Cyclic GMP controls Rhodospirillum centenum cyst development

Authors


E-mail cbauer@bio.indiana.edu; Tel. (+1) 812 855 6595; Fax (+1) 812 855 6705.

Summary

Adenylyl cyclases are widely distributed across all kingdoms whereas guanylyl cyclases are generally thought to be restricted to eukaryotes. Here we report that the α-proteobacterium Rhodospirillum centenum secretes cGMP when developing cysts and that a guanylyl cyclase deletion strain fails to synthesize cGMP and is defective in cyst formation. The R. centenum cyclase was purified and shown to effectively synthesize cGMP from GTP in vitro, demonstrating that it is a functional guanylyl cyclase. A homologue of the Escherichia coli cAMP receptor protein (CRP) is linked to the guanylyl cyclase and when deleted is deficient in cyst development. Isothermal calorimetry (ITC) and differential scanning fluorimetry (DSF) analyses demonstrate that the recombinant CRP homologue preferentially binds to, and is stabilized by cGMP, but not cAMP. This study thus provides evidence that cGMP has a crucial role in regulating prokaryotic development. The involvement of cGMP in regulating bacterial development has broader implications as several plant-interacting bacteria contain a similar cyclase coupled by the observation that Azospirillum brasilense also synthesizes cGMP when inducing cysts.

Introduction

The cyclic nucleotide second messengers cAMP and cGMP regulate numerous cellular processes across kingdoms, altering the activities of sensor kinases (Francis and Corbin, 1994), ion channels (Finn et al., 1996), phosphodiesterases (PDEs) (Heikaus et al., 2009), guanine nucleotide exchange factors (Quilliam et al., 1995) and transcription factors (Harman, 2001). Their synthesis from ATP and GTP is catalysed by six classes of nucleotidyl cyclases grouped by differences in primary amino acid sequence of catalytic domains. Class I, II, V and VI cyclases are restricted to bacteria, whereas class IV cyclases are also present in archaea (Danchin, 1993; Cotta et al., 1998; Sismeiro et al., 1998; Iyer and Aravind, 2002; Tellez-Sosa et al., 2002). Class III enzymes comprise the majority of adenylyl, and the entirety of guanylyl cyclases, and are present in eukaryotes, bacteria and archaea (Shenroy and Visweswariah, 2004; Schaap, 2005). While numerous cAMP regulated signalling pathways have been elucidated in eukaryotes and bacteria, cGMP regulated processes have only been well defined in eukaryotes.

Even though early research indicated the presence of cGMP in several bacteria, many of these studies are discredited due to difficulty in reproducibility, failure to isolate and biochemically characterize guanylyl cyclases, and the inability to definitively identify examples of cGMP regulated processes. Nevertheless, intracellular cGMP pools have been reported in several bacteria, with fluctuations often coinciding with changes in cellular growth states. For example, the existence of cGMP in Escherichia coli was reported in 1974 (Bernlohr et al., 1974), and later demonstrated to periodically increase from 0.02 to 0.2 pmol mg−1 protein per cell division in synchronized cultures (Cook et al., 1980). Intracellular cGMP in Caulobacter crescentus was also detected and found to decrease from 0.67 to 0.16 pmol mg−1 protein between exponential and stationary growth (Kurn et al., 1977). Similar low levels of cGMP have been reported in an assortment of other bacteria, including Bacillus megaterium (Setlow and Setlow, 1978), Bacillus licheniformis (Bernlohr et al., 1974) Myxococcus xanthus (Ho and McCurdy, 1980), Mycobacterium smegmatis (Bhatnagar et al., 1984) and Bradyrhizobium japonicium (Lim et al., 1979). Genetic evidence appeared to identify the source of cGMP in E. coli as an unintended side activity of adenylyl cyclase (Shibuya et al., 1977). However, partially purified protein fractions exhibiting guanylyl cyclase activity have been reported from cell lysates of E. coli (Macchia et al., 1975; 1981), C. crescentus (Sun et al., 1974), B. licheniformis (Clark and Bernlohr, 1972), M. smegmatis (Bhatnagar et al., 1984), M. xanthus (Devi and McCurdy, 1984) and Nocardia sp. strain NRRL (Son and Rosazza, 2000). In every instance, the identity of the enzyme in question has remained undetermined, precluding genetic and biochemical analyses that could prove the existence of a bona fide guanylyl cyclase in any of the above mentioned species. While intriguing, such studies left significant doubts as to the physiological significance of cGMP in bacteria.

In contrast to early studies that provided inconclusive evidence for cGMP in bacteria, higher intracellular concentrations of cGMP have recently been reported in several cyanobacterial species including Synechocystis PCC 6308 (17–35 pmol mg−1 protein), Plectonema (4.0 pmol mg−1 protein) and Nostoc PCC 8009 (1.0–3.5 pmol mg−1 protein) (Herdman and Elmorjani, 1988). In Synechocystis PCC 6803 cGMP levels are dependent on culture conditions, being relatively low during autotrophic growth (3.5–6.0 pmol mg−1 protein) as compared with mixotrophic and photoheterotrophic growth (∼10 pmol mg−1 protein) or growth in media lacking combined nitrogen (20 pmol mg−1 protein) (Herdman and Elmorjani, 1988). A membrane bound class III cyclase (Cya2) was implicated in cGMP synthesis in Synechocystis PCC 6803 when a cya2 disrupted strain exhibited attenuated intracellular levels of cGMP, but not cAMP (Ochoa De Alda et al., 2000). Biochemical and structural studies of a Cya2 truncated catalytic domain demonstrated that it is indeed a highly specific guanylyl cyclase (Rauch et al., 2008). cGMP has been proposed to regulate UV-B-induced photoacclimation in Synechocystis (Cadoret et al., 2005), although additional genetic and biochemical analyses are needed to verify this hypothesis, as well as to uncover regulators that actually interact with cGMP.

