Nucleic Acids Research

Nucleic Acids Research Advance Access originally published online on May 13, 2008
Nucleic Acids Research 2008 36(11):3676-3689; doi:10.1093/nar/gkn170

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Nucleic Acids Research, 2008, Vol. 36, No. 11 3676-3689
© 2008 The Author(s)
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.0/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

Molecular Biology

Streptococcus pyogenes pSM19035 requires dynamic assembly of ATP-bound ParA and ParB on parS DNA during plasmid segregation

Florencia Pratto¹, Aslan Cicek², Wilhelm A. Weihofen², Rudi Lurz³, Wolfram Saenger² and Juan C. Alonso^1,*

¹Department of Microbial Biotechnology, National Centre of Biotechnology, CSIC, 28049 Madrid, Spain, ²Institute of Chemistry and Biochemistry / Crystallography, Freie Universität Berlin, 14195 and ³Max-Planck Institute for Molecular Genetics, 14195 Berlin, Germany

*To whom correspondence should be addressed. Tel: +34 91585 4546; Fax: +34 91585 4506; Email: jcalonso{at}cnb.csic.es

Received January 8, 2008. Revised March 19, 2008. Accepted March 25, 2008.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION SUPPLEMENTARY DATA REFERENCES

 
The accurate partitioning of Firmicute plasmid pSM19035 at cell division depends on ATP binding and hydrolysis by homodimeric ATPase ₂ (ParA) and binding of ₂ (ParB) to its cognate parS DNA. The 1.83 Å resolution crystal structure of ₂ in a complex with non-hydrolyzable ATPS reveals a unique ParA dimer assembly that permits nucleotide exchange without requiring dissociation into monomers. In vitro, ₂ had minimal ATPase activity in the absence of ₂ and parS DNA. However, stoichiometric amounts of ₂ and parS DNA stimulated the ₂ ATPase activity and mediated plasmid pairing, whereas at high (4:1) ₂ : ₂ ratios, stimulation of the ATPase activity was reduced and ₂ polymerized onto DNA. Stimulation of the ₂ ATPase activity and its polymerization on DNA required ability of ₂ to bind parS DNA and its N-terminus. In vivo experiments showed that ₂ alone associated with the nucleoid, and in the presence of ₂ and parS DNA, ₂ oscillated between the nucleoid and the cell poles and formed spiral-like structures. Our studies indicate that the molar ₂ : ₂ ratio regulates the polymerization properties of (•ATP•Mg²⁺)₂ on and depolymerization from parS DNA, thereby controlling the temporal and spatial segregation of pSM19035 before cell division.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION SUPPLEMENTARY DATA REFERENCES

 
Accurate distribution of newly replicated chromosomes before cell division is imperative for the stable transmission of genetic information. In eukaryotic cells, after chromosomal DNA condensation and alignment at mid-cell, microtubule fibers anchored via the kinetochore to the centromere pull the sister chromatids apart (1,2). In bacterial cells the mechanism that moves the newly replicated chromosomes and plasmids to opposite sides of the division plane requires a genuine partition system (ParA and ParB, ParM and ParR, or TubZ and TubR) (2–6).

For active and faithful segregation, most bacterial chromosomes and low-copy-number plasmids have evolved genuine partitioning (par) loci. The par loci contain one or more cis-acting DNA segment(s) (parS) and encode two trans-acting proteins: an ATPase motor protein and a centromere binding protein (3–5,7,8). Three evolutionary different plasmid partition systems have been identified: the tubulin-like (TubZ or type III), the actin-like (ParM or type II), and the Walker-box (ParA or type I) ATPases (4–6,9,10). The ParA system, which is the most common and conserved one, can be subdivided into two subfamilies (ParA-Ia and ParA-Ib) (4). The mechanism of action of ParA systems is less clear than that of the other mentioned systems, although a similar mechanism to the one observed with the actin-like systems has been suggested (3–5,7,8). Among the Proteobacteria phylum a large number of plasmid- and chromosome-encoded partition systems have been studied, e.g. plasmids P1, F and RK2 encode ParA-Ia ATPases (P1-ParA, F-SopA and RK2-IncC), while plasmids pB171 and pTB228 and the Caulobacter crescentus (Ccr) chromosome encode ParA-Ib ATPases (pB171-ParA, pTB228-ParF and CcrParA). However, among the Firmicutes phylum the ParA ATPases studied thus far (e.g. Streptococcus pyogenes pSM19035 and the Bacillus subtilis chromosome) encode ATPase of the ParA-Ib (₂ and BsuSoj) subfamily [(3–5), this work]. The ParA-Ia ATPases feature an N-terminal helix-turn-helix (HTH) motif that specifically interacts with DNA to repress the expression of the par loci, while the ParA-Ib ATPases bind non-specifically to DNA [e.g. ₂ and Thermus thermophilus Soj (TthSoj)] (3–5).

