities of Tde-SAH against pppApp (left) or ppApp (right); 40 min incubation time. Red chromatograms, Tde-SAH; blue chromatograms, no-added protein (negative) controls. The respective sets of assays shown in panels A and B were performed under analogous conditions. (C) Specific molar activities of Tde-SAH for the hydrolysis of pppGpp, ppGpp, pGpp, pppApp, and ppApp substrates. All reactions were performed in triplicate, reporting the mean, ± standard deviation. (D) Divalent metal ion requirements of Tde-SAH. An overlay is shown of representative plots from continuous spectrophotometric pyrophosphatase-coupled phosphate-release assays quantifying the pyrophosphate released from the hydrolysis of ppGpp by Tde-SAH in buffer containing various metal ions, under standardized conditions. Assays containing 10 mM Mg²⁺ (red), 1 mM Mn²⁺ (green), 10 mM Mg²⁺ and 1 mM Mn²⁺ (magenta), 1 mM Co²⁺ (brown), 100 μM Zn²⁺ (yellow), 100 μM Fe³⁺ (orange), 1 mM Ca²⁺ (black), 1 mM Ni²⁺ (navy blue), and no added metal ions (blue). (E) Alarmone-hydrolyzing activities of Tde-SAH versus Tde-SAH_D74A mutant. An overlay is shown of representative plots quantifying the hydrolysis of ppGpp by Tde-SAH (blue), ppApp by Tde-SAH (red), ppGpp by Tde-SAH_D74A (green), ppApp by Tde-SAH_D74A (magenta), and ppGpp with no enzyme added (orange) in assay mixtures containing 1 mM Mn²⁺ ions under standardized conditions. See Materials and Methods for experimental details.

Further biochemical analysis using phosphate-release assays with/without the addition of a pyrophosphatase enzyme confirmed that Tde-SAH specifically hydrolyzed the diphosphate (PPi) unit from the 3′-ribose position of (pp)pGpp (data not shown). Consequently, pyrophosphatase enzyme-coupled continuous spectrophotometric assays were performed to determine the Michaelis-Menten kinetic parameters for (pp)pGpp hydrolysis by Tde-SAH, analogous to previously described methods (40). The results indicated that Tde-SAH hydrolyzed pppGpp, ppGpp, and pGpp with similar catalytic efficiencies (Fig. 8). The k_cat values were in the order ppGpp > pppGpp > pGpp, ranging from 0.79 ± 0.07 to 0.54 ± 0.04 s⁻¹. The K_m values were fairly similar for all three alarmones, being lowest for ppGpp and ppGpp at ~100 μM. Correspondingly, Tde-SAH exhibited the highest catalytic efficiency for ppGpp hydrolysis (9.06 ± 0.42 mM⁻¹ s⁻¹), with pGpp and pppGpp hydrolyzed ~40% and 60% less efficiently, respectively. There was some evidence of positive cooperativity for pGpp and ppGpp hydrolysis, with h coefficients of 1.45 ± 0.24 and 1.89 ± 0.25, respectively, but not for pppGpp.

FIG 8.

Enzymatic kinetic parameters for (pp)pGpp hydrolysis by Tde-SAH. (A) Plots of initial reaction velocity (V₀) versus substrate concentration ([S]) used to calculate the kinetic parameters for pppGpp (left), ppGpp (center), and pGpp (right) hydrolysis by Tde-SAH. The y axes show the reaction velocity in units of micromoles of pyrophosphate (PPi) formed per minute per micromole of protein. The x axes show the substrate concentration (pppGpp, ppGpp, pGpp) in micromolar units. (B) Table summarizing the respective kinetic parameters obtained for the hydrolysis of pppGpp, ppGpp, and pGpp by Tde-SAH. Shown are the maximum reaction velocity (V_max) in units of micromoles of pyrophosphate (PPi) formed per minute per micromole of protein; the Michaelis constant (K_m) for the respective pppGpp, ppGpp, or pGpp substrates in millimolar units; and the turnover number (k_cat) in units of pyrophosphate molecules formed per second, enzymatic catalytic efficiency (k_cat/K_m). The data were obtained from sets of continuous spectrophotometric pyrophosphatase-coupled phosphate release assays performed under optimal conditions, in triplicate, reporting the mean values ± standard deviation. See Materials and Methods for experimental details.

Tde-SAH hydrolyzed pppApp/ppApp, to ATP/ADP + pyrophosphate, respectively, in a Mn²⁺ ion-dependent manner (Fig. 7B). pApp could not be enzymatically synthesized in sufficient quantities and was not tested. It may be noted that under the assay conditions used (pH 8.4, 1 mM Mn²⁺), (p)ppApp molecules are more prone to nonenzymatic (metal ion-facilitated) hydrolysis than (p)ppGpp. Consequently, peaks corresponding to ppApp and pApp are observed in the chromatograms of the control reactions. The specific molar rates of hydrolysis for (pp)pGpp and (p)ppApp were evaluated under identical conditions to quantify the substrate selectivity (Fig. 7C). The results indicated that Tde-SAH hydrolyzed (pp)pGpp ~2-fold faster than pppApp or ppApp under the conditions employed.