Recently we have undertaken genetic analysis of the complex developmental life cycle present in Rhodospirillum centenum that features three morphologically distinct cell types: swim cells, swarm cells and resting cyst cells (Favinger et al., 1989; Ragatz et al., 1995; Berleman et al., 2004; Berleman and Bauer, 2004; 2005). Metabolically dormant cysts are thought to form as a means of surviving environmental stresses such as desiccation, and have been reported in a wide range of proteobacterial genera. Ultrastructural and morphological characteristics of cyst cell formation have been well studied in the Azotobacter and Azospirillum genera (Stevenson and Socolofsky, 1966; Sadasivan and Neyra, 1987), the latter to which R. centenum is closely related (Fani et al, 1995; Stoffels et al., 2001). R. centenum encystment typically involves nutrient starvation triggering a multi-stage developmental process that includes rounding of cells, production of large intracellular storage granules of poly-hydroxybutyrate (PHB) and the excretion of highly refractive lipopolysaccharide and lipoprotein protective coatings that eventually envelop dormant cyst cells (Berleman and Bauer, 2004). While descriptive features of encystment have been reported for several species, molecular mechanisms governing encystment are poorly understood, certainly not as well understood as regulators that govern Bacillus endospore formation or myxospore development in Myxococcus (Kroos, 2008). In an effort to identify encystment regulatory elements, we undertook a screen for R. centenum mutants that suppressed a previously described hyper-cyst phenotype back to vegetative growth (Berleman et al., 2004). Two insertions fell within a single gene predicted to encode a class III cyclase domain that is shown to be involved in cGMP production required for encystment. Linked to the cyclase are two genes encoding proteins of unknown function that are also required for cGMP production, as well as a homologue of the E. coli transcription factor CRP that preferentially binds cGMP over cAMP, and is essential for inducing cyst formation in R. centenum.

Results

Identification of cyst regulatory elements by hyper-cyst suppression

Our previous screen for regulatory elements that control encystment centred on the isolation of hyper-cyst mutants displaying derepressed cyst formation (Berleman et al., 2004). The hyper-cyst screen successfully uncovered several regulatory elements, including a chemotaxis-like gene cluster (che3) involved in regulating encystment (Berleman et al., 2004; Berleman and Bauer, 2005). However, this screen did not identify downstream DNA binding elements, prompting the development of an alternative screening strategy. In this study, we undertook mini-Tn5 transposon mutagenesis of a Δche3 hyper-cyst strain and screened for mutants that suppressed the hyper-cyst phenotype. Disrupted loci were identified by cloning and sequencing chromosomal segments that flanked the inserted transposon, and then searching against the sequenced R. centenum genome (Lu et al, 2010).

Our hyper-cyst suppression screen generated several mutants that had mini-Tn5 insertions within ORFs encoding putative regulatory elements (Table 1), including two histidine kinase response regulator hybrids (rc1-0896, rc1-3465), a serine/threonine phosphatase (rc1-3466) and a σ70 transcription factor (rc1-1638). Additionally, three insertions were in a region containing a class III nucleotidyl cyclase (ORF rc1-3783) of indeterminate substrate specificity (key substrate discrimination residues are not conserved as indicated in Fig. 1C), multiple ORFs of unknown function and a homologue of the E. coli cAMP receptor protein (CRP) (ORF rc1-3788) (Fig. 1A, B and D). CRP is a founder of the CRP/FNR superfamily of prokaryotic transcription factors that possess an N-terminal cyclic nucleotide-monophosphate-binding domain (cNMP) and a C-terminal helix–turn–helix (HTH) DNA-binding motif. CRP homologues have been shown to directly or indirectly regulate processes such as energy metabolism, development, virulence, quorum sensing and motility in diverse bacteria. All determinant residues of cyclic nucleotide binding to CRP (Weber and Steitz, 1987; Passner et al., 2000; Youn et al., 2007) are identically or functionally conserved in RC1-3788 (residues G90, E91, R100, T101, R141, T145 and S146), indicating that this homologue likely binds a cyclic nucleotide (Fig. 1D). Because of the shared genomic context of the CRP-like transcription factor with the class III nucleotidyl cyclase, we speculated that these proteins may have congruent regulatory roles in controlling encystment. This potentiality is the focus of this study.

Table 1.  ORFs disrupted by transposition screening for hypo-cyst suppressor mutants.
HSM strainLocus tagDescription
Metabolic class  
 44rc1_0155Purine biosynthesis
 41, 52, 142rc1_2014Pyrimidine biosynthesis
 67rc1_1525Pyrimidine biosynthesis
 43, 66, 94, 98, 138rc1_2543Undecaprenyl-phosphate galactose phosphotransferase
 28, 118rc1_2544UDP-glucose-4-epimerase
 26, 30, 68rc1_3992NAD-dependent epimerase/dehydratase
 119rc1_2536Glycosyltransferase
 121, 123, 131, 146rc1_2537Glycosyltransferase
 101rc1_1410Polysaccharide biosynthesis/export protein
Regulatory class  
 69, 114, 115, 122rc1_3465Sensor kinase response regulator hybrid
 117rc1_3466Serine/threonine phosphatase
 90, 93, 97, 105, 108rc1_0896Sensor kinase response regulator hybrid
 70rc1_1638Sigma 70 transcription factor
 55, 76rc1_3783Class III nucleotidyl cyclase
 57rc1_3784/85Intergenic Space
Figure 1.

