Abstract
Sensory and regulatory domains allow bacteria to adequately respond to environmental changes. The regulatory ACT domains are mainly found in metabolic-related proteins as well as in long (p)ppGpp synthetase/hydrolase (SD/HD) enzymes. Here, we investigate the functional role of the ACT domain of SpoT, the only (p)ppGpp SD/HD of Caulobacter crescentus. We show that SpoT requires the ACT domain to hydrolyse ppGpp in an efficient way. In addition, our in vivo and in vitro data show that the phosphorylated version of EIIANtr (EIIANtr~P) interacts directly with the ACT to inhibit the hydrolase activity of SpoT. Finally, we highlight the conservation of the ACT-dependent interaction between EIIANtr~P and SpoT/Rel along with the PTSNtr-dependent regulation of (p)ppGpp accumulation upon nitrogen starvation in Sinorhizobium meliloti, a plant-associated α-proteobacterium. Thus, this work suggests that α-proteobacteria might have inherited from a common ancestor, a PTSNtr dedicated to modulate (p)ppGpp levels.
Introduction
Bacteria use a wide range of sensory and regulatory domains to integrate environmental signals. In response to nutrient limitation, most bacteria synthesize a second messenger, the guanosine (penta) tetra-phosphate commonly referred to as (p)ppGpp. This molecule helps in reallocating cellular resources notably by reprograming transcription and interfering with cell cycle progression (Hallez et al., 2017; Hauryliuk et al., 2015). In Escherichia coli, (p)pppp levels are regulated by RelA and SpoT, two long RelA/SpoT homolog (RSH) enzymes. SpoT carries a synthetase domain (SD) that is poorly activated by starvation signals (carbon, fatty acids, phosphate, …) and a functional hydrolase domain (HD). By contrast, RelA harbours a SD activated by amino acids scarcity and a degenerated and inactive HD domain (reviewed in (Potrykus and Cashel, 2008) and (Hauryliuk et al., 2015)). Despite these differences, both RSH enzymes present the same domain architecture with catalytic domains (SD and HD) located towards the N-terminus (NTD) and regulatory domains (TGS for ThrRS, GTPase and SpoT and ACT for Aspartokinase, Chorismate mutase and TyrA) at the C-terminal end (CTD) (Figure 1). The CTD is thought to play a critical role in RSH by sensing the starvation signal and transducing it to the catalytic domains. RelA for example is known to be associated with the ribosome where it detects deacylated tRNA in the ribosomal acceptor site (A-site) as a signal for amino acid starvation, which activates SD activity (Haseltine and Block, 1973). Recent studies using cryo-electron microscopy revealed that E. coli RelA adopts an extended “open” conformation on stalled ribosomes with the A-site/tRNA in contact with the TGS domain in CTD (Arenz et al., 2016; Brown et al., 2016; Loveland et al., 2016). Likewise, activation of SpoT SD activity in E. coli cells starved for fatty acids has been shown to require an acyl carrier protein (ACP) that binds to the TGS of SpoT (Battesti and Bouveret, 2006). Although the role of CTD is undoubtedly critical for regulating catalytic functions of RSH, the mechanisms by which this regulation is mediated remain unknown.
In contrast to E. coli, most α-proteobacteria encode bifunctional (SD/HD) RSH usually referred to as SpoT or Rel (Atkinson et al., 2011). This is the case of Caulobacter crescentus that harbours a single RSH (SpoT) sensitive to nitrogen or carbon starvation (Boutte and Crosson, 2011; Lesley and Shapiro, 2008). C. crescentus divides asymmetrically to generate two dissimilar progeny, a motile swarmer cell and a sessile stalked cell. The stalked cell initiates a new replication cycle (S phase) immediately at birth whereas the swarmer cell first enters into a non-replicative G1 phase before starting replication and concomitantly differentiating into a stalked cell (Curtis and Brun, 2010). Once accumulated, (p)ppGpp modulates cell cycle progression by specifically extending the G1/swarmer phase (Chiaverotti et al., 1981; Gonzalez and Collier, 2014). Recently, we reported a key role played by the nitrogen-related phosphotransferase system (PTSNtr) in regulating (p)ppGpp accumulation in response to nitrogen starvation (Ronneau et al., 2016). We showed that EINtr, the first protein of PTSNtr, uses intracellular glutamine concentration as a proxy for nitrogen availability, since glutamine binds to the GAF domain of EINtr to inhibit its autophosphorylation. Therefore, glutamine deprivation strongly stimulates autophosphorylation of EINtr, which in turn triggers phosphorylation of the downstream components HPr and EIIANtr. Once phosphorylated, both HPr~P and EIIANtr~P modulate activities of SpoT to quickly increase (p)ppGpp levels. Whereas HPr~P stimulates SD activity of SpoT by an unknown mechanism, EIIANtr~P interacts directly with SpoT to interfere with its HD activity (Ronneau et al., 2016). The plant-associated α-proteobacterium Sinorhizobium meliloti also accumulates (p)ppGpp upon nitrogen or carbon starvation from a single RSH called Rel (Krol and Becker, 2011; Wells and Long, 2002), but the mechanism beyond this regulation remains unknown.