The Asp74 residue of Tde-SAH forms part of the HD motif (HD3; Fig. 6B), which has previously been shown to be essential (or critically important) for hydrolytic activity in other RSH homologues (24, 38). The biochemical activities of the Tde-SAH_D74A (His-Asp → His-Ala) mutant were determined using assays analogous to those described above. the results revealed that the Tde-SAH_D74A protein had undetectable hydrolytic activities against ppGpp or ppApp (Fig. 7E), confirming that Asp74 played an essential role in alarmone hydrolysis.

Tde-SAS synthesizes alarmones in vivo.

The in vivo (p)ppGpp synthesis activities of Tde-SAS and Tde-SAS_1–246 were determined using a well established growth assay in E. coli ΔrelAΔspoT (CF1693) (23). The CF1693 mutant is deficient in (p)ppGpp synthesis and degradation, exhibits multiple amino acid auxotrophies, and therefore cannot grow in minimal medium (66). Complementation with a plasmid-based RSH protein capable of synthesizing (p)ppGpp to sufficient levels, but not to excessive (i.e., toxic) levels, restores growth in minimal medium (23, 36). The wild-type E. coli MG1655 strain (CF1648) was included as a reference.

Genes encoding the Tde-SAS, Tde-SAS_1–246, Tde-SAH, and S. aureus RelP (Sa-RelP) proteins were respectively cloned into the medium-copy number and arabinose-inducible pBAD33 plasmid (67) and established in the E. coli ΔrelAΔspoT strain. The pBAD33-Sa-RelP and empty pBAD33 plasmids were included as positive and negative controls, respectively. Cultivation in MOPS minimal medium lacking arabinose led to modest increases in growth rates for the pBAD33-Tde-SAS, pBAD33-Tde-SAS_1–246, and pBAD33-Sa-RelP complemented strains, compared to the pBAD33-Tde-SAH and pBAD33 strains (Fig. 9A). Analogous experiments performed in MOPS minimal medium containing arabinose (0.2%) led to notable increases in growth rates for the strains complemented with the pBAD33-Tde-SAS, pBAD33-Tde-SAS_1–246, and pBAD33-Sa-RelP plasmids, indicative of (p)ppGpp production to subtoxic levels (Fig. 9B). In contrast, there were negligible changes in the growth rates of the strains containing the empty pBAD33 and pBAD33-Tde-SAH plasmids. The growth rate of the wild-type CF1648 (MG1655) strain complemented with an empty pBAD33 plasmid remained essentially unchanged in absence and presence of inducer, which were considerably higher than those of all the complemented ΔrelAΔspoT strains under all conditions employed. Taken together, these results strongly support the premise that both the Tde-SAS and Tde-SAS_1–246 proteins can synthesize (p)ppGpp in the E. coli ΔrelAΔspoT strain to levels that can maintain its effective growth in minimal medium.

FIG 9.

Evaluation of in vivo (p)ppGpp synthesis/hydrolysis activities of Tde-SAS, TdeSAS_1–246, and Tde-SAH. The E. coli ΔrelAΔspoT (CF1693) strain is deficient in (p)ppGpp synthesis and exhibits negligible growth in minimal medium without complementation with a protein capable of mediating (p)ppGpp synthesis to subtoxic levels. (A, B) Growth curves for CF1693 complemented with (arabinose-inducible) pBAD33-Tde-SAS (purple), pBAD33-Tde-SAS_1–246 (brown), pBAD33-Tde-SAH (red), and pBAD33-Sa-RelP (green), as well as noncomplemented CF1693 (orange) and wild-type E. coli MG1655 (CF1648, blue). All strains were cultivated in MOPS minimal medium at 37°C. (A) In the absence of arabinose. (B) In the presence of 0.2% arabinose. Growth curves were measured spectrophotometrically (optical density at 600 nm [OD₆₀₀]) over 10 h, plotting the mean value from three biological replicates (shading indicates standard deviation). There were notable enhancements in growth rates for CF1693 strains complemented with pBAD33-Tde-SAS, pBAD33-Tde-SAS_1–246, and pBAD33-Sa-RelP cultivated in the presence of arabinose, indicative of (p)ppGpp synthesis. (C) Growth curves for CF1693 complemented with pBAD- and pGEX-based plasmids, cultivated in MOPS minimal medium at 37°C. The plasmid combinations tested were pBAD33-Tde-SAS + pGEX-empty (green), pBAD33-Tde-SAS + pGEX-Tde-SAH_D74A (purple), pBAD33-Tde-SAS + pGEX-Tde-SAH (light-blue), pBAD33-empty + pGEX-Tde-SAH (red), and pBAD33-empty + pGEX-empty (dark-blue). Expression was induced by the addition of isopropyl β-d-1-thiogalactopyranoside (IPTG; 0.2 mM) and arabinose (0.2%) after 2.5 h (arrow). The growth rates were highest for the pBAD33-Tde-SAS + pGEX-empty and pBAD33-Tde-SAS + pGEX-Tde-SAH_D74A strains. The growth rates of the strains complemented with pBAD33-empty + pGEX-Tde-SAH and pBAD33-empty + pGEX-empty were very low. The growth rate of the pBAD33-Tde-SAS + pGEX-Tde-SAH strain was intermediate. See Materials and Methods for detailed experimental protocols.