Genetic context of the guanylyl cyclase-CRP gene cluster.
A. Mini-Tn5 insertion sites are denoted by black triangles.
B. Identified domains in RC1-3783 include an adenylyl/guanylyl cyclase catalytic domain (CYCc) (aa 14–197), a predicted partial ATPase (aa 221–595), followed by seven tetratricopeptide repeats (TPR). Domains in RC1-3788 include a cyclic nucleotide-monophosphate-binding domain (cNMP) (aa 21–139) and a helix–turn–helix (HTH) domain (aa 176–226).
C. Amino acid sequence alignment of the RC1-3783 cyclase domain with representative class III cyclase domains. Species abbreviations are: Rhodospirillum centenum (Rc) RC1-3783 (YP_002299938); Synechocystis sp. PCC 6803 (Sy) Cya2 (NP_440289); Homo sapiens (Hs) GUCY2C (NP_004954); Chlamydomonas reinhardtii (Cr) CYG12 (XP_001700847); Dictyostelium discoideum (Dd) ACRA (AAD50121); Arthrospira platensis (Ap) CyaC (BAA22997) and CyaG (BAB19924); Mycobacterium tuberculosis H37Rv (Mt) Rv1264 (NP_215780). Positions of residues involved in metal binding (yellow), substrate specification (blue) and transition state stabilization (red) are indicated by asterisks (*), inverted triangles (▾) and arrows (↓). Shading indicates identically and functionally conserved residues at 100% (black), 87.5% or 75% (dark grey shading with white letters) and 62.5% (light grey shading with dark letters) conservation.
D. Amino acid sequence alignment of RC1-3788 and representative CRP homologous. Sequences and species abbreviations used are as follows: Rhodospirillum centenum (Rc) RC1-3783 (YP_002299938) Mycobacterium tuberculosis (Mt) Rv3676 (NP_218193); Escherichia coli (Ec) CRP (NP_417816), Streptomyces coelicolor (Sc) CRPSco (NP_627768), Synechocystis PCC 6803 (Sy) SyCRP1 (NP_440289). The alignment was constructed in MEGA v.4.0 utilizing a BLOSUM matrix, with respective pair-wise and multiple alignment gap opening and extension penalties of 20 and 0.2. Intra-subunit E. coli CRP residues which directly contact cAMP are shaded red and denoted by an asterisk (*) and a single inter-subunit contact (S128) is shaded blue and marked by an inverted triangle (▾). Residues which make direct contacts with DNA shaded yellow and denoted by arrows (↓). Additional shading indicates identically and functionally conserved residues at 100% (black) and 80% (dark grey) conservation.

Deletion of the nucleotidyl cyclase and CRP homologue disrupts cyst formation

To address the function of the nucleotidyl cyclase (rc1-3783) and CRP homologue (rc1-3788) in cyst cell development, we constructed in-frame deletions of these genes in a wild-type background. Wild-type and deletion mutants were then phenotyped with respect to encystment by plating on vegetative (CENS) and cyst-inducing (CENS-8xN) culture media. When grown on CENS medium, wild-type and deletion strains (Δrc1-3783, Δrc1-3788) developed shiny, convex colonies that contained vegetative cells (Fig. 2A and B). However, when grown on cyst-inducing CENS-8xN medium the wild-type strain rapidly underwent encystment, as evidenced by the formation of dry intricately ridged colonies that contain many round, refractive cyst cells (Fig. 2A and B). In stark contrast, strains deleted for the nucleotidyl cyclase (Δrc1-3783) and CRP homologue (Δrc1-3788) exhibited shiny vegetative colonies and vibroid shaped cells on cyst-inducing medium (Fig. 2A and B). We also quantified encystment by assaying the percentage of viable cells that exhibit desiccation resistance, a trait only exhibited by cyst cells (Berleman and Bauer, 2004). When grown for 72 h on vegetative cyst-inducing CENS-8xN medium ∼1% of viable wild-type cells were desiccation resistant, while cells of strains deleted for the nucleotidyl cyclase or the CRP homologue exhibited no assayable desiccation resistance (Fig. 2C).

Figure 2.

Analyses cyst cell formation in wild-type, Δrc1-3783 (nucleotidyl cyclase) and Δrc1-3788 (CRP homologue) strains after 3-day growth on agar solidified cyst-inducing CENS-8xN media.
A. Wild-type colonies formed dry/ridged cyst containing colonies under all growth conditions whereas a strain deleted for the cyclase (Δrc1-3783) only formed cyst colonies in the presence of 50 µM cGMP. A strain deleted for the CRP homologue (Δrc1-3788) only formed shiny vegetative colonies.
B. Microscopic observations of cyst and vegetative cells within the above-shown colonies.
C. Desiccation resistant quantification of cyst cells in the above colonies. The double asterisk (**) indicates that no cysts were detected in any cell dilution assayed.

cGMP is a signal controlling encystment

We assessed whether R. centenum is capable of producing cGMP when undergoing cyst cell differentiation by assaying for the presence of cGMP during growth in butyrate minimal liquid medium that effectively induces cyst formation (Statwald-Demchick et al., 1990). The results show significant extracellular accumulation of cGMP in wild-type cultures that are undergoing cyst formation up to 33.0 nM (Fig. 3). This is contrasted by strains deleted for the nucleotidyl cyclase (Δrc1-3783) or the CRP homologue (Δrc1-3788) that fail to form cysts and have < 1 nM cGMP production (Fig. 3). Similar measurements also showed that cAMP levels did not significantly accumulate during cyst induction in either the wild-type or deletion strains (data not shown). We also determined whether cGMP accumulates intracellularly by assaying cell-free extracts derived from washed cell pellets. Intracellular cGMP levels in cells from 11-day-old cultures were measured at 13.2, 0.4 and 5.2 pmol mg−1 of total protein in respective cell pellets of the wild-type, Δrc1-3783 and Δrc1-3788 strains.

Figure 3.

Accumulation of exogenous cGMP in cyst-inducing liquid CENBA medium by wild-type cells (grey bars), the cyclase deletion strain Δrc1-3783 (black bars) and the CRP homologue strain Δrc1-3788 (white bars). While cGMP accumulates to appreciable levels in the wild-type strain, in the Δrc1-3783 and Δrc1-3788 strains levels remain < 1 nM.

Previous studies have shown that addition of exogenous cAMP is capable of complementing disruptions of microbial adenylyl cyclases (Terauchi and Ohmori, 1999; Roberts et al., 2007). We therefore attempted to complement the cyst deficient (hypo-cyst) phenotype exhibited by deletions of the nucleotidyl cyclase and CRP homologues by growing cells on CENS-8xN encystment media supplemented with cAMP or cGMP. When grown on cyst-inducing CENS-8xN medium, the wild-type strain formed cysts after 3-day growth under all assayed conditions (Fig. 2). In contrast, the strain deleted for the cyclase (Δrc1-3783) failed to encyst, except when supplemented with cGMP at a final concentration of 50 µM, which allowed production of wild-type levels of cysts (Fig. 2). Although Fig. 2 shows complementation with 50 µM cGMP, effective complementation of the cyclase deletion also occurs with as little as 5 µM cGMP (data not shown). Addition of 50 µM cAMP, or any other adenine or guanine nucleotide (AMP, ADP, ATP, GMP, GDP and GTP), did not promote induction of cysts in the cyclase mutant (Fig. 2; Fig. S1). With respect to the CRP homologue mutant (Δrc1-3788), there was no discernable response to the addition of cGMP nor to any other nucleotide (cAMP, AMP, ADP, ATP, GMP, GDP and GTP), as colonies from these plates were identical in appearance to those grown in the absence of nucleotides and contained only vegetative cells (Fig. 2; Fig. S1). Interestingly, cyst formation is also rapidly induced in both the wild-type and cyclase mutant strains by the addition of 200 µM cGMP to vegetative CENS media, which is a growth medium that normally does not induce cyst formation (Fig. 4). Again, the strain deleted for the CRP homologue (Δrc1-3788) remained unable to induce development of cysts (Fig. 4).