In this work, we investigate the role of the ACT domain in regulating the activity of RSH enzymes. In particular, we show that the ACT domain is indispensable for HD activity of C. crescentus SpoT and that EIIANtr~P inhibits (p)ppGpp hydrolysis by directly interacting with ACT of SpoT. In addition, we show that the EIIANtr~P-mediated regulation of RSH is conserved in S. meliloti, suggesting PTSNtr plays a critical role in sensing metabolic state and regulating cell cycle progression in α-proteobacteria.
Results
EIIANtr~P interacts directly with the ACT domain of SpoT
We showed previously that only the phosphorylated version of C. crescentus EIIANtr (EIIANtr~P) was able to interact with SpoT in a bacterial two-hybrid (BTH) assay (Ronneau et al., 2016). To map the domains of SpoT interacting with EIIANtr~P, we used truncated versions of SpoT in a BTH assay (Figure 1a). We found that deleting the ACT domain (SpoTΔACT) precluded the interaction with EIIANtr~P and with the isolated ACT domain (ACT506-742), but not with full-length SpoT (Figure 1b-c). Moreover, the ACT domain alone was able to strongly interact with EIIANtr~P (Figure 1b-c) as well as with itself (Figure 1 – figure supplement 1). Together, these results suggest that ACT-ACT interactions occur between SpoT subunits and show that the ACT domain is required to mediate the interaction between SpoT and EIIANtr~P.
Inactivating the ACT domain increases (p)ppGpp levels
Disrupting the interaction between SpoT and EIIANtr~P should release the hydrolase (HD) activity of SpoT, since strains that either did not express EIIANtr (ΔptsN) or expressed only the non-phosphorylated form of EIIANtr (ΔptsP, ΔptsH or ptsNH66A) actively hydrolysed (p)ppGpp in vivo (Ronneau et al., 2016). Accordingly, a C. crescentus strain expressing spoTΔACT as the only copy of spoT should phenocopy the ΔptsP strain by decreasing motility and shortening G1 lifetime (Ronneau et al., 2016). Surprisingly, we found that spoTΔACT cells phenocopied rather the HD-dead mutant spoTD81G than ΔptsP (Ronneau et al., 2016) Indeed, the spoTΔACT mutation led to an increase of motility, an accumulation of G1/swarmer cells and a growth delay in PYE complex medium (Figure 2a-d). As for spoTD81G, these phenotypes might be due to a (p)ppGpp excess in spoTΔACT. In support of this, spoTΔACT cells accumulated (p)ppGpp without stress (Figure 2e). In addition, abolishing the synthetase (SD) activity in spoTΔACT cells, either by deleting ptsP (EINtr) or by incorporating a Y323A substitution into SpoT (spoTY323A ΔACT) known to specifically inactivate SD activity (Boutte and Crosson, 2011; Ronneau et al., 2016), suppressed all the phenotypes (Figure 2a-e and Figure 2 – figure supplement 1a). The (p)ppGpp accumulation observed in spoTΔACT cells could be due to an increase of SD activity or a decrease of HD activity. To discriminate between these two possibilities, we measured (p)ppGpp levels in nitrogen-deplete (-N) conditions, which mainly relies on SD (Ronneau et al., 2016). We observed that spoTΔACT and SpoTD81G protein levels in the corresponding mutant strains were slightly higher than the SpoT level in the wild-type strain (Figure 2 – figure supplement 1b-c). Nevertheless, upon nitrogen starvation, neither spoTΔACT nor spoTD8IG cells produced more (p)ppGpp than wild-type cells (Figure 2 – figure supplement 1d). This suggests that SD activity is not enhanced in the of spoTΔACT nor spoTD8IG strains. The slight increase of spoTΔACT nor spoTD8IG levels is likely a result of the positive feedback loop of (p)ppGpp on spoT promoter. Indeed, PspoT displayed higher activity in strains accumulating (p)ppGpp (ptsPL83Q and Pxylx::relA-FLAG) and lower activity in ppGpp0 strains (ΔptsP, ΔptsH and ΔptsN), so that SpoT protein levels varied according to (p)ppGpp levels (Figure 2 – figure supplement 1e-g). Thus, the higher concentration of (p)ppGpp detected in sporΔACT cells may come from an inactivation of HD activity, thereby suggesting that ACT is required for HD activity.
