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Response regulator PorX coordinates oligonucleotide signalling and gene expression to control the secretion of virulence factors

Abstract The PglZ family of proteins belongs to the alkaline phosphatase superfamily, which consists of metallohydrolases with limited sequence identity but similar metal-coordination architectures in otherwise divergent active sites. Proteins with a well-defined PglZ domain are ubiquitous among prokaryotes as essential components of BREX phage defence systems and two-component systems (TCSs). Whereas other members of the alkaline phosphatase superfamily are well characterized, the activity, structure and biological function of PglZ family proteins remain unclear. We therefore investigated the structure and function of PorX, an orphan response regulator of the Porphyromonas gingivalis TCS containing a putative PglZ effector domain. The crystal structure of PorX revealed a canonical receiver domain, a helical bundle, and an unprecedented PglZ domain, similar to the general organization of the phylogenetically related BREX-PglZ proteins. The PglZ domain of PorX features an active site cleft suitable for large substrates. An extensive search for substrates revealed that PorX is a phosphodiesterase that acts on cyclic and linear oligonucleotides, including signalling molecules such as cyclic oligoadenylates. These results, combined with mutagenesis, biophysical and enzymatic analysis, suggest that PorX coordinates oligonucleotide signalling pathways and indirectly regulates gene expression to control the secretion of virulence factors.


INTRODUCTION
The revolution in genome sequencing has revealed large numbers of protein sequences with entirely unknown biological roles as well as domains with a well-recognized fold or conserved amino acid sequence but no known function. One example is the PglZ family, which belongs to the alkaline phosphatase superfamily (APS) of metalloenzymes. APS proteins hydrolyse P-O, S-O and P-C bonds, and possess a similar metal coordination architecture at the active site despite their limited sequence identity (1). The APS includes well-characterized enzymes such as alkaline phosphatases, phosphodiesterases, phosphotransferases, phosphomutases, sulfamidases and sulfatases (2). However, no activity or biological function has been assigned to members of the PglZ family, even though the Pfam database contains 1744 sequences with this domain derived from 1424 prokaryote species (3). PglZ family proteins are encoded by various polycistronic operons with essential roles in the protection of bacteria against phage infection, including the well-known phage growth limitation (Pgl) and bacteriophage exclusion (BREX) systems (4)(5)(6)(7)(8)(9). The PglZ domain is also found in many response regulator (RR) proteins of two-component systems (TCSs), which are essential signal transduction systems that allow bacteria to respond to environmental cues. However, a TCS featuring a PglZ family protein has yet to be characterized in detail.
Multiple TCSs responding to different stimuli are present in all prokaryotes, each featuring a signalling histidine kinase (HK) and corresponding RR protein. A signal detected by the HK sensory input domain leads to the autophosphorylation of a conserved His residue in the cytoplasmic domain, followed by the transfer of the phosphate group to a strictly conserved Asp residue in the receiver domain (RD) of the cognate RR (10). The phosphorylated RR usually activates an effector domain, which in >60% of TCSs is as a direct transcriptional regulator of target genes (10). Effector domains not only control transcription, but also perform many other well-characterized biological functions (11). However, one of the most remarkable exceptions is the PglZ effector domain, which has not been assigned a function. A PglZ effector domain is found in 75 species, including several oral commensals such as Prevotella spp. and periodontal pathogens such as Tannerella forsythia and Porphyromonas gingivalis (http://www.p2cs. org/) (12).
In P. gingivalis, the PorX protein is an orphan RR possessing a canonical RD and a PglZ effector domain. It is thought to regulate the secretion of virulence factors via the type 9 secretion system (T9SS), also known as PorSS or Pe-rioGate (13,14). Most proteinaceous virulence factors are secreted via the T9SS (including the gingipains, which are essential for pathogenicity) and deletion of the porX gene renders the mutant avirulent (15). PorX is thought to interact with PorY, a classical (but orphan) signalling HK in the inner membrane (13,16). Three distinct (but not mutually exclusive) signal transduction pathways affecting the function of the T9SS have been proposed for PorX. First, an unknown stimulus is assumed to trigger the autophosphorylation of PorY, which transfers the phosphate to the conserved Asp residue in the RD of PorX, and the latter induces genes encoding components of the T9SS by binding to the extracytoplasmic function (ECF) sigma factor, SigP (13,17). Second, activated PorX has been shown to bind directly to the sigP and porT promoters, facilitating SigP-dependent transcription (15,18). However, this mechanism is controversial because PorX lacks a canonical DNAbinding motif (16). Notably, PorT is one of the essential components of the T9SS (19). Finally, activated PorX can bind to PorL, the inner membrane component of the T9SS, which enhances protein secretion (16). In all these pathways, the mechanistic involvement of the PglZ domain of PorX remains undetermined, despite the well conserved architecture of the catalytic site typical for APS proteins.
To shed light on the role of the PglZ domain in bacterial TCS pathways and in defence systems against phages, we solved the crystal structure of P. gingivalis PorX, determined the enzymatic activity of the PglZ domain and deciphered its molecular mechanism of action. These data not only reveal the biological activity of the PglZ family, but also define the novel regulatory functions of a RR protein, independent of DNA binding, involving the hydrolysis of key signalling molecules such as cyclic and linear oligoadenylates.

Bacterial strains and general growth conditions
Porphyromonas gingivalis strain W83 was grown in enriched tryptic soy broth (eTSB) comprising 30 g/l trypticase soy broth, 5 g/l yeast extract, 5 mg/l hemin, 0.5 g/l L-cysteine, 2 mg/l menadione (pH 7.5) or on eTSB blood agar (eTSB medium containing 1.5% (w/v) agar and 4% (w/v) defibrinated sheep blood) at 37 • C in an anaerobic chamber (Don Whitley Scientific) with an atmosphere of 90% nitrogen, 5% carbon dioxide and 5% hydrogen. Escherichia coli cloning strains were grown in lysogeny broth (LB) and on 1.5% agar LB plates, supplemented with 100 g/ml ampicillin. P. gingivalis mutants were grown in the presence of 5 g/ml erythromycin and/or 1 g/ml tetracycline.

Expression and purification of wild-type PorX and mutants
The porX gene (KEGG locus tag: PG 0928) was amplified by PCR from genomic DNA extracted from P. gingivalis W83 using primers PorX pGEX BamHI F and PorX pGEX SalI R (Supplementary Table S1). The purified DNA fragment was transferred to the BamHI/SalI sites of pGEX-6P-1 and introduced into E. coli strain BL21 (DE3). The transformants were cultured until the OD 600 reached ∼0.6 and the expression construct was induced with 0.2 mM isopropyl-1-thio-␤-D-galactopyranoside (IPTG). After incubation for 16 h at 20 • C, the cells were collected by centrifugation (5000 × g, 15 min, 20 • C), resuspended in PBS and lysed by passing once through a cell disruptor (Constant Biosystems) at 0.75 kW and 23 000 psi, followed by ultracentrifugation (200 000 × g, 45 min, 4 • C). The supernatant was loaded onto a Glutathione Sepharose 4 Fast Flow column (Cytiva) equilibrated with PBS, washed with PBS, and the tag-free PorX protein was released by cleaving the GST-tag with PreScission Protease for 24 h at 4 • C. The eluted protein was dialysed against 20 mM Bis-Tris (pH 6.5), loaded onto a Resource Q column (Cytiva) and eluted with a linear gradient of NaCl up to 1 M in 20 column volumes (CV). A final SEC step was performed in 5 mM Tris, 50 mM NaCl (pH 7.5) using a HiLoad 16/60 Superdex 200 column (Cytiva). A selenomethionine (SeMet) variant of PorX was similarly obtained, except that minimal medium containing Se-Met (Chem-Impex) was used instead of LB.