Figure 4.

Analyses of cyst cell formation in wild-type, Δrc1-3783 (nucleotidyl cyclase) and Δrc1-3788 (CRP homologue) strains after 3-day growth on agar solidified CENS vegetative media. Media supplemented with cGMP had a final concentration of 200 µM.
A. Top panel shows shiny vegetative colonies forming on CENS media whereas the bottom panel shows dry ridged cyst-forming colonies from wild-type and Δrc1-3783 strains grown on CENS medium containing 200 µM cGMP.
B. Microscopic observations of cyst and vegetative cells from the above panels.

ORF rc1-3783 encodes a guanylyl cyclase

An alignment of the catalytic core of the R. centenum cyclase with previously characterized eukaryotic and prokaryotic cyclases shows that several key catalytic amino acids are not present in the R. centenum cyclase (Fig. 1C). Specifically, while two critical metal-ion binding aspartates are conserved (D41, D85), the transition state-stabilizing asparagine and arginine residues are substituted by histidine (H157) and lysine (K161). In addition, glutamine (Q81) and proline (P157) residues occupy substrate-specifying positions, neither of which is similar to residues that are classic predictors of purine selectivity (Linder, 2005). The observed cellular response to cGMP, coupled with the presence of ambiguous residues at substrate-specifying positions, prompted us to ascertain substrate specificity and product formation by the R. centenum cyclase coded by ORF rc1-3783. For this analysis we expressed and purified a His-tagged version of this cyclase using an E. coli expression strain, and then used high-pressure liquid chromatography (HPLC) to resolve adenine and guanine nucleotide substrates and products from enzymatic assays. The ability of the recombinant cyclase to utilize either ATP or GTP as a substrate was tested in the presence of either Mg2+ or Mn2+, with substrate and product retention times compared with nucleotide standards (Fig. 5A and E). Synthesis of cAMP was not observed in reactions containing ATP and Mg2+ (Fig. 5C), whereas synthesis of cGMP was observed in an identical assay using GTP as a substrate (Fig. 5G). In reactions containing Mn2+ the production of cGMP was greatly enhanced (Fig. 5H), with cGMP increasing concomitantly with GTP substrate reduction over time. In a reaction containing ATP and Mn2+ a small peak corresponding to cAMP was observed (Fig. 5D), although cAMP formation was significantly less than the amount of cGMP produced in the presence of either Mg2+ or Mn2+. Small amounts of nucleotide mono and diphosphates observed in all assays were attributed to substrate degradation. Prolonged incubation of GTP at elevated temperatures has been reported to cause non-enzymatic formation of cGMP (Kimura and Murad, 1974). Since our reactions were terminated by heat denaturation, we addressed whether heat significantly contributed to spontaneous cGMP formation with a control reaction lacking enzyme, which produced no detectable cGMP (Fig. 5D).

Figure 5.

HPLC separation of products and substrates from assays using purified recombinant guanylyl cyclase (RC1-3783).
A–H. (A) Separation of adenine nucleotide standards AMP (1), ADP (2), cAMP (3) and ATP (4). (E) Separation of guanine nucleotide standards GMP (1), GDP (2), cGMP (3) and GTP (4). All reactions were incubated for 2 h and contained 0.5 mM ATP (B–D) or GTP (F–H), 10 mM MnCl2 (B, D, F, H) or MgCl2 (C, G) and 0.5 µg µl−1 purified RC1-3783 (C, D, G, H). Control reactions (B, F) had buffer added in place of enzyme. A small amount of cGMP indicated by a black arrow (↓) is produced in reaction G-containing GTP, Mg2+ and recombinant cyclase, with greater activity occurring in reaction H when Mn2+ is substituted for Mg2+.
I. Purified recombinant cyclase elutes as a dimer in the absence of NaCl (blue line) but as a monomer when chromatographed on a Superose 6 column equilibrated in 100, 50 or 25 mM NaCl (black, green and red lines respectively).

We observed that cyclase activity was salt inhibited, as reactions lacking NaCl produced ∼3 × more cGMP than those containing 100 mM NaCl (data not shown). Since class III cyclases are active as dimers, we assessed the effect of salt on oligomerization by size exclusion chromatography, which showed that purified cyclase eluted as a monomer in columns equilibrated with 25, 50 and 100 mM NaCl, and as a dimer in the absence of NaCl (Fig. 5I).

The CRP homologue RC1-3788 preferentially binds cGMP

We also characterized the cyclic nucleotide binding abilities of the CRP-like homologue coded by ORF rc1-3788. For this analysis the CRP homologue was purified after overexpression from an E. coli expression strain and assayed for binding of cyclic nucleotides using both differential scanning fluorimetry (DSF) and isothermal titration calorimetry (ITC). DSF measures protein thermal denaturation, or melting temperature (Tm), which is often stabilized upon binding to a substrate (Matulis et al., 2005; Niesen et al., 2007). Using this technique the Tm of the CRP homologue in the absence of a ligand was measured at 67°C. The thermal stability of RC1-3788 was significantly and maximally enhanced by the addition of 40 µM cGMP, which increased the overall Tm to 82°C (Fig. S2; Table 2). In stark contrast, an equivalent concentration of cAMP increased the overall Tm by only 1.5°C (Fig. S2; Table 2).