The ParB proteins are divided into three discrete subfamilies (ParB-I, -II and -III) (3–5). The ParB-I proteins (e.g. P1-ParB, F-SopB or RK2-KorB), and ParB-II proteins (e.g. BsuSpo0J or TthSpo0J) recognize parS DNA via an HTH fold and ‘spread’ around the parS site. The ParB-III proteins (e.g. pB171-ParB, pTB228-ParG, pSM19035-₂) work in concert with ParA-Ib ATPases and recognize parS (parC) DNA via a ribbon-helix-helix (RHH) motif. The cis-acting parS DNA consists of one (e.g. P1-parS, F-sopC), two (e.g. pB171-parC) or several copies of the centromeric parS site (e.g. pSM19035-parS, Bsu-parS) (3–5,8).

Plasmid partitioning has mainly been studied in species of the Proteobacteria phylum (3–5,8). The polymerization of (•ATP•Mg²⁺)₂ on DNA (see below), which is in stark contrast to ParA ATPases of proteobacterial plasmids that form proteofilaments in the absence of ParB and DNA, suggests that the dynamic movement of plasmid and bacterial chromosomes during faithful segregation in Firmicutes may not necessarily follow similar mechanisms as found for plasmids of Proteobacteria [(5), this work]. Indeed, the evolutionary distance between B. subtilis or Streptococcus pyogenes (Firmicutes) and E. coli ( Proteobacteria) exceeds that between plants and animals, and this raises the question whether bacteria of these two phyla share the same mechanism of plasmid partitioning. Here, we address this question by studying the segregation of plasmid pSM19035 originally isolated from the Firmicute and human pathogen S. pyogenes.

Plasmid pSM19035 replicates via a theta mechanism and is maintained stably at 1–3 copies per cell in B. subtilis, as well as in a wide range of species of the Firmicutes phylum (11–13). The par locus of pSM19035 encodes two trans-acting proteins, (ParA-Ib type) and (ParB-III type) and harbors six cis-acting parS sites [(14–16), this work]. Protein , which occurs as a homodimer (₂), shares sequence identity with bacterial and archaeal Walker-box ATPases, namely TthSoj and Pyrococcus furiosus MinD (PfuMinD) [(11), Figure S1 in the Supplementary Data available with this article online]. Protein , which occurs as a homodimer (₂), acts as a multifunctional repressor of genes involved in copy number control, plasmid addiction and accurate segregation [(15), this work]. Repressor ₂ negatively controls promoter utilization by binding cooperatively and with high affinity to the promoter regions upstream of copS, and genes (P_copS, P and P). These regions, which function as the cis-acting parS sites (parS1 or P, parS2 or P and parS3 or P_copS, Figure 1A and B), contain 10, 7 and 9 unspaced heptads with sequence 5'-WATCACW-3' in ( or ) orientations (15). The affinities of ₂ for the cognate sites P_copS, P and P are similar with a k_D of 6 nM [(17), Figure 1B]. The minimal cooperative yet high affinity binding sites for ₂ are two contiguous heptads in direct () or inverted () orientations (minimal centromere) (17).

View larger version (41K): [in this window] [in a new window] [Download PowerPoint slide]   Figure 1. Genome organization, nucleotide sequences of the 2-binding sites and proposed structure of the 2•parS1 (P) complex. (A) Organization of the genes that direct replication (repS) and its control (copS), segregation (, ) and post-segregation growth inhibition (, ). This DNA segment is duplicated in pSM19035 (14,15). The promoter (P) regions (Pcop, PIII, Prep, P, P and P,) of one arm of the plasmid, and the directions of the 2 binding sites (7-bp heptad repeats 5'-WATCACW-3', W is A or T, symbolized by or ) are identified by arrows. Repression of Pcop, P and P by 2 is indicated by the two ellipsoids. (B) Sequences of the Pcop (parS3), P (parS1) and P (parS2) regions with –35 and –10 regions are boxed and transcription start sites indicated by bent arrows. The heptads are indicated by arrows. (C) Model of nine 2 bound to parS DNA based on the crystal structures determined for [2N19]2-() and [2N19]2-() complexes (21) with the DNA shown in space filling (gray/blue) and 2 in ribbon representations (one monomer is orange, the other red).