Tde-SAH can hydrolyze (p)ppGpp produced by Tde-SAS in vivo.

Further sets of growth complementation experiments were performed to investigate the in vivo alarmone-hydrolyzing activities of Tde-SAH, as well as the Tde-SAH_D74A mutant. In these experiments, two plasmids were stably coestablished in E. coli ΔrelAΔspoT: (i) a high-copy number, isopropyl β-d-1-thiogalactopyranoside (IPTG)-inducible pGEX plasmid (Amp^R) containing the Tde-SAH wild-type or Tde-SAH_D74A mutant genes or an empty pGEX plasmid and (ii) a pBAD33 (Cm^R) plasmid containing the Tde-SAS gene or an empty pBAD33 plasmid. Growth in MOPS minimal medium was measured over 10 h at 37°C, adding both the IPTG (0.2 mM) and arabinose (0.2%) inducers at the 2.5-h point. Growth curves are shown in Fig. 9C.

The E. coli ΔrelAΔspoT strains complemented with the pBAD33-Tde-SAS + pGEX-empty and the pBAD33-Tde-SAS + pGEX-Tde-SAH_D74A pairs of plasmids grew most effectively. This supported the premise that the Tde-SAH_D74A mutant could not hydrolyze the (p)ppGpp synthesized by Tde-SAS. Complementation with the pBAD33-empty + pGEX-empty or the pBAD33-empty + pGEX-Tde-SAH plasmids resulted in similarly low growth rates. However, complementation with pBAD33-Tde-SAS + pGEX-Tde-SAH led to growth rates intermediate between these two levels. Taken together, these results suggest that Tde-SAS synthesized sufficient quantities of (p)ppGpp, which survived long enough within the cell to mediate various growth-promoting stringent response effects before being degraded by Tde-SAH (Fig. 9C).

DISCUSSION

To the best of our knowledge, there have been no investigations into the operation of the stringent response in T. denticola or any other Treponema species, and its biological significance remains unexplored. T. denticola is remarkable by encoding one SAS and one SAH protein but lacking a bifunctional long-RSH (long-Rel) homologue (16, 31, 59). Long-RSH proteins appear to be present in all other Treponema species, with the notable exceptions of Treponema pallidum, Treponema putidum, and Treponema pedis (Table S3) (16, 31, 68–70). Our results here indicate that Tde-SAS (TDE1711) is putatively the source of (pp)pGpp and (p)ppApp in T. denticola, and Tde-SAH (TDE1690) is the protein responsible for (pp)pGpp and (p)ppApp degradation.

More than 20 different families (phylogenetic lineages) of SAS proteins have been identified (31), but only a small number have yet been studied. Tde-SAS represents the first protein in the DsRel (SAS-TPR) family to be functionally characterized. Below, we compare and contrast the characteristics and biochemical activities of Tde-SAS with homologues from other SAS families.

Tde-SAS (and Tde-SAS_1–246) adopts a homotetrameric arrangement analogous to RelP and RelQ homologues from Firmicutes (40–42, 64, 71). Similar to RelP and C. glutamicum RelS (ActRel_Cg), Tde-SAS lacks the consensus KXKXKXXR (R1), EFVT (R2), and LAMNFWAT (R3) motifs present in RelQ homologues, which are responsible for the allosteric binding of (p)ppGpp molecules that stimulate (p)ppGpp synthesis activities (14, 40–42, 64). This is consistent with the synthetic activities of Tde-SAS not being notably stimulated by the addition of (p)ppGpp (Fig. 5). Furthermore, unlike S. aureus RelP, added (p)ppGpp did not inhibit Tde-SAS activities, and micromolar concentrations of Zn²⁺ ions do not stimulate its activities (41, 42). Tde-SAS lacks potential metal-chelating residues equivalent to the adjacent histidine residues (His73 and His74) of S. aureus RelP that bind Zn²⁺ ions at the dimer interface, which putatively modulate its catalytic activities in response to changes in oxidative stress (42). This highlights the subtle differences in modulatory mechanisms operating within different SAS protein homologues.