The hydrolase activity of SpoT requires the ACT domain
To test whether ACT was required for HD activity, we first measured the endogenous HD activity of SpoT in vivo (Ronneau et al., 2016). To this end, we used C. crescentus strains in which (p)ppGpp (i) cannot be produced anymore by the endogenous SpoT since its SD activity has been inactivated with the Y323A mutation (Boutte and Crosson, 2011), but (ii) can be synthesized upon addition of xylose by a truncated version of the E. coli RelA (p)ppGpp synthetase expressed from the xylose-inducible promoter (PxyIx::relA-FLAG) at the xylX locus (Gonzalez and Collier, 2014). Thus in these strains the only HD activity capable to degrade (p)ppGpp produced by E. coli RelA was supplied by endogenous SpoT variants. In agreement with our previous observations, inactivation of SpoT HD in such a background (spoTD81G y323A: P xyIx::relA-FLAG,) led to a strong (p)ppGpp accumulation, an extension of the G1 phase and a growth arrest (Figure 3; (Ronneau et al., 2016)). In contrast, releasing SpoT HD activity (ΔptsP spoTY323A: P xyIx::relA-FLAG) led to undetectable levels of (p)ppGpp, a reduced G1 phase and an optimal growth (Figure 3; (Ronneau et al., 2016)). Interestingly, deleting the ACT domain of SpoT (spoTY323A ΔACT,· Pxylx::relA-FLAG) led to a (p)ppGpp accumulation, a G1 proportion and a growth rate similar to the HD-dead mutant (spoTD81G y323A: Pxylx::relA-FLAG), and this independently of the presence of ptsP (Figure 3).
These data strongly suggest that the ACT domain is strictly required in vivo for the hydrolase activity of SpoT and that EIIANtr~P inhibits HD activity of SpoT by interfering with the ACT domain. To test these hypotheses, we performed in vitro HD assays with the full-length enzyme and mutants lacking the ACT domain (SpoTΔACT) or containing only the two catalytic domains (SpoT1-373). We observed that full-length SpoT could efficiently degrade [32P]ppGpp but the deletion of the ACT domain in both SpoTΔACT and SpoT1-373 strongly affected HD activity, since radiolabelled [32P]ppGpp remained mostly intact in the presence of both ACT-deficient SpoT variants (Figure 4a, compare lane 3 to lanes 4-7). To check whether phosphorylated EIIANtr could modulate the HD activity of SpoT, [32P]ppGpp hydrolysis was measured in the presence of EIIANtr and non-phosphorylatable (EIIANtrH66A) or phosphomimetic (EIIANtrH66E) version of EIIANtr. We found that only EIIANtrH66E was able to interfere with SpoT HD activity (Figure 4b-c). The role of phosphorylation was further confirmed using EIIANtr~P. Our results show that EIIANtr~P efficiently inhibited SpoT HD activity, protecting [32P]ppGpp from hydrolysis (Figure 4d and Figure 4 – figure supplement 1). Altogether, these data show that EIIANtr~P inhibits the HD activity of SpoT by directly interacting and likely interfering with the ACT domain.