 
The N-terminal region of protein ₂ is unstructured (18,19). Crystal structures have been determined for protein ₂ in which the monomers lack the first 20 N-terminal amino acid residues (₂N20) (20) and for ₂N19 in complex with two diheptads in () and () orientations (21). Chemical and enzymatic footprint data of ₂ binding to the centromere reveal a continuous protein super-structure consistent with the crystal structures (21). Extrapolating from these structures, ₂N19 molecules assemble as a left-handed protein helix that wraps parS sites consisting of multiple DNA heptad repeats (Figure 1C). The abilities of ₂N19 and wild-type (wt) ₂ to bind to parS DNA in vitro and to repress transcription in vivo are comparable, but substitution of Threonine 29 (that binds specifically to the central G–C base pair of the heptads) for Alanine (₂T29A) abolishes DNA binding (18,19,21).

Here, we provide the first crystal structure of a plasmid-encoded ParA-Ib type protein in the ATPS-bound state (•ATPS•Mg²⁺)₂ and show that plasmid pairing, polymerization of (•ATP•Mg²⁺)₂ on and depolymerization from parS DNA, which is dependent on wt ₂ bound to parS DNA, is fine tuned by the stoichiometry of ₂ and ₂. This is consistent with the dynamic assembly of a partition apparatus through the interaction of (•ATP•Mg²⁺)₂ with ₂•parS complexes and supports a model proposed here for DNA segregation in Firmicutes that is mediated by ₂ (ParA) plus ₂ (ParB) and differs from that in proteobacteria plasmids.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION SUPPLEMENTARY DATA REFERENCES

 
Plasmid stability test
The bacterial strains and plasmids used are listed in Supplementary Table S1. The numbers of plasmid-containing cells were determined by replica plating onto chloramphenicol-containing plates. The theoretical frequency of plasmid loss (L_th = 2^1–n) is the probability of plasmid-free cells arising per division, with n being the number of copies of the plasmid per cell at cell division (12). The frequency of plasmid loss (L) was calculated as L = 1 – (P)^1/g, where P is the number of cells bearing plasmids after growth for g generations.

Protein expression and purification
Proteins , N19 or T29A were expressed in E. coli BL21(DE3) pLysS cells and purified as described (17,18). Proteins , K36A, D60A or ₊₁₄ (having 14 extra N-terminal residues when compared to ) were expressed in E. coli ER2566 cells and purified by sequential heparin POROS 20HE (buffer A, 50 mM Tris–HCl, pH 8.0, 2 mM EDTA) containing 0.05 to 1 M NaCl concentrations, anion-exchange PL-SAX (buffer A containing increasing NaCl concentrations) and gel-filtration chromatography (buffer B, 20 mM Tris–HCl, pH 8.0, 200 mM NaCl). The protein concentrations were determined by absorption at 280 nm using molar extinction coefficients of 2980 M^–1 cm^–1 for ₂, ₂N19 and ₂T29A, and 38 850 M^–1 cm^–1 for ₂, ₂K36A and ₂D60A, and concentrations are specified for protein dimers.

Co-crystallization, data collection and structure determination
Crystals in space group P6₅22 grew at 18°C in sitting drop vapour diffusion setups from 1 µl protein solution (14 mg ₂/ml, buffer B with 2 mM ATPS and 5 mM MgCl₂) mixed with 1 µl of buffer C (1 M Hepes and 3% (v/v) ethanol pH 7.0). The mother liquor was supplemented with glycerol to a final concentration of 25% (v/v) prior to flash freezing the crystals in liquid N₂.

X-ray diffraction data were collected at 100 K at Protein Structure Factory beamline BL1 of Freie Universität Berlin at BESSY and processed with DENZO/Scalepack (22). The structure was determined by molecular replacement using a monomer of the TthSoj protein structure (pdb code 2BEJ) as a search model. The model of was built and water molecules were located with ARP/wARP (23). Restrained refinement cycles in REFMAC5 (24) converged at an R factor (R_free) of 19.2% (21.8%) (Table 1). Atomic coordinates and structure factors have been deposited with the Protein Data Bank under accession code 2OZE.

View this table: [in this window] [in a new window]   Table 1. Crystallographic data and refinement statistics

 
ATPase activity assay
ATPase activity was assayed by thin-layer chromatographic separation of the reaction products. Reaction mixtures (20 µl) contained buffer D [50 mM Tris–HCl, pH 7.5, 10 mM MgCl₂, 50 mM NaCl], 1 µM ₂, ₂K36A or ₂D60A, 10 µM ATP, 0–2.5 nM parS DNA, 0–4.2 µM ₂ or 1.4 µM ₂, ₂N19 or ₂T29A and were incubated for up to 180 min at 37°C.