Tde-SAS preferentially utilizes GDP for the synthesis of ppGpp. Broadly speaking, this is similar to the majority of RelP and RelQ homologues studied to date, under the majority of experimental conditions tested (14, 29, 32, 40–42, 64). C. glutamicum RelS synthesizes pppGpp most efficiently but exhibits complex kinetic behaviors (36). The bifunctional RNase-SAS RelZ protein from M. smegmatis has been reported to synthesize pGpp most efficiently (45). The recently described RelQ homologue from Clostridiodes (Clostridium) difficille (RelQ_Cd) is a notable outlier, having the ability to utilize GDP and GTP for the production of pGpp, putatively via an intermediate phosphohydrolase step (35). We found no evidence that Tde-SAS catalyzes any competing phosphotransferase or phosphohydrolase processes throughout our experimental work.

The K_m(GDP) value for Tde-SAS (5.29 ± 2.61 mM) is similar to that of C. glutamicum RelS (6.1 mM) (36) but is considerably higher than the values previously reported for M. smegmatis RelZ and Firmicutes RelP and RelQ homologues, which are in the range 0.1 to 1.7 mM (14, 40, 41, 45, 64). However, removal of the TPR domain (as in the Tde-SAS_1–246 protein) dramatically increased (pp)pGpp production levels. It increased the rate of (pp)pGpp synthesis ~3-fold and reduced K_m(GDP) more than 6-fold (to ~1 mM), making it comparable to the above-mentioned SAS proteins. Putatively, this would greatly increase ppGpp production at GDP concentrations typically found within bacterial cells (13, 46, 72–74). However, removing the TPR domain did not alter substrate selectivity, with GDP strongly preferred over GTP or GMP. Thus, the TPR domain appears to act like a brake, repressing the overall alarmone-synthesizing activities of the Tde-SAS catalytic domain by reducing the effective substrate binding affinity (increasing K_m) and reducing the overall reaction rate (decreasing k_cat).

Both Tde-SAS and Tde-SAS_1–246 were capable of synthesizing sufficient quantities of (p)ppGpp to restore the growth of the E. coli ΔrelAΔspoT (CF1693) strain in minimal medium, without inducing toxic effects (due to the overproduction of (p)ppGpp) (Fig. 9). While these sets of experiments demonstrated in vivo alarmone synthesis activities in a model bacterial system, they do not directly show that Tde-SAS (or Tde-SAS_1–246) synthesizes alarmones within the native T. denticola host. It should also be noted that this E. coli reporter system is semiquantitative in nature and cannot be used to accurately evaluate levels of alarmone production or hydrolysis. These are notable limitations of our study.

Proteins containing TPR motif domains are widely distributed throughout prokaryotes, in which they typically function as protein “interaction modules” that promote the formation of multiprotein complexes, function as chaperones, or regulate the extracellular export of proteins or exopolysaccharides (60, 75–79). While TPR motif domains comprise multiple (typically 3 to 16) tandem repeats of a 34-aa structural motif that adopts a distinctive helix-turn-helix conformation, there is considerable sequence heterogeneity in their sequence composition (60, 75). There are high levels of sequence conservation within the four respective TPR motifs in SAS-TPR homologues encoded by diverse Treponema species, indicative of conservation of structure and/or function (Fig. 2). We speculate that the TPR domain of Tde-SAS and other treponeme SAS-TPR (DsRel) homologues play analogous regulatory roles in the stringent response (see below).

Tde-SAS synthesized three different adenine nucleotide-based alarmone products: pppApp, ppApp, and pApp, albeit at greatly differing rates (Fig. 5). The pattern for AMP/ADP/ATP utilization (as diphosphate acceptor) was equivalent to that of GMP/GDP/GTP, i.e., ADP was strongly preferred for ppApp synthesis, just as GDP was preferred for ppGpp production. Notably, the rates of (pp)pApp production were tens to hundreds of times lower than the corresponding rates of (pp)pGpp production. Tde-SAS_1–246 synthesized ppApp ~8-fold faster than Tde-SAS, indicating that the TPR domain of Tde-SAS repressed (pp)pGpp and (pp)pApp production by the catalytic domain via the same mechanism.

The physiological functions and molecular sources of (p)ppApp within bacterial cells remains poorly understood (1, 31, 38, 46, 47, 49, 80). Several phylogenetic lineages of SAS proteins (PhRel, CapRel, PhRel2, and FaRel) are prolific (p)ppApp synthesizers, leading to potent cytotoxic effects (31, 47). In addition to functioning as toxins, (p)ppApp directly modulates enzymatic activities (e.g., PurF) and alters transcriptional activities in a manner notably different from that of (p)ppGpp (47, 81–83). Thus far, Rel_Mex is the only long-RSH protein shown to possess notable (p)ppApp synthesis activities (49, 50). The majority of SAS and Rel (long-RSH) proteins characterized to date do not appear to function as sources of (p)ppApp (1, 5, 38, 46, 49, 50). The putative in vivo ppApp-synthesizing abilities of Tde-SAS remain to be verified. However, Tde-SAS synthesizes ppApp rather slowly in vitro, and results from our growth rate assays indicate that Tde-SAS does not function as a “Tox-SAS” in E. coli. Furthermore, the presence of (pp)pApp in T. denticola cells remains unknown. It is conceivable that Tde-SAS synthesizes low levels of (p)ppApp as an alarmone-like transcriptional modulator or for other regulatory purposes in T. denticola cells (3, 81–83). In these regards, our study has notable limitations, and these important issues require future investigation.