Table 2.  Changes in melting temperature (Tm) of the CRP homologue RC1-3788 in the presence of increasing cAMP or cGMP relative to a sample lacking ligand, as determined by DSF.
Concentration of ligand (µM)ΔTm (oC)ΔTm (oC)
RC1_3788 + cGMPRC1_3788 + cAMP
 000
108.50.5
2012.00.9
3014.01.5
4014.41.5
5014.42.0

Using ITC analysis for substrate binding, we observed a biphasic titration curve by the addition of cGMP (Fig. S3; Table 3). The initial binding event is characterized by exothermic enthalpy (ΔH = −0.9 ± 0.2) followed by an endothermic event (ΔH = 7.2 ± 0.2). The observed raw data were best fitted to a two-site sequential model, described in Experimental procedures. Binding of cGMP exhibited a negative cooperativity (Kd1 < Kd2) with a stoichiometry of two cGMP per peptide dimer and apparent Ka1 (2.6 ±  0.6 × 107 M−1) and Ka2 (0.8 ± 0.2 × 107 M−1) (Table 3) similar to those observed for the binding of cAMP to E. coli CRP (Lin and Lee, 2002a). Indeed the exothermic and endothermic binding events, negative cooperativity and Ka values are surprisingly similar to those previously reported for cAMP binding to CRP from E. coli (Gorshkova et al., 1995).

Table 3.  Binding affinities of CRP homologue RC1-3788 for cAMP and cGMP, as determined by ITC.
LigandModelnΔHKobsΔSΔG
(kcal mol−1)(× 107 M−1)(kcal mol−1 K−1)(kcal mole−1)
  • a. 

    Assumes an integral stoichiometry value of n = 1.

  • b. 

    According to this model, n1 = n2 = 1.

cAMPOne-site binding0.5 ± 0.033.3 ± 0.40.03 ± 0.010.04 ± 0.4−7.5 ± 0.2
cGMPTwo-site1b−0.9 ± 0.22.6 ± 0.60.03 ± 0.2−10.1 ± 0.1
Sequential bindinga1b7.2 ± 0.20.8 ± 0.20.1 ± 0.3−9.4 ± 0.1

Interestingly, the R. centenum CRP homologue exhibits slight, albeit much poorer, binding to cAMP with a binding constant (Ka) of 0.03 ± 0.01 × 107 M−1 (Fig. S3; Table 3). E. coli CRP also exhibits the capability of binding to cGMP, but at a much reduced affinity to that of cAMP (Gorshkova et al., 1995; Lin and Lee, 2002b). Therefore, the R. centenum CRP-like homologue and E. coli CRP bind both cAMP and cGMP, with the caveat that R. centenum preferentially binds cGMP over that of cAMP while E. coli CRP preferentially binds cAMP over cGMP. Thus, these homologues appear to have ‘swapped’ cyclic nucleotide substrate specificities.

The nucleotidyl cyclase gene cluster, and the ability to produce cGMP, is conserved in other soil inhabiting species

Searching the NCBI database, we discovered homologues of the R. centenum guanylyl cyclase, and of ORF rc1-3786 and ORF rc1-3787, clustered in several plant associated soil bacteria (Fig. 6A). Interestingly, Azospirillum sp. B510, which has a very similar life cycle as R. centenum (Berleman and Bauer, 2004), has linked homologues of the R. centenum cyclase, ORFs rc1-3786 and rc1-3787 as well as a homologue to the CRP family transcription factor coded by ORF rc1-3788 (Fig. 6A). Conservation of this gene cluster suggests that other soil inhabiting species may also produce cGMP and that ORFs rc1-3786 and rc1-3787 may be involved in regulating the production, transport or stability of cGMP. To test these possibilities we constructed in-frame deletions of ORFs rc1-3786 and rc1-3787 in R. centenum and assayed these mutants for their ability to produce cysts. Deletions of either rc1-3786 or rc1-3787 results in a phenotype that is indistinguishable from the strain deleted for the cyclase. Specifically, these additional deletion strains are defective in forming cysts on CENS-8xN cyst-inducing medium, and this defect can be complemented by exogenous addition of cGMP at a final concentration of 50 µM (Fig. 7A and B). Furthermore, deletion of either of these ORFs also results in defects in cGMP production when grown in liquid cyst-inducing CENBA media (Fig. 7C). Thus, in vivo production of cGMP in R. centenum requires three gene products coded by ORFs rc1-3783, rc1-3786 and rc1-3787.

Figure 6.

A. Species containing cyclases with similar genetic contexts as the R. centenum guanylyl cyclase coded by gcyA (rc1-3783), the downstream ORFs needed for cyclase activity (rc1-3786 and rc1-3787) and the CRP homologue coded by cgrA (rc1-3788) in Azosprillum sp. B510. Red (nucleotidyl cyclase), green (COG1944), orange (COG3482) and blue (CRP homologue).
B. Amino acid sequence alignment of the RC1-3783 cyclase domain with putative guanylyl cyclase domains. Species are: Rhodospirillum centenum, RC1-3783 (YP_002299938); Azospirillum sp. B510, AZL_d00190 (YP_003452389); Sinorhizobium meliloti 1021, SMc01491 (NP_386201); Mesorhizobium loti MAFF303099, mll0576 (NP_102350); Rhizobium leguminosarum bv. viciae 3841, pRL110256 (YP_771288); Rhizobium etli CIAT 652, RHECIAT_PA0000121 (YP_001985730); Rhizobium sp. NGR234, NGR_c20030 (YP_002826519); Sinorhizobium medicae WSM419, Smed_1991 (YP_001327660). Domains were identified using the SMART database and aligned in MEGA v.4.0 utilizing a PAM matrix with respective pair-wise and multiple alignment gap opening and extension penalties of 10 and 0.1. Relative positions of residues involved in metal binding (yellow), substrate specification (blue) and transition state stabilization (red) are additionally indicated by asterisks (*), inverted triangles (▾) and arrows (↓). Further shading indicates identically and functionally conserved residues at 100% (black), 80% (dark grey, white letters) and 60% (light grey, dark letters) conservation.

Figure 7.

Analyses of cyst cell formation and cGMP production in wild-type, Δrc1-3786 and Δrc1-3787 deletion strains.
A. After 3-day growth on cyst-inducing CENS-8xN media the wild-type strain displayed ridged colony morphology indicative of cyst cell formation, while both deletion strains remained vegetative in appearance. In contrast, on a plate supplemented to 50 µM cGMP, all three strains displayed encysted colony morphologies.
B. Microscopic observations of vegetative and cyst cells from colonies shown in (A).
C. Exogenous cGMP levels present in culture supernatants after 11-day growth in liquid CENBA media.