Polymerization of protein ₂
Dynamic light scattering (DLS) was measured in a 1.5 mm path length quartz cuvette at 90° angle in arbitrary units using a Laser Spectroscatter 201 (RiNA GmbH, Berlin) with 350 nm emission wavelength and plotted using the FL Solutions computer programme and Savitsky-Golay smooth data processing. Aliquots of HindIII-linearized 3.1 kb pUC57-borne parS DNA (25 nM) were incubated for 2 min on ice with protein ₂ or variants ₂K36A or ₂D60A (1 µM) and different concentrations of ₂ (0, 0.24, 0.48, 0.96, 1.8 or 3.6 µM) or ₂19N (2 µM) in buffer E (50 mM Tris-HCl, pH 7.5, 1 mM MgCl₂, 1 mM DTE, 5% glycerol). After pre-incubation of the protein•DNA complex for 1 min, ATP, ADP or ATPS was added to 1 mM final concentration and light scattering was measured as above at 30 s intervals for 5 min and subsequently every 2 min at room temperature. The measured intensity or count rate was the amount of scattered light expressed as photons detected per second and converted to particle size using the Stokes–Einstein relation. The intensity is given in arbitrary units (AU).

Filament formation of protein ₂ or variants ₂K36A or ₂D60A (1 µM) in the absence or presence of fixed (1 µM) or variable concentrations of ₂, parS DNA or ATP•Mg²⁺ was measured in a 1 cm path length quartz cuvette as the change in turbidity using a Hitachi F2500 scanning fluorometer equipped with a thermo-cuvette holder at constant temperature (37°C). Relative light scattering intensities were recorded. Linear 3.1 kb pUC57-borne parS DNA (25 nM) was pre-incubated with protein ₂ (1.2 µM) and different ₂ concentrations (0, 0.24, 0.48, 0.96, 1.8 or 3.6 µM) in buffer E for 1 min at 37°C. Then, ATP was added to 1 mM final concentration, and the samples were used for light scattering. In presence or absence of linear pUC57-borne parS DNA (2 nM), wt ₂ or variants ₂K36A or ₂D60A (1 µM) and ₂ (1 µM) were pre-incubated in buffer E for 1 min at 37°C. The ATP was added to a 1 mM final concentration and the samples were used for light scattering.

Fluorescence and electron microscopy
Aliquots of B. subtilis cultures grown overnight in LB medium at 30°C were diluted in fresh medium to OD₅₆₀ 0.05. IPTG (10 µM final concentration) was added to OD₅₆₀ 0.2 cultures to induce the synthesis of (-GFP)₂ or (K36A-GFP)₂, and incubation was continued until OD₅₆₀ 0.6. Samples of the cells present were fixed and visualized as described (25). Images were acquired using an Olympus BX61 fluorescence microscope with an Olympus DP70 color CCD camera. Z-stacks of 20–25 images, separated by 0.1 µm, were collected and image deconvolution was performed using Huygens Professional software (Scientific Volume Imaging). DNA was stained using 0.2 µg DAPI/ml before microscopy.

EcoRI-linearized 3.1 kb pCB30 or pUC57 DNA (2 nM) harboring parS DNA was incubated with the desired protein(s) (see figure legends) for 15 min at 37°C in buffers D or E, respectively, in the presence or absence of 1 mM ATP, as previously described (26). The DNA–protein complexes were visualized by electron microscopy (EM) after negative staining with 1% uranyl acetate (27) or after fixation with 0.2% (v/v) glutaraldehyde for 10 min at room temperature. The procedures for adsorption of the complexes to mica, rotational shadowing with platinum and EM image evaluation have been described previously (28).

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION SUPPLEMENTARY DATA REFERENCES

 
Crystal structure of (•ATPS•Mg²⁺)₂
The crystal structure of ₂ bound to the non-hydrolyzable ATP analogue ATPS and Mg²⁺ was determined using the structure of the TthSoj monomer in molecular replacement. The crystal asymmetric unit contains one •ATPS•Mg²⁺ complex that forms a dimer (•ATP S•Mg²⁺)₂ with the two subunits related by a crystallographic C2 axis (Figure 2A and B). The dimer is stabilized by a hydrophobic surface patch that buries 2197 Ų of otherwise solvent accessible surface area per subunit, augmented by two reciprocal inter-subunit salt bridges formed between R119 and D189 of each monomer. The structure, refined at 1.83 Å resolution, includes all 284 residues of the wt protein . The recombinant version of this protein that was also used in genetic assays (₊₁₄, Table S1 in the Supplementary Data) carries additional 14 residues at the N-terminus that are disordered in the crystal structure (Table 1).