There are many distinct phylogenetic lineages (families) of SAH proteins (31) (Fig. 6A), several of which have been functionally and/or structurally characterized. Tde-SAH represents the first homologue from the ActSpo2 family to be functionally characterized. The Metazoan SpoT homologue 1 (Mesh1) family, which is distributed across bacteria and metazoans, includes human MESH1 (PDB entry 3NR1; HsMesh1), Drosophila melanogaster Mesh1 (PDB entry 3NQW) (65, 84) and M. extorquens SAH (SAH_Mex; MexSAH) (50). The MixSpo (RelH) family includes C. glutamicum RelH_Cg (PDB entry 7QOD; CgSAH) (37, 39), L. levettii RelH_Ll (PDB entry 7QOE; LlevSAS) (39), and L. monocytogenes Lmo0812 (PDB entry 4YF1; LmSAH). The PbcSpo2 family includes P. aeruginosa SAH (PA0431; PDB entry 6YVC; PaSAH) (38), and the PbcSpo family includes C. marina ATFaRel (CmarSAH) (31).

Tde-SAH hydrolyzed pppGpp/ppGpp/pGpp to GTP, GDP, or GMP + pyrophosphate, respectively, with comparable catalytic efficiencies, dependent on Mn²⁺ ions (Fig. 7). Low-level hydrolytic activities were supported by Mg²⁺ ions or Co²⁺ ions. Tde-SAH hydrolyzed (p)ppApp to ATP/ADP + PPi in an analogous manner, at roughly equivalent rates. Thus, the hydrolytic activities of Tde-SAH functionally complement the synthetic activities of Tde-SAS. The respective catalytic efficiencies and K_m values for alarmone hydrolysis by Tde-SAH [K_m(pppGpp), 210 μM; K_m(ppGpp), 110 μM; K_m(pGpp), 120 μM] are very similar to those of the RelH_Cg SAH protein [K_m(pppGpp), 90 μM; K_m(ppGpp), 101 μM; K_m(pGpp), ~140 μM] (37). The (p)ppApp hydrolysis activities of RelH_Cg and RelH_Ll remain unexplored (39). SAH_Mex had negligible hydrolytic activities toward (p)ppGpp substrates but efficiently hydrolyzed pppApp and ppApp substrates (50). Drosophila Mesh1 hydrolyzed (p)ppApp and (p)ppGpp substrates with similar catalytic efficiencies, with K_m values ranging from 60 to 120 μM. While Mesh1 homologues probably function as (p)ppApp and/or (p)ppGpp hydrolases in bacteria, they are proposed to function primarily as NADPH phosphatases in metazoans (50, 84, 85). The PaSAH protein showed a notable preference for adenine-based alarmones, hydrolyzing (p)ppApp ~5- to 10-fold more rapidly and more efficiently than (p)ppGpp (38, 50). Taken together, SAH proteins across diverse taxa appear to selectively utilize Mn²⁺ cations but have “tunable” selectivities toward adenine versus guanine-based alarmone substrates.

Tde-SAH contains the HD1 to HD6 motifs identified in diverse SAH, SpoT, and bifunctional long-RSH homologues (Fig. 6B) (27). The strictly conserved HD diad (His73 and Asp74) are predicted to play a key role in binding the catalytic Mn²⁺ ion within the Tde-SAH active site (20, 24, 38, 39). Mutation of Asp74 to alanine abrogated the alarmone-hydrolyzing activities of Tde-SAH (Fig. 7E), putatively by disrupting Mn²⁺ cofactor binding. This is consistent with results from the in vivo growth rate assays, in which the E. coli ΔrelAΔspoT (CF1693) strain complemented with Tde-SAS, and Tde-SAH_D74A grew considerably faster than the strain complemented with Tde-SAS and Tde-SAH. This is putatively due to the inability of the Tde-SAH_D74A mutant to hydrolyze the (p)ppGpp synthesized by Tde-SAS in the cell. We have elucidated the X-ray crystal structure of the Tde-SAH protein, and hence a more detailed mechanistic description of this protein will be provided elsewhere (M. Wang and R.M. Watt, unpublished data).