The presence of linked homologues of rc1-3783, rc1-3786 and rc1-3787 in the genomes of several species that interact with plant roots suggest that the ability to produce cGMP may be a conserved trait of this bacterial niche. To test this possibility, we assayed for cGMP production in Azospirillum brasilense cultures that were grown in cyst-inducing MSM flocculation medium (Neyra and Van Berkum, 1977). We observed accumulation of up to ∼4 nM of cGMP in the culture supernatant that was concurrent with the formation of cysts in this species (Fig. 8).

Figure 8.

Accumulation of exogenous cGMP in culture supernatants of wild-type Azospirillum brasilense grown in cyst-inducing MSM medium. Cultures were grown aerobically in MSM media and the culture supernatant was measured for cGMP levels as described in Experimental procedures. Error bars represent standard deviation derived from three replicate assays.

Discussion

While cAMP is common in all three kingdoms, the existence of cGMP has only been reported for a few bacteria (Bernlohr et al., 1974; Kurn et al., 1977; Setlow and Setlow, 1978; Lim et al., 1979; Ho and McCurdy, 1980; Bhatnagar et al., 1984). There is also scant biochemical evidence as to any role cGMP has in controlling a prokaryotic cellular process, thus the importance of cGMP in bacteria has remained controversial. The only other example of a biochemically characterized bona fide prokaryotic guanylyl cyclase is a class III enzyme from Synechocystis PCC 6803 that has been crystallized, but as yet has an ill-defined role (Rauch et al., 2008). Our study of the role of cGMP in controlling R. centenum encystment thus represents the first example of prokaryote that produces cGMP with a clearly defined role in controlling a prokaryotic process. As such, we designate ORF rc1-3783 as gycA for guanyl cyclase and ORF rc1-3788 as cgrA for cyclic GMP regulator.

Rhodospirillum centenum secretes large quantities of cGMP into the extracellular environment as it transitions to cyst formation. Interestingly, cAMP is secreted by several bacterial species; for example, E. coli secretes > 99.9% of produced cAMP under certain conditions (Matin and Matin, 1982), the reason why this occurs remains unclear. Cyanobacteria also secrete cAMP, including Spirulina platensis (Sakamoto et al., 1991), which displays increased mat formation and gliding motility in response to exogenously added cAMP (Ohmori et al., 1992). In Streptomyces coelicolor, extracellular cAMP peaks during spore germination and again during aerial mycelium and actinorhodin production suggesting that cAMP may be a diffusible signal that co-ordinates production of this antibiotic (Susstrunk et al., 1998).

Our working model for cGMP signalling in R. centenum involves the synthesis of cGMP in response to a signal related to nutrient deprivation that activates the guanylyl cyclase (Fig. 9). The mechanism by which the R. centenum cyclase is activated is not immediately apparent, as the only other predicted domain present is a partial ATPase domain of the COG3899 functional group and several TPR protein–protein interaction domains at the C-terminus. While domains of known function are not present in the products of ORFs rc1-3786 and rc1-3787, it seems likely that they interact and form a complex with the cyclase as disruptions of either of these ORFs gives rise to the same hypo-cyst phenotype that, like the guanylyl cyclase-disrupted strain, can be rescued by cGMP. The majority of cGMP produced by the guanylyl cyclase appears to be secreted, and is presumably imported back into R. centenum via an unknown transport mechanism to allow intracellular cGMP levels to rise to a level where it binds to the CRP homologue to activate genes required for encystment. That most of the synthesized cGMP is secreted also indicates that cGMP production likely functions as a signal for a co-ordinated community wide change in cell type.

Figure 9.

Model of the regulation of R. centenum encystment by cGMP. The guanylyl cyclase coded by gcyA (rc1-3783) synthesizes cGMP in response to an unknown developmental signal. The cNMP-binding domain of the CRP homologue coded by cgrA (rc1-3788) binds and is activated by cGMP, resulting in expression of genes required.

Bioinformatic approaches have uncovered putative prokaryotic guanylyl cyclases similar to Cya2 of Synechocystis, in other cyanobacteria and more diverse species including Congregibacter litoralis, Mariprofundus ferrooxydans, Roseobacter denitrificans (γ- and α-proteobacteria respectively) and Mycobacterium ulcerans (an actinobacterium) (Rauch et al., 2008; Wu et al., 2008; Biswas et al., 2009). A constraining parameter in the identification of these enzymes is the presence of the substrate-specifying glutamate residue, which was a unifying feature of all previously characterized guanylyl cyclases. However, the R. centenum guanylyl cyclase contains a glutamine (Q81) substituted for the substrate-specifying glutamate and a proline (P150) at a second substrate binding position. Likewise, asparagine and arginine residues thought to be involved in transition state stabilization are replaced by histidine (H157) and lysine (K161), respectively, in the R. centenum cyclase. That these changes are present in the R. centenum guanylyl cyclase highlights the fact that there is an incomplete understanding of substrate specification in class III cyclases. Indeed, our demonstration that one of these non-canonical cyclases is a second identified bona fide prokaryotic guanylyl cyclase suggests that prokaryotic guanylyl cyclases may be more common than previously believed, especially given that there are abundant uncharacterized microbial cyclases that exhibit diverse substitutions in substrate-specifying residues (Shenroy and Visweswariah, 2004).

By performing genomic context searches using the two R. centenum ORFs rc1-3786 and rc1-3787, which are needed to observe in vivo cGMP production, we are intrigued to find that linked homologues of these genes are associated with nucleotidyl cyclases in the genomes of several sequenced Rhizobial species as well as in Azospirillum that produces cysts (Fig. 6A). An alignment of the catalytic cores from these cyclases revealed a variety of canonical catalytic residue deviations (Fig. 6B), with cyclases from Sinorhizobium meliloti (SMc01491), Sinorhizobium medicae (Smed_1991) and Rhizobium sp. NGR234 having a proline substitution at the second substrate-specifying position like the R. centenum cyclase. We have demonstrated that A. brasilense synthesizes and excretes cGMP as cells transition from vegetative to cyst cells (Fig. S4), suggesting that the cyst developmental cycle of this related species may be similarly regulated by cGMP. We have also observed that S. meliloti is also capable of synthesizing cGMP (data not shown), and it has been recently reported that this species is capable of making desiccation resistant cysts not unlike that of R. centenum and A. brasilense (Pogorelova et al., 2009). This suggests that the use of cGMP as a signal to control cyst formation may be a conserved trait among several species that interact with plants.