View larger version (89K): [in this window] [in a new window] [Download PowerPoint slide]   Figure 2. Structure of the (•ATPS•Mg2+)2 complex. (A) Structure of a monomer with -helices and β-strands shown as cylinders and arrows, respectively, and labelled sequentially. ATPS in sticks and Mg2+ as a purple sphere, N- and C-termini labelled. (B) The monomer shown in (A) is rotated by 90° about the vertical axis (black arrow) and forms (•ATPS•Mg2+)2 with the second monomer shown in blue. ATPS molecules as CPK models. A twofold crystallographic C2 axis (black vertical line with oval) relates both subunits in the crystal structure. (C) Superimposition of (•ATPS•Mg2+)2 (green/blue) with (Tth SojD44A•ATPS•Mg2+)2 (gray), viewed along the C2 axis. The monomers on the left were superimposed and the monomers on the right moved accordingly. (D) Electrostatic potential surface representation of (•ATPS•Mg2+)2. The bottom of the U-shaped dimer is negatively charged (left), whereas the upper tips are positively charged (right). (E) Electrostatic potential of the nucleotide binding pocket surface with bound ATPS and Mg2+ (green sphere). (F) Active site of (•ATPS•Mg2+)2 with the secondary structure elements indicated, hydrogen bonds drawn as yellow dashed lines and octahedrally coordinated Mg2+ as a magenta sphere. Lys36 and Asp60 that were substituted by Ala are labelled, water molecules as small red spheres, the water (labelled 1 near P150) is supposed to hydrolyze ATP.

 
Gel filtration and chemical cross-linking (Figure S2) confirmed that ₂ and the Walker A mutant ₂K36A (see below) form dimers in solution even in the absence of a nucleotide or the presence of ADP.

The monomer contains an eight stranded β-sheet surrounded by 12 -helices (Figure 2A). The N-terminal -helix (1) is not conserved in other Walker-box ATPases and shields the outward facing edge of the β-sheet (Figure 2B). The (•ATPS•Mg²⁺)₂ complex is U-shaped and each arm of the U represents one subunit with an ATP-binding site occupied by ATPS facing the cleft of the U (Figure 2A and B).

The surface charge of (•ATPS•Mg²⁺)₂ is negative near the bottom of the U and positive at the tips of the arms of the U (Figure 2D). The tip regions therefore most likely bind DNA and/or the negatively charged ‘bottom’ region of an adjacent ₂ when assembled into a ₂ polymer (see below).

The closest structural relatives of the monomer structure are TthSoj and PfuMinD although the underlying primary sequences exhibit only 25 and 14% identity, respectively (Figure S1 in the Supplementary Data). Superimposition of with 232 C atoms of TthSoj and 219 C atoms of PfuMinD shows root mean square deviations of 2.2 Å and 3.2 Å, respectively, indicating a high degree of structural similarity between these proteins. However, the structure of the dimer (•ATPS•Mg²⁺)₂ is significantly different from the hydrolysis-deficient (TthSojD44A•ATP•Mg²⁺)₂ variant.

TthSoj and PfuMinD dimerize only in the presence of ATP. In the dimers, each ATP molecule interacts with both monomers and becomes completely buried within the dimer interface (7,29). Based on these structures, it appears that ADP could only be released after dissociation of (TthSoj)₂ or (PfuMinD)₂ into monomers, whereas the wide and open cleft in the U-shaped (•ATPS•Mg²⁺)₂ allows free exchange of ATP and ADP without dissociation of the subunits.

The nucleotide-binding site
The ATP-binding sites in ₂ are positively charged (Figure 2E). The adenine N1 and amino group N6 of the two ATPS form hydrogen bonds to S240O and Y265O/K238O, respectively (Figure 2F). The C2'-endo puckered ribose is not engaged in hydrogen bonds but K36-S37-K38 within the Walker A motif at the N-terminus of helix 2 hydrogen bond with their peptide NH groups to - and β-phosphates of ATPS, whereas K36N forms salt bridges with the β- and -phosphates. Mg²⁺ is octahedrally coordinated by β- and -phosphate oxygen atoms, by S37O and by three water molecules (Figure 2E and F).