PTSNtr regulates (p)ppGpp accumulation in Sinorhizobium meliloti
The inhibition of EINtr autophosphorylation by glutamine was first observed in the γ-proteobacterium Escherichia coli (Lee et al., 2013) and the plant-associated α-proteobacterium Sinorhizobium meliloti (Goodwin and Gage, 2014). In addition, S. meliloti also accumulates (p)ppGpp in response to nitrogen starvation (Krol and Becker, 2011; Wells and Long, 2002). This suggests that PTSNtr stimulates (p)ppGpp accumulation in response to glutamine deprivation as shown for C. crescentus (Ronneau et al., 2016). To test this hypothesis, we checked whether the nitrogen-related EIIA component of S. meliloti (EIIANtr) was able to interact with the bifunctional (SD/HD) RSH of S. meliloti (Rel) in a BTH assay. As shown in Figure 5a, the full-length Rel fused to T25 (T25-Rel) interacted with EIIANtr fused to T18 (T18-EIIANtr). Similarly to C. crescentus, the deletion of ACT (T25-RelΔACT) abolished this interaction (Figure 5a). Another conserved feature is that phosphorylation of EIIANtr enhanced interaction with Rel. Indeed, we previously showed that (i) EIIANtr was phosphorylated in the BTH assay by the endogenous PTSNtr system of E. coli (EINtr and NPr) and (ii) the interaction between SpoT and EIIANtr was altered in an E. coli Anpr background (Ronneau et al., 2016). Likewise, the interaction between S. meliloti T18-EIIANtr and T25-Rel was strongly diminished in a Δnpr background (Figure 5b). This indicates that phosphorylation of EIIANtr also enhances the interaction with Rel. We constructed single in-frame deletion of ptsP (SMc02437) and rel (SMc02659) genes to test the PTSNtr-dependent accumulation of (p)ppGpp in S. meliloti in response to nitrogen deprivation. Upon nitrogen starvation (-N), neither Δrel nor AptsP cells accumulated (p)ppGpp in contrast to wild-type cells (Figure 5c). The increase of G1 proportion that results from the accumulation of (p)ppGpp upon nutrient limitation was previously used to synchronize S. meliloti (De Nisco et al., 2014). Thus if PTSNtr regulates (p)ppGpp levels, we reasoned that the G1 proportion of Δrel and ΔptsP populations should be reduced. This is exactly what we observed, with a proportion of G1 cells in the ΔptsP and Δrel strains reduced in comparison to the wild-type strain (Figure 5d-e). In contrast, the ectopic production of (p)ppGpp in non-starved S. meliloti cells with an IPTG-inducible version of RelAEc strongly increased the proportion of G1 cells and led to growth arrest (Figure 5 – figure supplement 1). Altogether, these results support a conserved role of PTSNtr in regulating (p)ppGpp accumulation in response to nitrogen starvation as well as in the control of cell cycle progression in S. meliloti.
Discussion
Two decades ago, mutational analysis of the spoT gene of E. coli suggested that the C-terminal end of SpoT including the ACT domain was required to bind regulators of the hydrolase activity of SpoT (Gentry and Cashel, 1996). Later, Mechold et al. reported that deleting the C-terminal domain of SpoT in Streptococcus equisimilis led to a strong inhibition of (p)ppGpp degradation in vitro of about 150 fold in comparison to the full-length protein (Mechold et al., 2002). In agreement with these studies, our work supports that the regulatory ACT domain modulates SpoT hydrolase (HD) activity in C. crescentus. In a nitrogen-rich environment (+N), the ACT domain is free to stimulate HD activity, thereby limiting (p)ppGpp concentration. Upon nitrogen starvation (-N), the last component of the nitrogen-related PTS (PTSNtr), EIIANtr, is phosphorylated and binds the ACT domain of SpoT. This physical interaction interferes with the ACT-dependent stimulation of SpoT HD activity (Figure 6). The recent RelA structures on stalled ribosomes highlighted the role of C-terminal domain (CTD) in sensing nutrient availability (Arenz et al., 2016; Brown et al., 2016; Loveland et al., 2016). When bound to the ribosome, RelA adopts an “open” conformation, which is thought to relieve the inhibitory effect of CTD on its synthetase (SD) activity and leads to (p)ppGpp synthesis (Gropp et al., 2001). This “open” conformation seems to be favoured by specific interactions between the ribosome stalk, the A-site finger and the tRNAs with the CTD (Loveland et al., 2016). Given the conserved domain architecture of RelA and SpoT proteins, we propose that EIIANtr~P modulates SpoT conformation to decrease its HD activity (Figure 6). Since ACT seems to interact with itself, it is tempting to speculate that dimerization of ACT induces a conformation that enhances the HD activity. Oligomerization of long RSH has already been proposed to regulate their activity (Gropp et al., 2001; Jain et al., 2006). In such a scenario, EIIANtr~P would preclude or interfere with the active conformational state. The ACT domain is highly prevalent among RSH enzymes, thus the role of the ACT domain in sustaining the hydrolase activity in these RSH is likely conserved as well.