BeF 3 -induced dimerization of PorX
PorX was incubated in a reaction mixture containing 50 mM Tris, 150 mM NaCl, 5 mM BeSO 4 , 30 mM NaF and 15 mM MgCl 2 (pH 8.0) for 1 h at room temperature to obtain the dimer. PorX incubated without with BeF 3 is hereinafter described as the monomer. Dimerization was confirmed by SEC using a Superdex 200 Increase 10/300 GL column (Cytiva) equilibrated with 10 mM Tris, 100 mM NaCl (pH 8.0) at a flow rate of 1 ml/min. Phosphorylated PorX (P-PorX) was prepared by incubating PorX with buffer A supplemented with 20 mM AcP and 10 mM MgCl 2 for 1 h at 37 • C. The AcP and MgCl 2 were then removed using Zeba spin desalting columns equilibrated with buffer A. The phosphodiesterase reaction was initiated by adding 50 l 2 M P-PorX (diluted in a test buffer, see below) to 50 l 10 mM bis(4-nitrophenyl)phosphate (bis-pNPP, Merck) and monitoring the release of the reaction product (p-nitrophenol) for 90 min at 37 • C by measuring the absorbance at 405 nm on a SpectraMAX microplate reader (Molecular Devices). The assay was performed in triplicate. For activity pH screening, P-PorX was incubated in 150 mM NaCl, 3 M ZnCl 2 , 100 mM Tris, 50 mM MES, 50 mM acetic acid (pH 5.0-9.5) or CAPS buffer (pH 9.5-10.5) for 30 min at 37 • C. For metal screening, P-PorX was incubated in buffer A containing 0.5 mM ZnCl 2 , CuCl 2 , MnCl 2 , MgCl 2 or CaCl 2 . PorX was used as a control of the reaction.

UHPLC-MS analysis
Samples were analysed by ultrahigh-performance liquid chromatography/mass spectrometry (UHPLC-MS) on an Acquity I-class UPLC system and Synapt G2-Si Q-Tof mass spectrometer with an electrospray ion source (Waters) follow. Samples were loaded onto an ACQUITY UPLC 1.7 m (2.1 × 100 mm) BEH Amide column (Waters) maintained at 45 • C, followed by isocratic elution in a mobile phase of 10 mM NH 4 HCO 3 (pH 9.0) in water and acetonitrile (1:2 v/v). The total analysis time was 6 min at a flow rate of 0.16 ml/min. The auto-sampler temperature was set to 8 • C, and 2 l of each sample was injected. A full MS scan in electrospray negative ionization mode (ESI -) was applied (capillary voltage = 2.5 kV, sampling cone = 40, source offset = 30, source temperature = 120 • C, desolvation temperature = 450 • C, desolvation gas flow = 800 l/h, cone gas flow = 40 l/h). Data were acquired in resolution mode using MassLynx v4.1. Leucine enkephalin (50 pg/l) was used as a lock spray interface at a flow rate of 10 l/min (m/z = 554.2615).

Site-directed mutagenesis of the porX gene
Mutations were introduced into the PorX-pGEX-6P-1 vector using primers with overhangs containing the required modifications (Supplementry Tables S1 and S2). The DNA was ligated using the NEBuilder HiFi DNA Assembly Cloning Kit (21). The resulting plasmids were analysed by DNA sequencing.
Nucleic Acids Research, 2022, Vol. 50, No. 21 12561 Crystallization, X-ray data collection and crystal structure determination Crystallization screens were carried out at 20 • C in a sitting drop setup, at the Barcelona Science Park Crystallization Platform. We mixed 300-400 nl of protein solution (PorX-SeMet in all cases) containing 5 mM Tris pH 7.5, 150 mM NaCl (PorX-BeF 3 crystal) or 50 mM NaCl (PorX-HR and T272A-pGpG crystals) with the same volume of precipitant solution, using nano-dispensing robots and 96-well plates (MRC96T-PS). BeF 3 reagents (at final concentrations of 15 mM MgCl 2 , 30 mM NaF, 5 mM BeSO 4 ) were added directly either to the protein solution (PorX-BeF 3 crystal) or to the precipitant solution (PorX-HR and T272A-pGpG crystals). Crystals were cryo-protected by adding 27.3% glycerol to the reservoir solution and dispensing a volume to the crystallization drop so that the final concentration was an estimated 18%. Crystals were harvested using cryo-loops (Molecular Dimensions) and were flash frozen in liquid nitrogen. PorX-BeF 3 (at 2.3 mg/ml) crystallized in 100 mM glycine pH 8.0, 100 mM sodium formate and a polyethylene glycol (PEG) mixture (1.5% each of 20K, 10K, 8K and 6K). PorX-HR crystals appeared in 100 mM glycine pH 8.0-8.5, 100 mM sodium formate, 3-4% PEG 8K at a protein concentration of 12.8 mg/ml with 0.1 mM ZnCl 2 added to the precipitant and 0.5 mM GMP to the cryoprotectant solution. Crystals grew as a cluster which was broken with CrystalProbes (Hampton Research) into single crystals after adding cryo-protectant. A second crystal from the PorX-HR crystal cluster was soaked for 20 min in solution containing 0.5 mM MnCl 2 . T272A-pGpG (12 mg/ml) crystallized like crystal PorX-HR but with 0.5 mM of ZnCl 2 and pGpG. X-ray data were collected at the ALBA synchrotron (Cerdanyola del Vallès, Spain; T272A-pGpG) and at the European Synchrotron Radiation Facility, beamline ID29 (ESRF, Grenoble, France; PorX-BeF 3 and PorX-HR) at 100 K with Pilatus 6M and Pilatus 6M-F detectors, respectively. PorX-BeF 3 and T272A-pGpG crystals were collected at the Se-absorption edge (0.97893 and 0.97923Å, respectively), whereas PorX-HR data were collected at the Zn 2+ absorption edge (1.28202Å). PorX-BeF 3 was collected at a single position, PorX-HR at five positions with three kappa orientations, and T272A-pGpG at two positions with different kappa orientations. The Mn-soaked fragment from the PorX-HR crystal cluster was diffracted at the Mn (1.89608Å) and Zn absorption edges to confirm metal identity.
Data were processed with XDS (22). Correction factors were applied, and multiple datasets were merged with XS-CALE (22). PorX-BeF 3 was further scaled with AIMLESS (23). PorX-HR and T272A-pGpG datasets were scaled, merged and truncated, and the resolution cut-off based on CC1/2 >50 was applied using ap SCALE (Staraniso suite) (24). Structures were solved using the MR-SAD PHASER-EP pipeline (25) as implemented in the CCP4i suite (23). The model used for MR-SAD was built on a previous SAD solution using SHELX C/D/E (26) and several merged datasets. Models were refined and maps calculated with PHENIX.refine (27) and BUSTER.refine (28) by keeping unmerged Friedel pairs, alternating with manual model building and real space refinement using COOT (29). Figures and RMSD values were generated with UCSF Chimera (30) and COOT (29). The overall structures of PorX-HR and PorX-BeF 3 are very similar, yet they provide complementary structural information. The LH subunit of PorX-BeF 3 was fully traced (chain B and D) at loops L␣3b-␣3c (amino acids 298-307) while these could not be traced in PorX-HR molB and molD. The high resolution of the PorX-HR data allowed us to unambiguously identify BeF 3 at the PglZ active site, place metals and coordinating waters with confidence, and the strong anomalous signal from zinc permitted metal identification. Further, the PorX-HR structure shows two clear conformations of loop L␤Ic-␤2 of the capping domain (aa 431-436) in molecules A and C, while in PorX-BeF 3 the same loop could only be tentatively traced due to weak density. The dataset collected at the Mn absorption edge did not give any anomalous signal, so an Mg was placed at the RD active site.