ATP hydrolysis requires a catalytic water molecule (W_cat) positioned in-line with the P-Oβ bond (W_cat···P-Oβ). In Walker-type ATPases, this W_cat is activated for nucleophilic attack on the -phosphate by an amino acid side chain in the Walker B motif that acts as catalytic base. However, in (•ATPS•Mg²⁺)₂, the position expected for W_cat is occupied by P150 within the Walker B motif (Figure 2F) but D60 of the Walker A' motif hydrogen bonds to and likely activates W_cat, which may in turn attack the -phosphate group of ATP (Figure 2F). This atypical positioning of amino acids likely participating in catalysis explains the relatively low ATPase activity of ₂ (see below).

The two above mentioned residues K36 and D60 (Figure S1) were considered to be engaged in the ATPase activity and replaced by Alanine to form ₂K36A and ₂D60A (see below).

The ATPase activity of ₂ is fine-tuned by ₂ levels in the presence of parS DNA and ATP
Since ₂, ₂ and cis-acting parS DNA interact and regulate pSM19035 segregation (see below), we investigated how the presence or absence of ₂ and/or parS DNA affect the enzymatic activity of ₂. Only experiments with parS2 DNA are described here because similar results were obtained with parS1 or parS3 DNA (Figure 1). The ATPase activity of ₂ was low and ₂ had no ATPase activity (Figure 3A). The ATPase activity of ₂ was not stimulated by the addition of parS DNA (₂ + parS, Figure 3A). Addition of stoichiometric amounts of ₂ stimulated the ATPase activity of ₂ by about 50% (₂ + ₂; Figure 3A). However, supplementation of parS DNA (₂ + ₂ + parS) resulted in a 3–4-fold stimulation of the ATPase activity of ₂, which was reduced to 2-fold when non-parS DNA was added (₂ + ₂ + non-parS, Figure 3A). However, when ₂ was added at nanomolar concentrations, the ATPase activity of ₂ was only stimulated by parS DNA (Figure 3C, see below).

View larger version (15K): [in this window] [in a new window] [Download PowerPoint slide]   Figure 3. Stimulation of the ATPase activity of 2 by 2 and DNA. (A) ATPase activity in reaction mixtures containing 1 µM 2 alone or with 2.5 nM parS DNA (2 + parS) or 1.4 µM 2 alone or after incubation of 1 µM 2 and 1.4 µM 2 (2 + 2) or additionally with parS DNA (2 + 2 + parS) (average of five experiments, n = 5) or with non-cognate DNA (2 + 2 + non parS) (n = 4). (B) ATPase activity of 2 after incubation of 1 µM 2 and 1.4 µM 2N19 or 2T29A with 2.5 nM parS DNA (2 + 2T29A + parS) or (2 + 2N19 + parS), or after incubation of 1 µM 2K36A or 2D60A and 1.4 µM 2 with 2.5 nM parS DNA (2K36A + 2 + parS) or (2D60A + 2 + parS). (C) ATPase activity of 1 µM 2 when incubated with increasing 2 concentrations (0.09–4.2 µM) and 2.5 nM parS2 DNA or non-cognate (non parS) DNA at 37°C. Values are averages of more than four independent experiments.

 
Stimulation of the ATPase activity of ₂ was marginal when ₂ was replaced by ₂T29A or ₂N19 (₂ + ₂N19 + parS; ₂ + ₂T29A + parS, Figure 3B). The ATPase activity of variants ₂K36A or ₂D60A amounted to 30% of wt ₂ activity and no stimulation of their ATPase activity was observed in the presence of ₂ and parS DNA (₂K36A + ₂ + parS or ₂D60A + ₂ + parS) (Figure 3B).

The stimulatory effect of increasing ₂ concentrations on the ATPase activity of ₂ in the presence of parS or non-parS DNA was also assayed (Figure 3C). In the presence of parS DNA and ₂ : ₂ molar ratios from 0.09:1 to 1.4 : 1 the ₂-catalyzed ATP hydrolysis was stimulated with the peak around 1.4 µM ₂, and the stimulation declined when the ₂ : ₂ ratio was further increased from 1.4 : 1 to 4.2 : 1 (Figure 3C). The observed characteristics of ATPase activity stimulation and alleviation of ₂ is genuinely associated with ₂ and parS DNA. In contrast, when parS DNA was replaced by non-parS DNA, increasing ₂ concentrations stimulated the ATPase activity of ₂ almost linearly (Figure 3C).