We also reported that SpoTΔACT is still able to increase (p)ppGpp concentration upon nitrogen starvation by stimulating SD activity in a PTSNtr-dependent way (Figure 2 – figure supplement 1d). This regulation could involve the TGS domain as described for E. coli, in which stimulation of SpoT SD activity upon fatty acid starvation depends on the TGS domain (Battesti and Bouveret, 2006). In support of this, deletion of the CTD harboring the TGS and the ACT domains completely abolished the ability of C. crescentus SpoT to produce (p)ppGpp upon nitrogen starvation (Boutte and Crosson, 2011). Altogether, our data support that regulators bind to SpoT via the regulatory CTD domains to modulate enzymatic activities.
Besides C. crescentus, another α-proteobacterium, Sinorhizobium meliloti, uses PTSNtr to sense nitrogen starvation and to stimulate (p)ppGpp accumulation (Figure 6). We found that EIIANtr interacts with Rel (Figure 5a) and that phosphorylation of EIIANtr promotes this interaction (Figure 5b). As glutamine also inhibits phosphorylation of EINtr in S. meliloti (Goodwin and Gage, 2014), glutamine deprivation likely leads to hyperphosphorylation of downstream PTSNtr components, favouring subsequent interaction between EIIANtr~P and Rel. In support of that, we showed S. meliloti required EINtr protein to accumulate (p)ppGpp upon nitrogen starvation (Figure 5c), as previously shown for C. crescentus (Ronneau et al., 2016). In addition, we observed that the physical interaction between Rel and the phosphorylated form of EIIANtr also occurs in another α-proteobacterium, Rhodobacter sphaeroides (Figure 5 – figure supplement 2). Altogether, our data support that the PTSNtr-dependent control of SpoT/Rel activities is another conserved feature in α-proteobacteria (Hallez et al., 2004).
Similarly to C. crescentus (Gonzalez and Collier, 2014; Ronneau et al., 2016), we found that (p)ppGpp accumulation also delays the G1-to-S transition in S. meliloti, (Figure 5d-e and Figure 5 – figure supplement 2). In fact, this feature has been used to synchronize a population of S. meliloti by transiently blocking bacteria starved for carbon and nitrogen in G1 phase of the cell cycle (De Nisco et al., 2014). The specific interference of (p)ppGpp with the G1-to-S transition of the cell cycle could be another conserved feature in α-proteobacteria.
Methods
Bacterial strains and growth conditions
Oligonucleotides, strains and plasmids used in this study are listed in Supplementary Table. 1, 2 and 3, together with construction details provided in the Supplementary Methods. Escherichia coli Top10 was used for cloning purpose, and grown aerobically in Luria-Bertani (LB) broth (Invitrogen) (Casadaban and Cohen, 1980). Electrocompetent cells were used for transformation of E. coli. All Caulobacter crescentus strains used in this study are derived from the synchronizable (NA1000) wild-type strain, and were grown in Peptone Yeast Extract (PYE) or synthetic M2 (20 mM PO43-, 9.3 mM NH4+; +N) or P2 (20 mM PO43-; -N) supplemented with 0.5 mM MgSO4, 0.5 mM CaCl2, 0.01 mM FeSO4 and 0.2% glucose (respectively M2G or P2G) media at 28-30 °C. Growth was monitored by following the optical density at 660 nm (OD660) during 24 hrs, in an automated plate reader (Epoch 2, Biotek) with continuous shaking at 30 °C. Motility was monitored on PYE swarm (0.3 % agar) plates. Area of the swarm colonies were quantified with ImageJ software as described previously (Ronneau et al., 2016). For kinetic experiments with Sinorhizobium meliloti (Figure 5 – figure supplement 1d), bacteria where cultivated overnight at 30 °C in LBMC (LB broth with 2.5 mM MgSO4 and 2.5 mM CaCl2) supplemented with kanamycin, then back-diluted in LBMC during 3 hrs before induction with 0.1 mM IPTG. Samples were taken each hour during 5 hrs. For E. coli, antibiotics were used at the following concentrations µg/ml; in liquid/solid medium): ampicillin (50/100), kanamycin (30/50), oxytetracycline (12.5/12.5) where appropriate. For C. crescentus, media were supplemented with kanamycin (5/20), tetracycline (1/2.5) where appropriate. The doubling time of Caulobacter strains was calculated in exponential phase (OD660: 0.2 - 0.5) using D = [ln(2).(T(B) – T(A))) / (ln(OD660(B))-ln(OD660(A))] and normalized according to the wild-type strain. E. coli S17-1 and E. coli MT607 helper strains were used for transferring plasmids to C. crescentus by respectively bi- and tri-parental mating. In-frame deletions were created by using pNPTS138-derivative plasmids and by following the procedure described previously (Ronneau et al., 2016).