Small angle X-ray scattering (SAXS)
The PorX monomer was concentrated using Amicon Ultra 0.5-ml centrifugal filters to concentration between 15.5 and 17 mg/ml and then diluted with appropriate buffers. Buffer A was used for PorX-monomer samples and BeF 3buffer (buffer A plus 5 mM BeSO 4 , 30 mM NaF and 15 mM MgCl 2 ) for PorX + BeF 3 dimer samples. The final concentrations of the PorX monomer were 0.5, 2.0 and 10 mg/ml. The final concentrations of the PorX + BeF 3 dimer were 0.5, 1.0 and 2.0 mg/ml. Samples PorX + Zn 2+ and PorX + BeF 3 + Zn 2+ were diluted to 0.5 mg/ml with buffer A or BeF 3 -buffer supplemented with 43 M ZnCl 2 (to avoid high zinc concentrations while still saturating all zinc binding sites) and concentrated to 2-3 mg/ml before redilution to 0.5, 1.0 and 2.0 mg/ml. Samples were centrifuged before SAXS data collection, and collected in batch mode at the BM29 BIOSAXS beamline (ESRF Grenoble, France). Data were processed and analysed with BioXTAS RAW (31), which included data reduction, Guinier analysis, molecular weight (MW) estimation and indirect Fourier transform (IFT) analysis with GNOM (32) from the ATSAS 3.0 package (33). The MW was calculated based on the Porod volume (V p ) (34), or the volume of correlation (V c ) (35). For monomeric PorX, datasets at 0.5, 2 and 10 mg/ml were superimposed and corresponding regions with the best signal/noise ratio were selected and merged into a new curve. PorX + Zn 2+ samples at 0.5 and 2 mg/ml showed different behaviour at the Guinier region and were processed separately, whereas the data at 1 mg/ml were discarded due to polydispersity. PorX + BeF 3 data were best at 2 mg/ml, whereas PorX + BeF 3 + Zn 2+ data were best at 1 mg/ml. The datasets were processed and analysed, whereas low-q regions showing intramolecular interactions were discarded. Ab initio electron density was reconstructed with DENSS (36) using the IFT calculated with GNOM. For the electron density maps of PorX and BeF 3 + PorX, 100 ab initio maps were calculated and averaged in slow mode with default parameters and refined against the P(r) function with an oversampling ratio of 5 to optimize map quality. The PorX monomer was modelled manually with CHIMERA v1. 16 (30) against the DENSS electron density (37), and the model was then refined and validated with SREFLEX (38) and CRYSOL (39) with 1000 points of the computed SAXS curve, 33 maximum order of harmonics (as suggested by CRYSOL) and constant background subtraction.

Multi-angle laser light scattering (MALLS)
MALLS data were collected at the Barcelona Science Park Protein Purification Facility. A DAWN-HELEOS-II detector (Wyatt Technology) equipped with a 664.3-nm laser was coupled to a Superdex 200 Increase 10/300 GL column. We separated 50 l of each sample at room temperature. The system was first equilibrated with the specific buffer and calibrated with bovine serum albumin (BSA). The samples included SeMet-PorX, protein variants D58A and D361A/H365A, and the corresponding phosphorylated forms. The running buffer contained buffer A plus 10 mM MgCl 2 (or in some cases 20 mM AcP + 10 mM MgCl 2 and/or 43 mM ZnCl 2 ). Proteins were concentrated to between 4.6 and 6.9 mg/ml, as measured on a GE NanoVUE spectrophotometer (at 280 nm) and diluted with concentrator filtrate to 2.7 mg/ml for the Zn 2+ -containing samples, or 2.6 mg/ml for the other samples. Salts were then added from stock solutions diluting protein concentrations to 2.0 mg/ml. Phosphorylated samples were pre-incubated with 20 mM AcP for 1 h at 37 • C before the corresponding run, whereas the Zn 2+ -containing samples were preincubated with 0.7 mM Zn 2+ . During incubation with Zn 2+ , partial protein precipitation was observed for the wildtype and D58A variant. Before injection, all samples were passed through 0.22-m cut-off filters and kept at 4 • C in the MALLS device autosampler. For each buffer, injections were done every 1.5 CV at a flow rate of 0.5 ml/min. Phosphorylated samples were injected first to minimize dephosphorylation.

PglZ comparison with BREX ortholog proteins
Protein sequences taken from the NCBI database included PorX (GenBank: BAG33538.1), BREX PglZ 1 B (NF033450), BREX PglZ 2 (NF033446), BREX PglZ 3 (NF033449), BREX PglZ 4 (NF033445), BREX PglZ 5 (NF033444) and BREX PglZ 6 (NF033443). For BREX-PglZs, we used the consensus sequences of all members of the corresponding protein family within the database. The 3D models were predicted using ColabFold, which is a combination of AlphaFold2 (40) and MMseqs2 (41). First, proteins were predicted as monomers, whose structures were then subdivided into smaller subdomains such as globular N-terminus, helical bundle, PglZ and for BREX-2, -4 and -6, an additional C-terminal domain. Predictions were limited by the number of 1000 residues. Therefore, in some cases, the prediction was calculated for fragments that overlapped at their ends, so that the structures of the fragments could be connected for further analysis. Sequences were aligned using the secondary structure predictions from the AlphaFold models, as well as predictions from JPRED 4.0 (41) in combination with subsequence alignments by Tcoffee, MAFFT, MUSCLE and CLUSTAL (42) followed by manual comparisons and readjustments in JALVIEW (43). JALVIEW was also used to calculate and export phylogenetic trees by average distance using Blosum62, and the trees were visualized using Dendroscope (44). The identity matrix was calculated using Clustal 2.1 (45) in test mode (without realignment). Structures/models were compared, aligned and visualized with UCSF CHIMERA (30).

PorX surface conservation
Representation of PorX surface conservation was done with USCF CHIMERA (30). Its 5 closest homologs within the PDB were structurally superposed at the APS catalytic core, sequences aligned and colored in a gradient by conservation. Zinc atom radii were enlarged to 2.5Å to visualize buried Zn2 at the surface. Coloring thresholds used for identical residues were 100%, similar residues 75%, and not conserved 50% or below.

PorX dimerizes in the presence of acetyl phosphate
In a canonical TCS, the activation of a RR protein leads to phosphorylation within the RD, which often results in the formation of RR dimers, but this has not yet been demonstrated for RR proteins containing a PglZ domain (10). RD phosphorylation in vitro can be achieved by incubating the protein with low-molecular-weight phospho-donors such as acetyl phosphate (AcP) in the presence of Mg 2+ (46). Using this approach, PorX was indeed phosphorylated with radiolabelled AcP ( Figure 1A). Size-exclusion chromatography (SEC) confirmed that phosphorylation increased the Stokes radius, which usually indicates a conformational change or the formation of multimers ( Figure 1B). We determined the absolute molecular weight (MW) of the protein by multiangle laser light scattering (MALLS) ( Figure 1C, D; Supplementary Table S3). The peak of non-phosphorylated PorX (theoretical MW = 61.1 kDa) corresponded to a monomer (MW = 70.2 kDa) whereas the phosphorylated sample corresponded to a dimer (MW = 118.2 kDa). Mg 2+ alone did not induce multimerization ( Figure 1C) (10). Due to lability of the acyl phosphate linkage, we incubated PorX with BeF 3 to mimic phosphorylation since, in the crystallization drop, this compound makes a reversible ionic interaction with the RR active site in a stable equilibrium, enabling crystallization (47). Subsequent attempts to crystallize the PorX monomer did not yield any crystals after extensive trials, whereas PorX-BeF 3 generated crystals that diffracted to a resolution of 1.9Å (Supplementary Table S4).