Based on our results, we tentatively speculate that Tde-SAS and Tde-SAH modulate alarmone (and [p]ppApp) levels in T. denticola cells via the following mechanism. In its default conformation, Tde-SAS maintains a basal level of (pp)pGpp (and [p]ppApp) synthesis. While Tde-SAS synthesizes ppGpp most efficiently, the precise ratios of pppGpp/ppGpp/pGpp produced would also be governed by the respective intracellular levels of GTP/GDP/GMP. The alarmones (and [p]ppApp) produced would be rapidly hydrolyzed by Tde-SAH due to its efficient catalytic activities. We further speculate that the TPR domain of Tde-SAS forms binding associations with components of the transcription or translational machinery that are indicative of nutrient starvation or certain extracellular stresses. These protein-binding events alter the conformation of the Tde-SAS catalytic domain, derepressing alarmone production (akin to Tde-SAS_1–246). Upon cessation of binding to the TPR domain, the catalytic domain returns to its default conformation, and alarmone production drops to basal levels. Further investigations are required to validate this proposed mechanism.

In brief conclusion, we have established the activities of two previously undescribed lineages of SAS and SAH protein to significantly enhance our molecular understanding of (pp)pGpp and (p)ppApp metabolism within bacterial systems. Taken together, our in vitro and in vivo data suggest that the activities of Tde-SAS and Tde-SAH may be sufficient for the metabolism of (pp)pGpp in the periodontal pathogen T. denticola.

MATERIALS AND METHODS

Plasmid construction.

The Tde-SAS (TDE_RS08190, formerly TDE1711) and Tde-SAH (TDE_RS08100, formerly TDE1690) genes were PCR amplified from T. denticola ATCC 35405^T genomic DNA and cloned into pET28a (+) (Novagen, Merck Millipore) via BamHI/XhoI to create plasmids pET28a-Tde-SAS and pET28a-Tde-SAH, respectively (Table S1). The catalytic domain (Tde-SAS_1–246) of Tde-SAS was analogously cloned to create plasmid pET28a-Tde-SAS_1–246. The FN0926 gene from Fusobacterium nucleatum ATCC 25586^T was analogously cloned into pET28a to create plasmid pET28-Fn-SAS. The Tde-SAH_D74A mutant was created using the Phusion site-directed mutagenesis kit (Thermo Fisher Scientific, USA). The Tde-SAH and Tde-SAH_D74A genes were respectively subcloned into pGEX-4T1 expression vectors via BamHI/XhoI to create plasmids pGEX-Tde-SAH and pGEX-Tde-SAH_D74A. The genes were analogously PCR amplified and cloned into pBAD33 plasmids (67). PCR primer details are shown in Table S2. Plasmid integrity was confirmed by Sanger sequencing. The plasmids were routinely maintained in E. coli DH10B (Invitrogen) and cultivated in Luria Bertani (LB) or LB-agar media (USB) containing kanamycin (50 μg/mL).

Protein expression and purification.

Proteins were expressed from the respective recombinant plasmids that had been established in E. coli BL21(DE3) (Invitrogen), as previously described (40, 86). Briefly, 5-mL overnight cultures inoculated from single colonies were expanded into 500 mL of Terrific Broth (TB) containing kanamycin (50 μg/mL) and grown aerobically with shaking at 37°C. Protein expression was induced at OD₆₀₀ 0.6 to 0.8 by the addition of 0.2 mM IPTG (GE Healthcare), and incubation was maintained at 25°C for 12 h. The cells were chilled on ice and collected by centrifugation (6,000 × g, 4°C,10 min), the supernatant was discarded, and cell pellets were washed with cold phosphate-buffered saline (PBS; pH 7.4, 25 mL) and then stored at −70°C for future use or used directly.

The cell pellets were resuspended in 15 mL ice-cold nickel-binding buffer (25 mM Tris-HCl, pH 7.4, 500 mM NaCl, 20 mM imidazole) containing protease inhibitors (cOmplete ULTRA tablet, EDTA-free, Merck Millipore) in a 50-mL Falcon tube (Corning) and lysed by sonication with ice cooling (Vibra Cell, Sonics & Materials Inc.; 36% amplitude, 2 s on, 8 s off, 10 min total). The lysates were centrifuged (16,000 × g, 4°C, 60 min), and the decanted supernatants were filtered (0.45-μm pore size nylon membrane syringe filter, Pall Corporation) for immediate purification by immobilized metal affinity chromatography (IMAC), on pre-equilibrated 5 mL HiTrap chelating HP columns (GE Healthcare) impregnated with nickel ions, using an AKTA purifier system (GE Healthcare), with 1 mL/min flow rate and monitoring eluent at 280 nm. After loading the filtered supernatant, the columns were washed with 5 column volumes (CVs) of nickel-binding buffer. The recombinant proteins were eluted by linear gradient elution of nickel-binding and nickel-elution buffer (25 mM Tris-HCl, pH 7.4, 500 mM NaCl, 250 mM imidazole) over 20 CV. The fractions were routinely analyzed by SDS-PAGE on 12% acrylamide/bis-acrylamide gels (37.5:1; Bio-Rad).

Bioinformatic methods.