Bacterial two-hybrid assays
Bacterial two-hybrid (BTH) assays were performed as described previously in (Ronneau et al., 2016). Briefly, 2 μl of MG1655 cyaA::frt (RH785) and MG1655 cyaA::frt Δnpr (RH2122) strains expressing T18 and T25 fusions were spotted on MacConkey Agar Base plates supplemented with ampicillin, kanamycin, maltose (1%), and incubed for 1 day at 30 °C. All proteins were fused to T25 (pKT25) or T18 (pUT18C) at their N-terminal extremity. The β-galactosidase assays were performed as described in (Ronneau et al., 2016). Briefly, 50 μl E. coli BTH strains cultivated overnight at 30° C in LB medium supplemented with kanamycin, ampicillin and IPTG (1 mM) were resuspended in 800 μl of Z buffer (60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4) and lysed with chloroform. After the addition of 200 μl ONPG (4 mg/ml), reactions were incubated at 30° C until color turned yellowish. Reactions were then stopped by adding 500 μl of 1 M Na2CO3, and absorbance at 420 nm was measured. Miller Units are defined as (OD420 × 1,000)/ (OD590 × t × v), where “ OD590” is the absorbance of the cultures at 590 nm before the β-galactosidase assays, “t” is the time of the reaction (min), and “v” is the volume of cultures used in the assays (ml). All the experiments were performed with at least three biological replicates.
Flow cytometry analysis
DNA content was measured using Fluorescence-Activated Cell Sorting (FACS) as described previously in (Ronneau et al., 2016). Briefly, cells were fixed in ice-cold 70% Ethanol. Fixed samples were then washed twice in FACS staining buffer (10 mM Tris pH 7.2, 1 mM EDTA, 50 mM NaCitrate, 0.01% Triton X-100) containing 0.1 mg/ml RNaseA and incubated at room temperature (RT) for 30 min. Cells were then harvested by centrifugation for 2 min at 8,000 x g, resuspended in 1 ml FACS staining buffer containing 0.5 μM Sytox Green Nucleic acid stain (Life Technologies), and incubated at RT in the dark for 5 min. Samples were analyzed in flow cytometer (FACS Calibur, BD Biosciences) at laser excitation of 488 nm. Percentage of gated G1 cells of each strain was then normalized using gated G1 cells of the wild-type strain as reference.