The PorX-BeF 3 crystal structure is an intertwined dimer
The PorX-BeF 3 crystal structure shows two dimers (AB and CD chains, respectively; A and C are structurally equivalent, and so are B and D) in the crystallographic asymmetric unit. Relevant conformational differences were found between monomers in a dimer (RMSD between molecules A/B = 1.32Å, and between molecules C/D = 1.30Å; differences between monomers are described below) but not between dimers (RMSD between molecules A/C = 0.469 A, and between molecules B/D = 0.381Å). We therefore chose dimer AB and molecule A (molA) as the representative structures (Figure 2A). PorX bears an N-terminal RD (Pro7-Ile128), followed by a three-helix bundle domain (HBD) comprising helices H1, H2 and H3 (Ile129-Arg211), and a C-terminal PglZ domain (Pro212-Lys518). Dimerization involves two interfaces, between the RDs and between the PglZ domains, and a binding energy of -39.9 kcal/mol according to QtPISA (23). The monomers intertwine, the RD of one subunit sitting on the HBD of the other ( Figure  2A, C).

The RDs of the PorX dimer adopt an activated conformation
Within the PorX dimer, both RDs display the canonical (␣/␤) 5 doubly-wound fold, which consists of a central parallel five-stranded ␤-sheet (strands ␤1-␤5) surrounded by ␣-helices (helices ␣1-␣5). Helices ␣1 and ␣5 are found on one face of the ␤-sheet, whereas helices ␣2, ␣3 and ␣4 are found on the other. Each strand is connected to an ␣helix by a loop (10) (Figure 2; Supplementary Figure S1). At the C-terminal end of the ␤-sheet, the connecting loops form a cavity that contains the highly conserved active site, which includes Asp14 and Asp15 (end of strand ␤1), Asp58 (␤3 tip), Thr86 (␤4 tip) and Lys108 (␤5 tip) (Figure 2A, B; Supplementary Figure S1). These residues coordinate BeF 3 and Mg 2+ via interactions typically found in all activated RDs (47,48). The highly conserved Asp58 is found at the conserved phosphorylation position in RDs, and its terminal carboxyl oxygen O ␦1 contacts the beryllium of BeF 3 . In addition, Asp58 O ␦2 together with one BeF 3 fluorine atom, Asp15 O ␦1 and a water molecule, form the equatorial coordination plane of Mg 2+ , which occupies the canonical po-sition of this metal in RDs (see methods), and is necessary for phosphorylation (10) ( Figure 2B, Supplementary Figure  S1). The Mg 2+ octahedral coordination sphere is fulfilled with Asn60 C=O (O60 in Figure 2B) and a second water molecule, which is contacted by Asp14 O ␦1 . BeF 3 is also stabilized by Thr86 O ␥ 1 and N Lys108, and main-chain amides from Glu59, Asn60 and Lys87 (atoms N59, N60 and N87, respectively; Figure 2B).
Thr86 not only interacts with BeF 3 but also forms the highly conserved TY pair (10,49) with Tyr105 in the centre of ␤5, at the RD dimer interface ( Figure 2D). Crystal structures of activated RR dimers have extensively shown that, following phosphorylation, the side chain of the conserved Thr residue in the TY pair re-orients towards the active site phosphate, thereby creating a space that is occupied by the conserved Tyr residue (which is otherwise oriented to the solvent) (10). Because the Tyr residue is in the centre of ␤5, its reorientation causes the whole ␣4-␤5-␣5 surface to be remodelled. In the OmpR subfamily of RR proteins, this remodelling enables the formation of RD dimers (10,49). The PorX structure shows that RDs of molA and molB dimerize precisely via this surface (Figure 2A, C, D), with Tyr105 oriented towards Thr86, the latter providing the O ␥ 1 atom that contacts BeF 3 ( Figure 2B). This is consistent with the active conformation of the RDs.

The intermediate HBD joins the RD and PglZ domain
Beyond RD helix ␣5, the polypeptide chain bends by 80 • (Figure 2A, C) and folds as helix H1 (Gln125-Met148) of The residues that coordinate BeF 3 (beryllium and fluorine atoms in pistachio and magenta, respectively) and Mg 2+ (light green sphere) are shown as sticks and are labelled. Oxygen and nitrogen atoms are represented in red and blue, respectively. Polar contacts are represented as dashed lines. N59, N60, O60 and N87 correspond to the main-chain atoms of the corresponding residues. Water molecules are represented as light blue spheres. (C) Interface between the two RDs. In one monomer, the RD is represented in green and the HBD in orange, the other monomer is depicted in grey. The RD is followed by the HBD, whose ␣-helices are indicated (H1, H2 and H3). The arrow shows the kink in H1. (D) RD dimer from the top. The interface helices ␣4 and ␣5 and ␤-strand 5 (␤5) are indicated. Tyr105 and Thr86 (in blue and green in molA and molB, respectively), BeF 3 and Mg 2+ are depicted. (E) Sharp bend between RD helix ␣5 and H1 from the HBD. The hydrophobic core that stabilizes the inter-subunit interaction involves the burial of molB Ile98 (helix ␣4) into the hydrophobic cavity built by molA Leu117, Lys120 and Lys121 methylene side chains, Gln124 from helix ␣5, and Gln125, Ile128 and Ile129 from helix H1. (D) and (E) use the same colour code as in (C). the HBD. This bend creates a hydrophobic concavity occupied by molB helix ␣4 ( Figure 2E). H1 also contacts molB RD helix ␣3 through an extensive network of water molecules. It kinks (black arrow in Figure 2C) to pack against the central hydrophobic core of the HBD, which also contains the buried H2 (Phe154-Gln173). H3 (Arg176-Ala206) contacts laterally the hydrophobic core while making extensive contacts with the PglZ domain ( Figure 2A). Accordingly, whereas H1 contacts the other subunit's RD, the remaining HBD forms a block that sits on the PglZ domain of its own monomer.