The DNA and protein sequences were download from the NCBI and routinely manipulated using BioEdit version 7.2.0 (87). Amino acid multiple sequence alignments (.fas files) were constructed using the ClustalW program within BioEdit, visualizing the results in EsPript 3.0 (88). TPRpred in the MPI Bioinformatics Toolkit (89) was used to identify the TPR motif locations using default parameters. TPR structure modeling was performed using the SWISS-MODEL webserver using Tde-SAS residues 256 to 410 as the input sequence with default settings (90).

Representative SAS/SAH homologues were identified using the NCBI Basic Local Alignment Search Tool (BLAST) (91), searching the NCBI GenBank nonredundant (nr) amino acid sequences and PDB sequences (www.rcsb.org) (92), and were classified into families based on previously published studies (16, 31). Phylogenetic relationships between protein amino acid sequences were inferred using GARLI v.2.0 using the default maximum likelihood (ML) approach (93). Constructed ML phylograms were visualized and edited using ITOL (94) to highlight branches from distinct SAS/SAH protein families. Details of the SAS and SAH protein sequences used for phylogenetic tree construction are summarized in Supplemental File 2.

Enzymatic preparation of (pp)pGpp and (p)ppApp.

Nucleotides were purchased from Sigma and were of the highest purity available. The pppGpp, ppGpp, and pGpp alarmones were prepared enzymatically using the recombinant E. faecalis RelQ (EF-RelQ) protein and were purified by ion-exchange chromatography (1-mL Resource Q anion-exchange columns; GE Healthcare, USA) followed by desalting (5-mL HiTrap desalting columns; GE Healthcare, USA) as previously described (40, 86). The pppApp and ppApp nucleotides were synthesized using analogous enzymatic approaches using the recombinant SAS protein (FN0926) from F. nucleatum ATCC 25586^T. Briefly, reaction mixtures contained 50 mM Bis-Tris propane, pH 6.8, 150 mM NaCl, 20 mM MgCl₂, 1 mM dithiothreitol (DTT), 10 mM ATP + 10 mM ADP (to make ppApp), or 20 mM ATP (to make pppApp) and were incubated overnight at 37°C. The (p)ppApp products were purified analogously to (pp)pGpp. The concentrations of purified (pp)pGpp and (p)ppApp product solutions were determined via UV spectroscopy (Nanodrop 2000c spectrometer) based on the respective molar extinction coefficients for GTP (13,700 L.mol⁻¹ · cm⁻¹ at 252 nm) or ATP (15,400 L.mol⁻¹· cm⁻¹ at 259 nm), using a dilution series, as has previously been reported (40, 86).

Nucleotide product analysis and quantification.

Enzymatic product mixtures were analyzed by anion-exchange chromatography (1 mL MonoQ 5/50 GL; GE Healthcare) using an AKTA purifier system using the following program: 100% buffer A (25 mM Tris-HCl, pH 8.0, 25 mM NaCl) for 3 CV, a linear gradient of 100% buffer A increasing to 44% buffer B (25 mM Tris-HCl, pH 8.0, 1 M NaCl) over 13 CV and then 100% buffer B for 3CV, at a flow rate of 2 mL/min. The UV absorption of the eluent was monitored at 254 nm. Nucleotides and alarmone products were identified based on their unique elution volumes, in comparison with reference standards, as previously described (40). Reference chromatograms of AMP, ADP, ATP, GMP, GDP, GTP, pGpp, ppGpp, pppGpp, ppApp, and pppApp standards run under identical conditions are shown in Fig. S1. The respective levels of AMP biproduct formed (which are equimolar to those of the corresponding enzymatic (pp)pGpp or (p)ppApp products) were quantified by measuring the respective peak areas on the chromatograms, which were then compared with a standard curve prepared from a set of AMP solutions of known concentrations (0 to 1,000 μM). The unit of activity was defined as the number of micromoles of AMP (or alarmone) synthesized per minute per micromole of protein (μmol · min⁻¹ · μmol⁻¹ protein). All reactions were performed in triplicate. The y axes of the chromatograms shown in the figures (plotted in units of milli-absorbance units, mAU) were routinely removed for the sake of clarity.

Qualitative assays for determining (pp)pGpp/(p)ppApp/ppIpp synthesis activities.

Reaction mixtures (20 μL) containing 50 mM Tris-HCl, pH 8.8 (optimal pH), 150 mM NaCl, 10 mM MgCl₂, 1 mM DTT, 5 mM ATP, 5 mM GMP/GDP/GTP/ADP/ATP/IDP (as indicated), and 250 nM protein were incubated at 37°C for 2 h, before being quenched by the addition of 2 mM EDTA and then snap-frozen in liquid nitrogen for future analysis. Each product mixture (20 μL) was diluted to 200 μL with Milli-Q water and analyzed by anion-exchange chromatography on a Mono Q column 5/50 GL (1 mL) as described above. All reactions were performed in triplicate.

Kinetic analysis of alarmone synthesis by Tde-SAS and Tde-SAS_1–246.