Detection of intracellular (p)ppGpp levels
(p)ppGpp levels were visualized as described previously in (Ronneau et al., 2016) for Caulobacter crescentus (Cc) and in (Krol and Becker, 2011; Wells and Long, 2002) for Sinorhizobium meliloti (Sm). Briefly, strains were grown overnight in PYE and then diluted for a second overnight culture in M5GG (Cc) or grown overnight in LBMC medium (Sm). Then, cells were diluted a second time in M5GG (Cc) or in LBMC medium (Sm) and grown for 3 hrs to reach an OD660 of 0.5 (Cc) or 0.7 (Sm). Cells were then split into two parts and washed twice with P5G-labelling buffer (Cc; (Ronneau et al., 2016)) or with MOPS-MGS without glutamate (Sm; (Mendrygal and Gonzalez, 2000)). One milliliter of cells were then resuspended in 225 μl of P5G-labelling (-N) or M5G-labelling (+N) for C. crescentus and in 225 μl of MOPS-MGS with (+N) or without (-N) glutamate and 0.05% NH4+ for S. meliloti. In addition, media were supplemented with 25 μl of KH232PO4 at 100 μCi ml-1 and incubated for 1 hr (Sm) or 2 hrs (Cc) with shaking at 30 °C. Then, samples were extracted with an equal volume of 2 M formic acid, placed on ice for 20 min and then stored overnight at -20 °C. All cell extracts were pelleted at 14,000 rpm (18,000 x g) for 3 min and 6 x 2 μl (Cc) or 3 x 2 ill (Sm) of supernatant were spotted onto a polyethyleneimine (PEI) plate (Macherey-Nagel). PEI plates were then developed in 1.5 M KH2PO4 (pH 3.4) at room temperature. Finally, TLC plates were imaged on a MS Storage Phosphor Screen (GE Healthcare) and analysed with Cyclone Phosphor Imager (PerkinElmer). For hydrolase experiments (Figure 3C), cells were incubated 1 hr in P5G supplemented with xylose (0.1%). Then, cells were washed twice with P5G-labelling and resuspended in P5G-labelling (-N) supplemented with KH232PO4, xylose (0.1%) and glutamine (9.3 mM).
β-galactosidase assay
The β-galactosidase assays performed to measure PspoT-lacZ activity were essentiality done as for the BTH assay with the following modifications. One milliter of Caulobacter strains harbouring the PspoT-lacZ fusion was resuspended in 800 μl of Z buffer and the absorbance of the cultures at 660 nm (OD660) instead of 590 nm was measured before the β-galactosidase assays. All the experiments were performed with three biological replicates and were normalized according to the wild-type strain harbouring the PspoT-lacZ fusion cultivated at 30°C.
Immunoblot analysis
Immunoblot analyses were performed as described in (Beaufay et al., 2015) with the following primary antibodies: anti-MreB (1:5,000) (Beaufay et al., 2015), anti-SpoT (1:5,000) and secondary antibodies: anti-rabbit linked to peroxidase (GE Healthcare) at 1:5,000, and visualized thanks to Western Lightning Plus-ECL chemiluminescence reagent (Biorad) and ImageQuant LAS400 (GE Healthcare).
Proteins purification
The purification of B. subtilis EI was essentially done as described in (Galinier et al., 1997), with the modifications indicated below. Genes encoding C. crescentus SpoT, SpoTD81G, SpoT 1-373, SpoTΔAcτ, as well as HPr, EIIANtr, EIIANtrH66E and EIIANtrH66A were transformed into E. coli BL21 (DE3) for protein production. In the case of B. subtilis EI, E. coli NM522 was transformed with plasmid pQE-30-ptsI_Bs as described in (Galinier et al., 1997) for protein expression.
Each protein contains an N-terminal 6His-tag for purification by Ni-NTA chromatography. Cells were grown to an OD600 of ~ 0.7 in LB medium (1 L) at 37 °C. Isopropyl-β-D-thiogalactoside (IPTG) was added to a final concentration of 0.5 mM and incubated overnight at 28 °C. Cells were harvested by centrifugation for 20 min at 6,000 x g, 4°C, resuspended in KCl 8 mM, TCEP 1 mM, MgCl2 2 mM, Tris 50 mM at pH 8, Protease Inhibitor Cocktail (Roche) and lysed with a cell disruptor at 60 psi in KCl 500 mM, TCEP 2 mM, Tris 50 mM at pH 8, NaCl 500 mM, glycerol 1%, Protease Inhibitor Cocktail. The lysis extract was centrifuged for 20 min at 25,000 x g, 4°C. Supernatants were loaded onto a HisTrap HP 1 ml column (GE Healthcare) in HEPES 50 mM, KCl 500 mM, NaCl 500 mM, MgCl2 2 mM, TCEP 1 mM, glycerol 2%, mellitic acid 0.002%, Protease Inhibitor Cocktail (Roche), imidazole 5 mM, pH 7.5 and eluted with imidazole. The fractions recovered from the Ni-NTA were further purified by size exclusion chromatography (Superdex 200, for SpoT, SpoTD81G, SpoTcat, SpoTΔACT and EINtr, and Superdex 75 for HPr, EIIANtr, EIIANtrH66E and EIIANtrH66A). Purified samples of SpoT were also used to immunize rabbits in order to produce anti-SpoT polyclonal antibodies.