The two PglZ active sites within the dimer adopt different conformations
The PglZ domain displays a canonical AP-core (␣/␤) 6 fold as found in the APS (50), which consists of a ␤-sheet flanked by ␣-helices ( Figure 3A). In PorX, a ␤-rich capping subdomain (51), hereafter termed cap subdomain, is inserted between ␤5 and ␣6, so the overall topology of the PglZ domain is ␣1-␤1-␣2-␤2-␣3(a-c)-␤3-␣4-␤4-␣5(a-d)-␤5cap-␣6-␤6 ( Figure 3A; Supplementary Figure S1). The two PglZ domains from molA and molB are highly similar, except precisely at the conserved phosphodiesterase active site core residues ( Figure 3A inset), in proximity to the dimerization interface, where there are striking structural differences. Specifically, the Phe359-Arg367 residues of molA fold as helix ␣5a ( Figure 3B, in magenta) whereas the same residues in molB fold in a well-defined extended loop (Figure 3C, in yellow). The region Arg367-Ser370, which also belongs to helix ␣5a in molA, forms a new N-terminus in the downstream helix ␣5b in molB, which is described as helix ␣5b . Therefore, molA features two consecutive helices (␣5a-␣5b, conformation HH), whereas molB features a loop and a helix (loop + ␣5b , conformation LH). Furthermore, in helix ␣5a of molA, Asp361 O ␦1 and His365 N ε2 coordinate a Zn 2+ ion (Zn1) ( Figure 3B, Suppl. Figure S1), which is absent in molB due to the L loop conformation that relocates both side chains and places Met363 near to Zn1 instead (compare Figure 3B and C). In molA, Zn1 interacts with BeF 3 , which in turn coordinates a second Zn 2+ (Zn2). Notably, BeF 3 and Zn2 are also present in molB despite the absence of Zn1 ( Figure 3B, C, Supplementary Figure S1). Both Zn1 and Zn2 were confirmed by anomalous diffraction at the Zn absorption edge (see methods and Suppl. Fig S1). The protein atoms that coordinate BeF 3 in molA include Thr272 O ␥ 1 and the corresponding backbone amide (atom N272 in loop ␤3-␣3 a , also in molB; Figure 3B, C). In both molA and molB, Zn2 is also coordinated by Thr272 O ␥ 1 , as well as Asp415 O ␦2 and His416 N ε2 (in the loop connecting to the cap subdomain at one side of the active site), and Asp239 O ␦1 (at the end of PglZ ␤1) (Figure 3B, C). Thr272 occupies the conserved catalytic Thr/Ser position of phosphodiesterases in the APS (1), whereas the two Zn 2+ ions and BeF 3 in molA occupy the same positions as the metals and phosphate found in many APS structures (1,50,52) (Supplementary Figure S2). Notably, Zn 2+ ions were not added during any production nor crystallization step of PorX-BeF 3 , indicating that the metal was spontaneously acquired from E. coli. In the unprecedented PglZ dimer, the second Zn therefore reconstitutes the canonical bi-metal site of APS phosphodiesterases in molA, but in molB the site is disrupted due to the LH conformation.
The contacts between PglZ domains not only involve helices ␣5a and ␣5b, but also the following helix ␣5c, which makes an antiparallel interaction with ␣5c from the other monomer. Remarkably, none of the other available APS structures dimerise by this surface, and despite they possess helix ␣5a (in which Asp361 and His365 are fully conserved for phosphoesterases; the active site amino acids are highly conserved among APS, see Figure 3A, Supplementary Fig S2) they do not show helices ␣5b and ␣5c (Supplementary Fig S2). In these other structures, a short turn connects ␣5a with a long helix that eventually coincides with the C-terminus of PorX ␣5d, suggesting a more rigid region. Thus, among APS members, ␣5b and ␣5c are PorX-specific insertions that configure the dimer interface.

The PorX RD and PglZ domains show extensive crosstalk
To gain more insight into the functional relationship between dimerization, phosphorylation, the binding of Mg 2+ in the RD, and the binding of Zn 2+ in the PglZ domain, we prepared a series of structure-guided protein variants mutated at the active site or dimerization interfaces, and analysed their ability to be phosphorylated by 32 P-AcP and [␥ -32 P]-ATP ( Figure 4A), as well as their behaviour as un-and phosphorylated proteins in SEC (Figure 4B-G; please see Figure 1B for SEC of the wt protein for comparison). Notably, RDs are generally phosphorylated by 32 P-AcP (46), and the PglZ domain from APS phosphoesterases can be transiently phosphorylated at the conserved Thr/Ser residue (Thr272 in PorX) during the catalytic turnover of [␥ -32 P]-ATP (51,53,54). Consistent with this, we found that wild-type PorX bound efficiently to the radioactive moiety from both 32 P-AcP and [␥ -32 P]-ATP ( Figure 4A). Mutation of the RD phosphorylation site Asp58 (D58A) abolished phosphorylation by 32 P-AcP and prevented dimerization ( Figure 4A, B). Surprisingly, it also prevented phosphorylation by [␥ -32 P]-ATP ( Figure 4A), suggesting that the RD regulates the PglZ domain. Accordingly, the mutation of residues involved in Zn-binding or hydrolysis (D361A/H365A and T272A, respectively) caused the anticipated complete loss of phosphorylation by [␥ -32 P]-ATP but also attenuated phosphorylation by 32 P-AcP as well as dimerization, suggesting the PglZ domain can attenuate the effect of the RD ( Figure 4A, C, D). Disruption of the interface between RDs (M94K/D104A/I129A) or PglZ domains (S385E/S389E, S385E/S389E/D361A/H365A) completely abolished dimerization but, interestingly, also blocked phosphorylation by 32 P-AcP or [␥ -32 P]-ATP (Figure 4A, E-G). These results suggest that the activating signal at the catalytic site (of either the RD or PglZ domain) was transmitted to the corresponding dimerization surface. The converse was also true: intact interfaces were required for the active site to function. The two domains were therefore interdependent, with mutations causing functional impairment in one domain also disrupting the activity of the other.
To analyse the effect of Zn 2+ in more detail, a threefold molar excess of Zn 2+ was added to the SEC buffer, which surprisingly induced the dimerization of wild-type PorX, RD mutants (D58A, M94K/D104A/I129A) and PglZ mutants (T272A), but neither D361A/H365A nor S385E/S389E were able to dimerize (Supplementary Figure  S3). These results suggest that, by binding to Asp361 and His365, Zn1 facilitates PglZ dimerization by inducing the HH fold to create the appropriate interface contacts abolished by bulky residues in the S385E/S389E mutants. These effects were not observed in the presence of Mg 2+ alone (Supplementary Figure S4). To confirm the nature of the SEC elution peaks, we used MALLS to compare the absolute MW of wild-type PorX and that of the D58A and D361A/H365A mutants (affecting the metal-binding sites) in the presence of Mg 2+ and Zn 2+ , with or without phosphorylation. The results confirmed that the peak shifts induced by adding Zn 2+ were due to dimerization (Supplementary Figure S5; Supplementary Table S3).

The PorX PglZ domain shows phosphodiesterase but not monophosphatase activity
RR proteins can be assigned to functional families by looking for similar architectures (55). We therefore screened for structural homology using DALI (56), and found that the top 12 closest structural homologs (Zscores > 14.9) were phosphodiester hydrolases, phosphonoacetate hydrolases or alkaline phosphatases, and contained two zinc metal centres each (Supplementary Table S5). The highest-ranked hits (PhnA, XAC and Ecto-Nucleotide Pyrophosphatases/Phosphodiesterases, ENPPs 1 to 7) showed the closest structural similarity near the Zn 2+ -binding sites whereas the surrounding cavities were more diverse. Indeed, ENPPs hydrolyse mononucleotides, dinucleotides and (cyclic) polynucleotides (including Ap3A and Ap4A) and feature large active site clefts with several pockets similar to PorX, suggesting that PorX may accept dinucleotides or longer (cyclic) polynucleotides (Supplementary Figure S6). We therefore tested whether nonactivated and AcP-activated PorX (P-PorX) were able to hydrolyse the phosphodiesterase substrate analogue bis(pnitrophenyl) phosphate (bis-pNPP) in the presence of different divalent cations (Zn 2+ , Cu 2+ , Mn 2+ , Mg 2+ and Ca 2+ ) or without. Only Zn 2+ stimulated the activity of PorX (Figure 5A, B), and this was dependent on the pH ( Figure  5C, D). The optimal pH differed between PorX (pH 8-8.5) Figure 4. Phosphorylation of PorX mutants analysed by size-exclusion chromatography (SEC). The profiles represent PorX structure-based mutants at the RD or PglZ domain active sites or at domain interfaces following incubation with AcP (orange curves) and corresponding non-phosphorylated controls (blue curves). The SEC corresponding to the wild-type protein is shown in Figure 1B  and P-PorX (pH 8.5-9), confirming that RD phosphorylation and dimerization alters the activity of PglZ bound to Zn 2+ ( Figure 5C, D). In contrast, there was no activity against the monophosphatase substrate p-nitrophenyl phosphate (pNPP) under any conditions (data not shown).
We next tested the cleavage kinetics of bis-pNPP for wild-type PorX and mutants affecting the active site and domain interfaces, in the presence of Zn 2+ . Apparently, the Zn-induced dimerization can induce enzymatic activity even in the absence of AcP. Therefore, the observed effect of AcP was only moderate and only slightly increased the reaction turnover rate (k cat ) of wild-type PorX ( Figure 5E, F; Supplementary Table S6). In contrast, AcP did not stimulate the activity of the D58A or M94K/D104A/I129A RD interface mutants ( Figure 5E Table S6). AcP only slightly stimulated the PglZ interface mutants S385E/S389E, and no activity was detected for the other mutants. We concluded that PorX is a Zn 2+ -dependent phosphodiesterase by virtue of the PglZ domain.