Kinetic parameters (V_max, k_cat, and K_m) were calculated using the Michaelis-Menten model, incorporating results from sets of assays performed in triplicate. The assays were performed at the pH value optimal for the Tde-SAS protein (Fig. S3). Reaction mixtures (20 μL) contained 50 mM Tris-HCl, pH 8.8, 150 mM NaCl, 10 mM MgCl₂, 1 mM DTT, 5 mM ATP, 0 to 10 mM GMP/GDP/GTP, 250 nM protein (Tde-SAS/Tde-SAS_1–246), with or without 100 μM pppGpp/ppGpp (as a putative allosteric modulator). Alarmone synthesis rates were determined by quantifying AMP levels as described above.

Quantitative determination of Tde-SAH (pp)pGpp and (p)ppApp rates of hydrolysis.

The respective rates of pppGpp, ppGpp, pGpp, pppApp, and ppApp hydrolysis by Tde-SAH were determined using an enzyme-coupled continuous fluorescent assay (EnzChek phosphate assay kit; Thermo Fisher Scientific, USA) as previously described (40). Assays were performed at the pH value optimal for the Tde-SAH protein (Fig. S3). Briefly, reaction mixtures in 96-well plates (200 μL) contained 50 mM Tris-HCl, pH 8.4, 150 mM NaCl, 1 mM MnCl₂, 1 mM DTT, 200 μM 2-amino-6-mercapto-7-methylpurine ribonucleoside (MESG), 0.2 units PNP, 3 μM recombinant S. aureus inorganic pyrophosphatase protein (Sa-PpaC, SAV1919), and 250 nM Tde-SAH protein. The assay mixtures were preincubated at 37°C for 10 min, and then the assays were initiated by the addition of the respective pppGpp, ppGpp, or pGpp substrate (0 to 500 μM; for assays used for Michaelis-Menton kinetic parameter determination) or 200 μM pppGpp, ppGpp, pGpp, pppApp, or ppApp (for assays used to determine specific rate of hydrolysis). Reaction mixtures were incubated at 37°C for 15 min with OD₃₆₀ readings taken every 30 s, using a SpectraMax M2e multilabel microplate reader (Molecular Devices). The rate of hydrolysis was normalized using a phosphate standard curve constructed according to the manufacturer’s recommended protocol.

Evaluation of in vivo (p)ppGpp production using E. coli ΔrelAΔspoT reporter system.

Cellular (p)ppGpp levels were evaluated using growth assays with the E. coli ΔrelAΔspoT (CF1693) strain (from Michael Cashel) (23), which was cultivated in MOPS minimal medium (1× MOPS mixture, 1.32 mM K₂HPO₄, 1% glucose as carbon source) as previously described (31, 36) with minor modifications. Plasmids and strains used are listed in Table S1. pBAD33 plasmids containing the Tde-SAS, Tde-SAS_1–246, S. aureus RelP (Sa-RelP, NWMN_2405), or Tde-SAH gene or no insert (pBAD33-empty, negative control) were transformed into E. coli CF1693 and were propagated at 37°C in LB medium containing chloramphenicol (Cm, 25 μg/mL) to ensure plasmid maintenance. Wild-type E. coli MG1655 (CF1648) (23) transformed with pBAD33 was included as an additional control. Aliquots (10 μL) of overnight cultures (single colonies inoculated into 5 mL of LB + Cm, incubated at 37°C for 16 h) were diluted to a final OD₆₀₀ of 0.08 into MOPS minimal medium (990 μL) containing chloramphenicol (25 μg/mL) and 0.2% arabinose to induce protein expression. Aliquots (200 μL) were pipetted into 96-well plates, which were incubated at 37°C for 10 h with OD₆₀₀ readings taken every 10 min (immediately after a few seconds of automated agitation), using a SpectraMax M2e multilabel microplate reader (Molecular Devices).

Analogous sets of growth assays were performed in E. coli CF1693 transformed with pairs of pBAD33 and pGEX-4T1 plasmids respectively containing Tde-SAS, Tde-SAH, Tde-SAH_D74A (hydrolytically inactive), or no genetic insert (negative control). Transformed E. coli CF1693 strains were routinely propagated at 37°C in LB medium containing ampicillin (100 μg/mL) and chloramphenicol (25 μg/mL) for stable plasmid maintenance. These strains were used in growth assays performed analogously to those described above, except using MOPS minimal medium containing ampicillin (100 μg/mL), chloramphenicol (25 μg/mL), arabinose (0.2%), and IPTG (0.2 mM) to induce protein expression. Throughout these experiments, we were fastidious with the cultivation and restreaking of this “(p)ppGpp zero” strain to reduce the likelihood of spontaneous suppressor (e.g., RNA polymerase) mutations occurring (66). Additional experimental details are included in the Supplemental Methods.

Data availability.

All research materials and primary data sets are available upon request (Rory M. Watt, rmwatt@hku.hk).