Pairing of parS regions by proteins ₂ and ₂
The complexes formed by ₂ and parS DNA, in the absence or presence of ₂, ₂N19 or ₂T29A, were visualized by EM at low protein concentrations (Figure 4B–D). The substrate was the linear 3.1-kb pCB30 DNA containing parS DNA located at 320 bp from one end (Figure 4A). In presence of ATP•Mg⁺², ₂ (100 nM) assembled to form discrete clusters on 85% of the DNA molecules (n = 200) at random locations (Figure 4B), whereas 40% of the DNA molecules (n = 250) that were incubated only with ₂ showed clusters of ₂ bound to parS on the plasmid (Figure 4C). The parS DNA region on linear DNA was not significantly distorted by ₂ binding, consistent with the prediction based on crystal structures that protein ₂ would wrap around parS sites without significantly bending the DNA double helix [(21), Figure 1C].

View larger version (58K): [in this window] [in a new window] [Download PowerPoint slide]   Figure 4. Electron micrographs of protein•DNA complexes. In the illustrated cartoons parS DNA, located 320 bp from one end (orange) of 3.1-kb linear pCB30 DNA, was incubated with 2 (red spheres) and/or 2 (blue spheres), in the presence of 1 mM ATP and the complexes formed were visualized by EM. Linear DNA (1 nM) alone (A), plus 2 (100 nM) (white arrows) (B), plus 2 (60 nM), showing 2•parS complexes (black arrows) (C). (D) Linear parS DNA (1 nM) incubated with 2 (100 nM) and 2 (60 nM) in the presence of 1 mM ATP with paired molecules indicated by empty arrows. The bars indicate 500 nm.

 
At 100 nM ₂ and 1 nM parS DNA but in the absence of (•ATP•Mg²⁺)₂ only 1% of the ₂•DNA complexes (n = 300) contained two DNA molecules that were paired at the position where ₂ was bound at the parS region. The frequency of these complexes was not increased by raising the ₂ concentration. However, when 1 nM parS containing DNA was incubated with 100 nM ₂, 60 nM ₂ and 1mM ATP•Mg²⁺, 20% of the parS DNA molecules were paired with DNA molecules juxtaposed at their ₂•parS regions (n = 200, Figure 4D), indicating that (•ATP •Mg²⁺)₂ is required for plasmid pairing (Figure S4A). Plasmid pairing was absent at a 2 : 1 ₂ : ₂ molar ratio (Figure S4B). Replacing ₂ with ₂N19 or ₂T29A or ₂ with ₂K36A abolished DNA pairing (Figure S4C–S4E).

₂ polymerization on parS DNA is dependent on ₂ and ATP
DLS data in Figure 5A show that (•ATP•Mg²⁺)₂ (1 µM) polymerizes onto linear 3.1-kb parS DNA in the presence of a 2 : 1 molar ratio of ₂ : ₂. No ₂ polymers were formed when either parS DNA was omitted (see below), protein ₂ was substituted by ₂N19, or ATP was substituted by ADP (Figure 5A).

View larger version (15K): [in this window] [in a new window] [Download PowerPoint slide]   Figure 5. Polymerization of 2 in the presence of parS DNA, 2 or 2N19 and ATP, ADP or ATPS. (A) Dependence of light scattering (AU) on time (min). Reaction mixtures containing linear 3.1-kb parS DNA (20 nM), 1 µM 2 and 2 µM 2 or 2N19 proteins were pre-incubated on ice in buffer E and DLS was measured at room temperature. At time zero, we added 1 mM of ATP to 2 and 2 (red line) and to 2 and 2N19 (green line), 1 mM ATPS to 2 and 2 (blue line) or 1 mM ADP to 2 and 2 (black line). (B) Light scattering increases with increasing 2 : 2 ratio. Reaction mixtures containing 3.1-kb parS DNA (20 nM), 1.2 µM 2 and the indicated 2 concentration were pre-incubated at 37° C in buffer E. Polymerization was initiated by the addition of 1 mM ATP (denoted by an arrow). Light scattering was measured at 37°C. Values are averages of three independent experiments. (C) Polymerization of protein 2 requires 2, parS DNA and ATP•Mg2+, as shown by an increase of light scattering. Reaction mixtures containing 3.1-kb parS DNA (20 nM), 1 µM 2, 2D60A or 2K36A and 2 µM 2 were pre-incubated at 37°C in buffer E. Polymerization was initiated by the addition of 1 mM ATP (denoted by an arrow). Light scattering was measured at 37°C. Values are averages of four independent experiments.