The crystal structure of PorX in a complex with pGpG
We initially considered the cyclic dinucleotide c-di-GMP as a potential natural substrate for PorX because this is a major signalling molecule that controls motility, biofilm formation and virulence (57). P. gingivalis produces diguanylate cyclase PGN 1932 (58) that catalyses the synthesis of c-di-GMP, but it does not possess EAL or HD-GYP homologues, which degrade this cyclic dinucleotide (59,60). Both c-di-GMP and its derivative pGpG would fit in the wide active site of PorX, similar to mouse ENPP1 and rat ENPP3, which were crystallized with pApG and Ap4A, respectively (61,62) (Supplementary Figure S6). Therefore, we attempted to crystallize PorX and inactive variant T272A as complexes with c-di-GMP, pGpG and GMP, to catch any of the three steps of c-di-GMP cleavage, as well as GTP, GTP␥ S analogue and ATP, by co-crystallization or  Table S4). pGpG binds all four molecules (A and B chains in one dimer, C and D in the other, all of which with HH conformation) and the pGpG 5 phosphate is coordinated by Zn1 and Zn2, similar to BeF 3 in the unliganded structure ( Figure 6A; Supplementary Figure S6A, S7). The presence of two Zn 2+ ions in all molecules may reflect the high Zn 2+ concentration (0.5 mM). However, PorX under the same conditions (0.5 mM Zn 2+ ) but in the presence of c-di-GMP instead of pGpG crystallized as dimers with LH/HH conformation, without a bound nucleotide. Therefore, pGpG stabilized dimers with di-Zn and HH/HH conformation.
From the Zn binding site, pGpG emerges occupying part of the wide catalytic centre, which features several additional pockets (Supplementary Figure S6A). The 5' guanine (Gua1) is located between pockets A and B, aligned with a loop between helices ␣3b and ␣3c (L␣3b-␣3c, residues 297-307; Figure 6A). Notably, two guanine-specific atoms, Gua1 N2 and O6, are contacted by the protein. Gua1 N2 is contacted by main chain C=O Glu302 in molecules B, C and D (O302 in brackets in molA; Figure 6A), whereas Lys305, also at L␣3b-␣3c, stacks its extended methylene chain against the guanine rings. In addition, Lys305 N in molC contacts Gua1 O6, a contact that could easily be established also in molecules A, B and D with a slight reorientation of the side chain tip. In all molecules, the amide from Asn306 backbone (atom N306 in Figure 6A), at the end of loop L␣3b-␣3c, contacts O2 from the Gua1 sugar, further stabilizing the ligand in the active site. Note that in molB and molD from the ligand-free PorX structures with the LH conformation, loop L␣3b-␣3c is highly flexible and difficult to trace (in molA and molC, the loop is stabilized by contacts with the second dimer) whereas in the T272A-pGpG structure, molB and molD show this loop stabilized by adding further contacts to the ligand. Gua2 extends along pocket A towards the protein surface (Figure 6A), but the electron density is well defined only at its phosphate ( Figure 6B). Next to it, Asn358 and Arg276 contact each other, close to where a third metal ion (Mg 2+ ) is found in the active sites of alkaline phosphatases ( Figure  6A). This contact creates a platform, which is occupied by an electronegative phosphate oxygen from Gua2. Beyond the phosphate, the density around Gua2 purine rings becomes fainter close to the solvent. Gua2 is sandwiched between Leu307 and poorly-defined Tyr332, making stacking interactions with the latter that rotates the Tyr side chain by 60 • compared to PorX-BeF 3 structure ( Supplementary Figure S7). Apart from the local changes, the main differences between PorX with and without bound pGpG were mostly found in the flexible loop of the cap subdomain (Ser432-Thr435; Suppl. Figure S1, S7) and are probably caused by crystal packing. The pGpG molecule does not fill all pockets of the active site cavity, as also shown in the structures of ENPP1-pApG and ENPP3-Ap4A (61,62) (Supplementary Figure S6A-C), suggesting the site accepts larger substrates or allows the reorientation of the ligand during catalysis.

P-PorX cleaves cyclic oligoadenylates and linear oligonucleotides
The structure of PorX in a complex with pGpG suggested that the physiological substrate could be a signalling nucleotide, so we screened the activity of PorX against a library of nucleotides (AMP, ADP, ATP, GMP, GDP, GTP, cdi-GMP, c-di-AMP, 2 ,3 -cGAMP, 3 ,3 -cGAMP, c-AMP, c-GMP, cA3, cA4, cA6, pApA, pApG, pGpG, AP3A, AP4A and AP4G) by high-performance liquid chromatography (HPLC). PorX showed moderate reactivity in the presence of the cyclic adenylates (cOAs) cA3, cA4 and cA6 (Figure 7A-C). The main product of these reactions was a linear oligoadenylate, resulting from a single cleavage (pA-pApA for cA3, pApApApA for cA4 and pApApApApApA for cA6), together with intermediate hydrolysis products en route to the final product AMP (Figure 7A-C; Supplementary Table S7). We used pApA as a positive control (Figure 7D) and tested guanine-containing linear dinucleotides such as pApG ( Figure 7E) and pGpG ( Figure 7F), which were cleaved to single nucleotides with similar efficiency, indicating that PorX also accepts guanine. PorX showed high selectivity for cyclic oligoadenylates or linear oligonu- cleotides because it did not cleave other nucleotides from the library (Supplementary Figure S8).