Experimental procedures

Strains and growth conditions

Escherichia coli strains DH5α, DH10B and SCS1101 were used for general cloning, BL21 Rosetta II (Invitrogen) for protein overexpression and S17-1 (λpir) for plasmid conjugation. E. coli strains were grown and cultured on agar-solidified or liquid Luria–Bertani (LB) media at 37°C or 16°C with appropriate antibiotics. The parental R. centenum strain used in this study was a Δche3 derivative of wild-type R. centenum (ATCC 51521). R. centenum strains were cultured aerobically in CENS or CENBA liquid media (Statwald-Demchick et al., 1990) at 37°C, or grown at 42°C on agar-solidified CENS or CENS-8xN, which is CENS supplemented with 8 mM NH4Cl. A. brasilense (ATCC 29145) cells were grown on agar-solidified NB, or cultured aerobically in liquid NB or MSM media (Neyra and Van Berkum, 1977) at 37°C.

Desiccation resistance analyses

Rhodospirillum centenum colonies were harvested from CENS-8xN plates after 3-day growth and resuspended in 1 ml of phosphate buffer. Cell suspensions were sonicated 2 × for 5 s at low power to disperse cellular aggregates as needed. Total viable cell counts were enumerated from serial dilutions pipetted onto CENS plates incubated at 42°C for 2 days. Cysts cells were quantified by pipetting identical dilutions onto 0.45-µm-pore-sized filters (Grade: A045FO47A; Advantec), which were dried briefly, then desiccated at 42°C for 3 days. Filters were subsequently transferred to CENS plates and incubated at 42°C for 2 days to allow surviving cell outgrowth. All assays were repeated in triplicate, and desiccation resistance was calculated as total viable cells/total desiccation-resistant cells.

Hypo-cyst suppression screen

Transposon mutagenesis and the construction of in-frame deletion mutants of rc1-3783, rc1-3786, rc1-3787 and rc1-3788 were performed as described by Berleman et al. (2004). Mutagenesis of the Δche3 strain was performed as previously described by Berleman and Bauer (2004) and Berleman et al. (2004). Briefly, a modified mini-Tn5 spectinomycin-resistant (Spr) interposon was delivered to a hyper-cyst Δche3 R. centenum strain (Berleman et al., 2004) on plasmid pZJD17 from E. coli S17-1 (λS17) using a filter mating procedure. Overnight R. centenum and E. coli cultures were washed 3× with CENS to remove antibiotics, then were applied in a 5:1 ratio to a 0.45-µ-pore-sized filter (No. 245-0045; Nalgene). After 4 h incubation at 37°C on an antibiotic-free CENS plate, cells were resuspended in 5 ml of CENS and spread onto CENSKmSp plates in 200 µl aliquots. Spr transconjugates displaying suppression to vegetative growth after 72 h were selected for further scrutiny of colonial encystment morphologies. Cells from overnight cultures were washed 3 × and resuspended in 1/20 volume of phosphate buffer (40 mM KH2PO4/K2HPO4, pH 7), pipetted in 5 µl aliquots onto CENS and CENBA plates that were incubated for 3 and 6 days respectively.

Interposon insertion loci from mutants growing vegetatively on both media were mapped on the chromosome by isolating and digesting mutant genomic DNA with the PstI restriction enzyme and ligating fragments into the corresponding site in pBluescript SK+. Plasmids were electroporated into E. coli DH10B and transformants with a chromosomal DNA fragment containing a Spr interposon were selected for by growth on LBAmpSp. Transformant plasmid DNA was used in sequencing reactions with primers specific to the 5′ and 3′ ends of the interposon (Table S1). Resultant DNA sequences were then used in blast searches against the R. centenum genome to identify disrupted loci. Domains in RC1-3783 and RC1-3788 were identified by SMART and NCBI Conserved Domain Databases, with the TPR domains in RC1-3783 identified using TPRpred.

Quantification of cyclic nucleotides in bacterial cultures

Wild-type, Δrc1-3783, Δrc1-3786, Δrc1-3787 and Δrc1-3788 R. centenum cells taken from agar-solidified CENS plates were resuspended in phosphate buffer to an OD600 of ∼2.0, 5 µl of which was then subcultured into 25 ml of liquid CENBA medium and cultured at 37°C with shaking at 250 r.p.m. Measurements of extracellular cGMP produced in wild-type, Δrc1-3783 and Δrc1-3788 cells were from supernatants of liquid cultures. Culture supernatants were treated with 1/100 volume of concentrated HCl, centrifuged at 13 000 r.p.m. at room temperature and directly assayed by EIA. Cell pellets were washed 2 × and resuspended in 0.5 ml of phosphate buffer, lysed by sonication and then clarified by centrifugation. The supernatant was mixed with an equal volume of 10% ice-cold trichloroacetic acid (TCA) and kept on ice for 30 min with TCA-insoluble material removed by centrifugation. TCA was then extracted from the supernatant by three successive washes with 2 vols of diethyl ether. The final aqueous phase was lyophilized and resuspended in 450 µl of 0.1 M HCl with concentrations of cAMP and cGMP determined with commercially available immunoassay kits (AssayDesigns).

In-frame deletions of rc1-3783, rc1-3786, rc1-3787 and rc1-3788

In-frame deletion mutants were constructed using a modified sucrose selection technique, performed as described previously (Berleman et al., 2004; Masuda and Bauer, 2004; Blomfield et al., 1991). Gene-flanking 500 bp fragments were PCR-amplified using wild-type cells as template for colony PCR and primer sets detailed in Table S1. PCR-amplified fragments were separately cloned and sequenced in either pTOPO Blunt-II or pGEM-T vectors. The rc1-3783 and rc1-3788 fragments were assembled in pUC18 prior to being subcloned into the suicide vector pZJD29a, whereas fragments for the rc1-3786 and rc1-3787 constructs were ligated directly into pZJD29a. Plasmids were mated into wild-type R. centenum from E. coli S17-1 (λpir), and initial recombinants were selected for on CENSGm agar plates. Second recombinants carrying gene deletions were later identified through phenotypic (GmS/SucR) and colony PCR analyses.