In vitro phosphorylation of EIIANtr
To phosphorylate EIIANtr, B. subtilis EI, C. crescentus HPr and C. crescentus EIIANtr were mixed at final concentrations of 2.5 µM, 2.5 µM and 50 µM respectively, in phosphorylation buffer [25 mM Tris pH 7.4, 10 mM KCl, 10 mM MgCl2, 1 mM DTT] and incubated at 37 °C for 10 min. The phosphorylation mix was then incubated in 2 mM of PEP at 37 °C for 30 min. Phosphorylated EIIANtr (EIIANtr~P) was further purified by size exclusion chromatography on a Superdex 75 and dialysed for 72 hours at 4°C, in HEPES 50 mM, KCl 500 mM, NaCl 500 mM, MgCl2 2 mM, TCEP 1 mM, glycerol 2% and stored in glycerol 20%, at -20°C.
In vitro hydrolase assay
To assess SpoT hydrolase activity, 32PppGpp was synthesized by enzymatic reaction catalysed by RelA from Chlorobaculum tepidum. To synthesize 32PppGpp, 1 µM of RelACtep was incubated in TCEP 1 mM, MgCl2 1 mM, NaCl 50 mM, Tris 10 mM at pH 7.4. Then, 100 µM GDP were added to the synthesis reaction and incubated for 10 min at 37 °C, followed by an addition of 3 pM [γ32P] ATP (PerkinElmer) incubated for 45 min, 37°C. 32PppGpp was extracted from the reaction medium by centrifugation in Amicon 3K Centrifugal filter (Millipore) at 13,000 x g for 25 min. The hydrolase assays were performed by incubating for 10 min at 37 °C (i) 1 or 2 µM of SpoT1-373 or SpoTΔACT (Figure 4a), (ii) 1 µM of SpoT with increasing concentrations of EIIANtr, EIIANtrH66A or EIIANtrH66E (5, 10 and 20 µM) (Figure 4b-c) or (iii) 1 µM of SpoT with 20 µM of purified EIIANtr~P (Figure 4 – figure supplement 1). The reaction mix was then incubated with 10 µl of 32PppGpp to a final volume of 30 µl. The reaction was stopped by adding 2 μl of 12 M formic acid. Reaction products were separated by thin layer chromatography (TLC) by transferring 2 μL of the reaction medium onto a TLC PEI Cellulose F membrane (Millipore). Chromatography membranes were placed in 1 M of KH2PO4 buffer, pH 3.0, for 50 min at room temperature. The dried TLC membrane was placed in a Phosphor Screen plate (GE Healthcare) for 1 hr and the Phosphor Screen revealed with a phosphoimager.
Author Contributions
S.R. and R.H conceived and designed the experiments. S.R. performed all the experiments except otherwise stated. A.M. did the cloning and preliminary tests for proteins purification and in vitro phosphorylation assays. J.C-M purified the proteins for the biochemical assays and performed the in vitro HD assays (Figure 4). S.R., J.C-M, A.G-P and R.H analyzed the data. S.R. and R.H. wrote the paper.
Competing financial interests
The authors declare no competing financial interests.
Acknowledgements
We are grateful to Emanuele Biondi, Gabriele Klug, Anne Galinier and Josef Deutscher for providing strains and/or plasmids. We thank the members of the BCcD team for critical reading of the manuscript and helpful discussions; Guy Houbeau at the Animal Care Facility of the University of Namur for immunizing rabbits with purified SpoT. This work was supported by a Research Credit (CDR J.0169.16) from the Fonds de la Recherche Scientifique – FNRS to R.H and FNRS-EQP U.N043.17F, FRFS-WELBIO grant (CR-2017S-03), FNRS-PDR (PDR-T.0066.18) to A.G-P, the Programme ‘Actions de Recherche Concertée’ 2016-2021 from the ULB and the Fonds d’Encouragement à la Recherche ULB (FER-ULB) to A.G-P. S.R. was and J.C-M. is holding a FRIA (Fund for Research Training in Industry and Agriculture) fellowship from the Fonds de la Recherche Scientifique – FNRS. R.H. is a Research Associate of the Fonds de la Recherche Scientifique – FNRS.