The monomer in solution reveals the reorientation of helix H1, the cap subdomain and a highly flexible HH/LH region
We were unable to crystallize the PorX monomer, so we analysed the structure of PorX and PorX-BeF 3 in solution by small-angle X-ray scattering (SAXS) (Figure 8, Suppl. Figure S9). The oligomeric state of PorX (monomer) did not change at protein concentrations up to 10 mg/ml according to the MW based on the Porod volume (V p ). In contrast, PorX-BeF 3 was already a dimer at concentrations of 1 mg/ml (not shown) and 2 mg/ml (Supplementary Table  S8). The dimerization of PorX was therefore concentrationindependent but required activation. The monomers and dimers were both compact, non-flexible molecules (see Kratky plot in Supplementary Figure S9B). In the absence of BeF 3 , the addition of Zn 2+ had different effects depending on the protein concentration, inducing the formation of dimers at 2 mg/ml but not at 0.5 mg/ml. The overall volume of all samples recorded as the radius of gyration (R g , based on P(r) in Supplementary Table S8) was 29.6Å for the monomer and 33.4Å for the dimer. The R g of the PorX-Zn 2+ dimer was 39.0Å and the largest maximum dimension (D max ) was 118Å (Supplementary Figure S9B; Supplementary Table S8). In contrast, the remaining monomers and dimers had similar D max values of 103 and 105Å, respectively. These similar R g and D max values are consistent with monomers of elongated shape that dimerize along the largest dimension, as we see in the X-ray structure. In contrast, Zn 2+ ions stimulated a non-native dimerization of the non-phosphorylated monomers.
We then tested different models to find the one that fitted best the SAXS data obtained from the PorX monomer. We calculated theoretical scattering curves from molA and molB from the crystal structure, but they fitted the experi-mental data poorly, indicating significant structural deviations. In parallel, an ab initio 3D electron density map (not biased by any model) was calculated from the SAXS data (Supplementary Table S9). We tried to fit crystal monomers against such ab initio map but the HBD H1, PglZ cap subdomain and LH/HH segments of the Zn-binding site did not fit inside the density (Supplementary Figure S9). After relocating these regions by applying iterative calculations guided by automated domain border definition, we generated a final model that fitted precisely to both the ab initio map and experimental SAXS curve ( Figure 8A, B). This model features a rotated RD that connects to the HBD via displaced helix H1 ( Figure 8C, see Supplementary Figure S9 for details). The cap subdomain faces away from the PglZ dimerization surface and apparently covers the active site ( Figure 8C). The SAXS map showed no electron density in the HH or LH region, which suggests high flexibility in the monomer (Supplementary Figure S9). Note that the cap subdomain contacts the HH/LH region in the crystal structure and probably stabilizes it, so the displacement of the cap subdomain in the soluble monomer likely affected the HH/LH region. In contrast, the crystallographic dimer model fitted reasonably the PorX-BeF 3 data (Supplementary Figure S9E, F). Therefore, our SAXS results suggest that, during dimerization, RD and H1 are reoriented to contact the second monomer appropriately, whereas the cap subdomain near the PglZ domain is repositioned in close proximity to the HH or LH region at the dimer interface and increases its stability.