Cloning, overexpression and purification RC1-3783 and RC1-3788

rc1-3783 was amplified in two separate fragments with primers rc1-3783NdeI-f, rc1-3783SbfI-r, rc1-3783SbfI-f and rc1-3783XhoI-r, whereas rc1-3788 was cloned as one fragment with the primers rc1-3788NdeI-f and rc1-3788XhoI-r (Table S1). PCR fragments were gel-purified, cloned and sequenced in pTOPO-Blunt II. Fragments where then subcloned into pET28a vectors in ligation reactions using the external NdeI and XhoI restriction sites, and were internally joined in rc1-3783 by a SbfI site common to both fragments. Each construct was sequenced and transformed into E. coli Rosetta II (DE3).

For overexpression of RC1-3783, an overnight culture of the Rosetta II cells containing the RC1-3783 expression plasmid was subcultured 1/50 into 2 l of LBKmCm, grown to an OD600 ∼ 0.6, then induced at 0.2 mM IPTG and grown overnight at 16°C. Cells were pelleted by centrifugation at 6000 r.p.m. for 5 min, washed once with 20 mM Tris 100 mM NaCl pH 8.0, then repelleted and resuspended in 40 ml of binding buffer (20 mM Tris, 100 mM NaCl, 25 mM Imidazole, pH 8). Cells were lysed by four passes through an M-110L Micro Fluidizer Processor at ∼2000 p.s.i. (Microfluidics). Insoluble matter was removed by centrifugation at 15 000 r.p.m. for 30 min and the decanted supernatant was incubated with 4 ml of Ni2+ charged resin (Novagen) with gentle shaking for ∼2 h. After 5 min of centrifugation at 1000 r.p.m., the pelleted resin was washed 3 × with ∼45 ml of wash buffer (20 mM TrisHCl, 100 mM NaCl, 50 mM Imidazole, pH 8) and then loaded onto a 25 ml disposable column (Bio-Rad). The resin was washed with an additional 20 ml of wash buffer and bound protein was then eluted with elution buffer (20 mM TrisHCl, 100 mM NaCl, 200 mM Imidazole, pH 8). Elution fractions were concentrated in a 30k Centriprep Concentrator (Millipore), then was loaded onto a hand-packed Superose 12 size exclusion column (GE Healthcare) equilibrated in binding buffer lacking imidazole. After concentration, the protein was dialyzed against storage buffer (20 mM Tris-HCl, 100 mM NaCl, 50% glycerol, pH 8) and stored at −20°C until use.

Overexpression and purification conditions for RC1-3788 were essentially identical to those of RC1-3783, with the exception that all buffers had a pH 7.5, and the size exclusion purification step differed. The size exclusion equilibration buffer additionally contained 5% glycerol when RC1-3788 was to be used for DSF analyses, whereas a separate buffer (50 mM KH2PO4/K2HPO4, 200 mM KCl, 0.2 mM Na2EDTA, 0.2 mM DTT, 5% glycerol, pH 7) was used for protein to be used in ITC experiments. RC1-3788 was stored at 4°C and used within 1 week of purification.

Adenylyl and guanylyl cyclase assays

Adenylyl and guanylyl cyclase reactions were undertaken in 100 µl of reactions containing 0.5 µg µl−1 purified RC1-3783, 20 mM Tris (pH 8.0), 100 mM NaCl, 0.5 mM ATP or GTP and 10 mM MnCl2 or MgCl2. Reactions were incubated at 25°C, stopped by heating at 75°C for 10 min and then clarified by centrifugation at 15 000 r.p.m. for 10 min. HPLC separation of nucleotides was accomplished as described by Gebelein et al. (1992).

ITC and DSF determination of RC1-3788 cyclic nucleotide binding

Sodium salts of cAMP and cGMP (Sigma) were dissolved in RC1-3788 storage buffer and quantified using molar coefficients of 1.23 × 104 cm M−1 at 260 nm for cAMP and 1.34 × 104 cm M−1 at 250 nm for cGMP. All reactions contained 10 µM RC1-3788, 2.5 × SYPRO Orange dye (Cat. S6650; Invitrogen) and cyclic nucleotide concentrations of 0–80 µM (cAMP) or 0–50 µM (cGMP) tested in at least three independent assays with control reactions of protein alone, protein without ligand and ligand alone. DSF experiments were performed using an Eppendorf Mastercycler EP Realplex thermal cycler, with excitation and emission at 470 and 550 nm respectively. Denaturation curves were generated as a function of [–R′(T)] for every °C between 25°C and 95°C, and Tm values.

For ITC analyses, both RC1-3788 and cyclic nucleotide stocks were equilibrated in degassed assay buffer (50 mM KH2PO4/K2HPO4, 200 mM KCl, 0.2 mM Na2EDTA, 0.2 mM DTT, 5% glycerol, pH 7). All experiments were performed at 25°C using a Microcal VP-ITC calorimeter with a stirring speed of 310 r.p.m. A minimum of two titrations were carried out for each ligand and the experimental values reported are the average of the individual best-fit values collected using a one-site or sequential binding model (ITC Data Analysis in Origin® v.7). The best fit to data produces a binding constant (Ka), enthalpy (ΔH), entropy (ΔS) and reaction stoichiometry (n). ΔG and ΔS were then calculated using thermodynamic definition, ΔG = −RT ln K and ΔG = ΔH − TΔS. The observed raw data were best described by the following reactions:

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Acknowledgements

We thank Dr Yves V. Brun for the use of his microscope facility; Dr Todd Stone, Facility Manager of the Indiana University Physical Biochemistry Instrumentation Facility and Dr Jonathan A. Karty, Facility Manager of the Indiana University Chemistry Department Mass Spectrometry Facility, for training and assistance with instrumentation. This work was supported by the Metabolomics and Cytomics Initiative (MetaCyte), which is supported by a major grant from the Lilly Endowment. The second (Q.D.) and third (S.R.) authors contributed equally to this study.

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