DISCUSSION
The PglZ domain is often found in the RR proteins of TCSs (11)--including PorX, which is required for the activation of the T9SS (18). In an apparently unrelated context, proteins containing a PglZ domain are necessary for the BREX anti-phage defence system (7,9). Here, we determined the crystal structure of PorX, its structure-function relationships, and the enzymatic activity of the PglZ domain. Biochemical experiments and mutagenesis revealed that phosphorylation at RD Asp58 regulates the dimerization of PorX via the RD and PglZ domain, as well as PglZ enzymatic activation. The structure of the dimer indicated that Zn 2+ is present at both PglZ active sites, which adopt different conformations in both PorX-BeF 3 and PorX-HR structures, which confirmed the findings. Specifically, two consecutive helices ␣5a and ␣5b at one edge of the active site feature the 'HH conformation' in one monomer, and these helices are remodelled as a loop and a longer helix ␣5b' in the 'LH conformation' of the other monomer. Whereas the LH conformation shows a Zn bound at the active site, only HH features the canonical bi-metal (Zn 2+ ) site typical for APS phosphodiesterases (1, 50,52). This changed to a double-HH conformation following the binding of pGpG, suggesting that one HH conformation is sterically necessary for PorX dimerization, while ligand binding may induce the HH conformation in the other monomer.
In PorX, conserved Asp361 and His365 belong to helix ␣5a and bind the second Zn in the HH conformation. However, helix ␣5a is also present in all available structures of bimetallo-(two Zn ions) APS, which do not dimerize by this surface, therefore the transition from loop to helix ␣5a is required but seems not sufficient for PglZ dimerization in PorX. Notably, following helix ␣5a there are helices ␣5b and ␣5c that also contribute to the dimer interface. Interestingly, our SAXS data shows that precisely helices ␣5a, ␣5b and ␣5c are disordered in the monomer, suggesting rearrangement during dimerization, which might be facilitated by bi-metal Zn binding that stabilizes ␣5a in at least one subunit. Surprisingly, whereas in APS helices ␣5b and ␣5c are exclusive to PorX, a similar helical arrangement is found in the BREX dimers predicted by AlphaFold, which dimerize precisely by this region. This is an intriguing feature that might be related to a dimerization-dependent activity at the nearby active sites.
We found evidence for extensive crosstalk between the PorX domains. Phosphodiesterase activity was abolished by mutating the PglZ catalytic amino acid Thr272 (homologous to Ser/Thr in the active site of APS proteins) but also by disrupting the PglZ dimerization interface, which borders the active sites and probably influences their stability. The same phenomenon was observed for Asp58 and the RD interface. However, the effect propagated beyond the mutated domain. For example, mutation T272A at the PglZ active site impaired PorX dimerization induced by phosphorylation at Asp58. Conversely, mutation D58A in the RD prevented dimerization and phosphorylation at PglZ Thr272. Excess Zn 2+ compensated for the disassembly of the D58A dimer and promoted phosphodiesterase activity. However, due to the limited intercellular concentration of free Zn 2+ (63), this phenomenon probably does not contribute to the biological function of PorX in vivo. A previous crystallization study with PorX from Flavobacterium johnsoniae also showed crystals containing dimers of nonphosphorylated protein and without addition of Zn (64). Despite the corresponding structure not being available, this indicates that under their crystallization conditions dimers also might be formed spontaneously.
Structural homology searches revealed that the PorX catalytic center is similar to that of enzymes that cleave mononucleotides, dinucleotides and (cyclic) polynucleotides such as cyclic and linear oligoadenylates (Suppl. Table S5). The similarity was high for mouse ENPP1 and rat ENPP3, which have been crystalized with the dinucleotides pApG and Ap4A, respectively (61,62). Like the PglZ domain of PorX, ENPP1 and ENPP3 also feature broad active site clefts with connected pockets. The broad active site of PorX could also accommodate cyclic oligoadenylate (cOA) bases and, in all cases facilitate the flipping of adenosines between pockets, so that all phosphodiester bonds reach the catalytic site, as proposed for substrate catalysis in ENPP1 (61). By screening a library of nucleotides, we showed that dimeric PorX cleaves cOAs and linear oligoadenylates to NMPs. Considering these results, we can hypothesize that the T272A-pGpG structure is representative of a reaction state intermediate with incomplete substrate degradation due to the T272A mutation. All polynucleotides were linked 3'-5'. Therefore, the scissile bond was between the O3' from one adenine and the P from the phosphate of the following adenine. PorX did not cleave mononucleotides, cyclic dinucleotides or other nucleotides. The cleavage mechanism of PorX must differ from that of ring nucleases, which do not require metals, and oligoribonucleases, which are Mn 2+ -dependent 3 →5 exonucleases (65,66).
Cyclic oligoadenylates are generated by type III CRISPR-Cas systems and cyclic oligonucleotide-based anti-phage signalling systems (CBASS) (67)(68)(69)(70). In type III CRISPR-Cas, when the effector complex detects foreign RNA, the Cas10 subunit converts ATP into adenylates cA2 to cA6 (68,69), which bind to the CARF domain of RNAse Csm6/Csx1 to induce the non-specific degradation of invading RNAs (68,69). On the other hand, each CBASS employs a cGAS/DncV-like nucleotidyltransferase (CD-NTase), which synthesizes a nucleotide second messenger (including cyclic trinucleotides) in response to viral infection (67,70) Subsequently, the nucleotide messenger activates an associated Cap effector protein which kills the host cell and, as a consequence, blocks phage propagation (71). All sequenced P. gingivalis strains encode at least one CRISPR-Cas system, but only 36% possess a type III CRISPR-Cas that synthesizes cOAs. Moreover, P. gingivalis strains do not encode CD-NTase nor any known effector of CBASS. Therefore, unless there are other, yet unknown cOA synthesis pathways, it seems unlikely these molecules play a critical role in PorX-regulated signalling. Given that the BREX system and recently discovered associated genes including brxU (72) and brxR (73,74) are also absent in P. gingivalis, it is puzzling that bacteriophages have not yet been shown to infect P. gingivalis, which would indicate the presence of a resistance system (75). Because cOAs regulate defence systems, it is tempting to speculate that they activate an unknown anti-phage mechanism involving PorX, and this should be investigated in more detail.
In addition to cOAs, PorX also cleaves linear oligoadenylates ( Figure 7A-C) and ribodinucleotides ( Figure 7D-F) to mononucleotides. Accordingly, PorX may facilitate RNA turnover, the cycle of transcription and RNA degradation used by cells when they adjust to environmental changes, and this would match the role of PorX as a RR protein. In E. coli, mRNA decay is initiated by the endonucleolytic activity of RNase E, followed by 3 →5 exonucleolytic degradation catalysed by other enzymes (76). The exonucleases cannot complete the degradation cycle because they are unable to cleave dinucleotides through to pentanucleotides (nanoRNAs), which are digested by oligoribonucleases such as Orn in E. coli (77), NrnA and NrnB in Bacillus subtilis (78), and NrnC in Bartonella spp. (79) NanoR-NAs are not only the intermediate products of RNA decay, they are also produced during abortive transcriptional initiation (80) and the single-step cleavage of cyclic dinucleotide second messengers (65,66). NanoRNAs play important roles in cellular physiology including the priming of transcriptional initiation (81), changing promoter-specific gene expression (81), and changing auto-aggregation and biofilm phenotypes by inhibiting enzymes that break down c-di-GMP (66). P. gingivalis W83 encodes NrnA homologue PG 1091, but this does not exclude the possibility that the oligoribonuclease activity of PorX is physiologically relevant, because many bacteria produce multiple oligoribonucleases with redundant functions (78). It is tempting to speculate that an additional route of PorX-dependent gene regulation involves the cleavage of nanoRNAs.
In P. gingivalis, PorX has been associated with PorY (13,16), an orphan signalling HK with a periplasmic sensor domain. However, there is little phenotypic overlap between the porX and porY mutants (18,82), making it unlikely that a PorY/PorX TCS regulates the secretion of virulence factors. Even so, it is possible that PorX and PorY operate as a TCS that regulates T9SS activity in response to environmental cues in other Bacteroidetes. Notably, although P. gingivalis produces only five or six HKs (depending on the strain) including the two orphan proteins PorY and GppX (12), we cannot exclude the possibility that another HK is involved. Alternatively, PorX phosphorylation may depend on an as yet unknown Asp-kinase or AcP. Interestingly, several orphan RR proteins are activated by AcP in vivo (83). The concentration of AcP in P. gingivalis is not known, but in some bacteria such as E. coli it can reach at least 3 mM which suffices for the efficient autophosphorylation of RR proteins (84).
To the best of our knowledge, PorX is unique among RR proteins by combining enzymatic and protein-binding capabilities, both of which depend on PorX phosphorylation and dimerization. PorX regulates the expression of T9SS genes by interacting with the ECF sigma factor SigP (13). In E. coli, a similar case occurs between the RR RssB and the stress-inducible sigma factor RpoS, which regulates the expression of ∼500 genes (85). The C-terminal effector domain of RssB features a PPM/PP2C family phosphatase fold, which it is enzymatically inactive, but binds to RpoS and triggers its degradation. Like the C-terminus of RssB, the C-terminus of PorX may be involved in the interaction with SigP. Specifically, parts of the PglZ cap subdomain or with the PorX-PglZ dimerization interface, which is disordered in the monomer. Despite many similarities, the RssB and PorX RR proteins might have opposite effects on their cognate sigma factors, with RssB marking RpoS for proteolysis but PorX stabilizing SigP (13). Moreover, the phosphorylation of RssB strengthens its interaction with RpoS but the impact of PorX phosphorylation on its interaction with SigP is unknown. PorX also interacts with PorL, which forms an inner membrane complex with PorM to provide energy for the T9SS (16,86). This interaction is reminiscent of CheY binding to the FliM and FliN switch complex proteins of the flagellar C-ring, thus reversing the rotational direction of the filament (87). Indeed, the T9SS was shown to promote the rotary movement of the SprB filamentous adhesin on the cell surface of Flavobacterium johnsoniae (88). Moreover, the binding of CheY to FliN is mediated by a hydrophobic patch on the surface of FliN and a similar patch in PorL is necessary for the interaction with PorX (16).
Although the protein-protein interactions of PorX are well documented, its DNA-binding activity is more controversial. This has been investigated in two independent studies with contradictory results (16,18). The original study reported that full-length PorX does not bind the promoter regions of T9SS-related genes, including porT, sov and porP (13,16), whereas more recent experiments identified two PorX-binding sites in the porT promoter region (18). The structural data obtained herein confirm that PorX lacks canonical DNA-binding motifs, thus a potential mechanism of gene expression regulation by interacting directly with promoters remains unclear. So far, this function has been clearly proven for SigP, which binds directly to the promoter region of porT (13).
Our analysis of sequenced bacterial genomes revealed that PorX is limited to the Fibrobacteres-Chlorobi-Bacteroidetes (FCB) group with most examples found in the phylum Bacteroidetes. This agrees with the phylogenetic distribution of 19 components of the T9SS, which is characterized by monophyletic clades representing all Bacteroidetes classes, and PorX is always present even in species possessing only the core subset of T9SS components (89). This argues that PorX is indispensable for the regulation of T9SS activity. In contrast, although the BREX system is widely distributed across bacterial phyla, it is only sporadically found in the phylum Bacteroidetes, with the exception of the genus Prevotella. Furthermore, whereas PorX and its orthologues are composed of 500-530 amino acids, BREX proteins featuring a PglZ domain are composed of at least 600. Despite the low level of identity between PorX and BREX-PglZ domains (19-25%), the latter possess a remarkably conserved fold with a high degree of similarity to the PglZ domain of PorX. The BREX-PglZ proteins are therefore phylogenetically related to PorX, and it is fascinating that although PorX and its closest homologue BREX-3 apparently evolved from a common ancestor, the BREX-3 system is not found in Bacteroidetes (7) whereas PorX is present in all classes within this phylum (89).
PorX and similar RR proteins in other bacteria are apparently important components of complex signalling pathways that involve protein-protein interactions and the cleavage of signalling nucleotides to modulate the activity of the T9SS. Our structural and functional characterization of PorX provides insight into the biological activity of orphan RR proteins from TCSs, which is based on the specific hydrolysis of cOAs and linear oligoadenylates by the PlgZ domain. This regulatory function might be independent from the interaction of PorX with SigP and PorL. It is tempting to speculate that protein-protein interactions directly affect T9SS activity, whereas the enzymatic activity of the PglZ domain is functionally related to the BREX PglZ proteins, as revealed by their sequence conservation and close but non-overlapping phylogenetic relationship. In the future, we will focus on AcP as a hypothetical regulator of PorX activity, the potential crosstalk between CRISPR-Cas and PorX, and the role of cOAs and/or linear oligoadenylates in the physiology of P. gingivalis and the function of PglZ-domain proteins in the BREX-3 system.

DATA AVAILABILITY
The data supporting the findings of this study are available from the corresponding authors upon reasonable request. Coordinates and structure factors have been deposited in the Protein Data Bank with accession codes 7PV7 (PorX-BeF 3 ), 7PVA (PorX-HR) and 7PVK (PorX-pGpG).