 
ATP binding but no hydrolysis is required for ₂ polymerization on parS DNA in the presence of ₂ because ATPS satisfied the cofactor requirement (Figure 5A). In presence of ATP or ATPS, the size increment of the polymers showed a sigmoidal pattern consistent with cooperative polymerization and levelled off after 60 min at room temperature. It remained at this level in the presence of ATPS (Figure 5A, blue line) contrasting the presence of ATP, where polymerization decreased to initial values after 90 min (Figure 5A, red line), indicating that ATP hydrolysis induces depolymerization, and the formed (•ADP•Mg²⁺)₂ complex did not support the integrity of the polymer. When fresh ATP was added to this solution after 120 min, ₂ polymerized again in a new cycle (data not shown). This indicates that the binding of ATP to ₂ enhances high affinity DNA binding (Table S2), and interaction of (•ATP•Mg²⁺)₂ with ₂•parS DNA leads to ₂ polymerization onto DNA, whereas ATP hydrolysis induced depolymerization. This is consistent with the observation, using atomic force microscopy, that no ₂ nucleoprotein filaments are formed onto linear or supercoiled parS DNA in the absence of (•ATP•Mg²⁺)₂ (F.P., K. Takeyasu and J.C.A., unpublished results).

EM revealed that (•ATP•Mg²⁺)₂ (1 µM) assembled and polymerized along the full-length of linear 3.1-kb parS-containing DNA molecules in presence of saturating amounts of ₂ (1 µM) (Figure S3A). However, such polymers were not observed when ATP was omitted (Figure S3B). The estimated protein volume of the nucleoprotein filament was only compatible with ₂ polymerization on DNA. Indeed, the molecular mass of ₂ is 4.3-fold larger than that the one of ₂ (the molecular masses are 68.8 and 15.9 kDa, respectively) (11,15).

Protein (•ATP•Mg²⁺)₂ polymerizes rapidly on parS DNA in the presence of an excess of ₂ at 37°C. However, no (•ATP•Mg²⁺)₂ polymers formed on parS DNA when the ₂ : ₂ molar ratio was 0.2 : 1 or below, suggesting that a minimal concentration of ₂ is needed under the experimental conditions used (Figure 5B). The extent of ₂ polymerization onto DNA increased with ₂ concentrations, because the light scattering signal was lower at ₂ : ₂ ratios of 0.75 : 1 compared to ₂ : ₂ molar ratios of 1.5 : 1 to 3 : 1 (black and blue lines; Figure 5B). Alternatively, at high ₂ : ₂ ratios, the elevated ₂ concentrations promoted ₂ polymerization even onto non-parS DNA.

Using 90° light scattering, we investigated the component requirements for ₂ polymerization (Figure 5C). In presence of ATP, no polymerization was observed when ₂ [(•ATP•Mg²⁺)₂ + parS] or parS DNA [(•ATP•Mg²⁺)₂ + ₂] were omitted or in presence of ATPS when protein ₂ was omitted (₂ + parS + ATPS) (Figure 5C), suggesting that ₂ polymerization on DNA requires the interaction with ₂. To further evaluate the effect of the nucleotide cofactor, the ₂D60A and ₂K36A variants were also analyzed. As shown in Table S2, protein ₂ or ₂D60A bound with 12-fold higher affinity to parS DNA in the presence of ATP than ADP, while binding of ₂K36A to parS DNA was weak, regardless of the presence of ATP or ADP. Wild type ₂ and variant ₂D60A feature similar polymerization kinetics (₂ + ₂ + parS vs ₂D60A + ₂ + parS) (Figure 5C), albeit the equilibrium was reached earlier in case of ₂D60A and the formed filaments were shorter. In contrast, variant ₂K36A showed a near-linear increase in light scattering or slowly assembled on parS DNA in the presence of ₂ and ATP (₂K36A + ₂+ parS; Figure 5C).

Plasmid segregation requires (•ATP•Mg²⁺)₂, ₂ and parS DNA
To test the importance of proteins ₂ and ₂ and a cis-acting parS site for plasmid segregation, we combined the respective genes and variants in the rolling-circle replicating and segregationally unstable vector pHP13 (Table S1). This vector was reported to replicate far from mid-cell and its replication imposes a metabolic burden that compromises its maintenance in host cells (30). The frequency of plasmid-loss was measured in B. subtilis cultured in LB medium at 30°C. Plasmids bearing a parS1 site and genes that directed the synthesis of ₂ and ₂ (pCB706) or ₂ and (-GFP)₂ (₂ with C-terminally fused GFP, pCB702) were retained in progeny cells at 10-fold higher frequencies (Table S3) than predicted if the plasmid were randomly distributed (see Materials and methods section). However, random distribution was observed if plasmids lacked either the or gene or carried genes that encoded variants K36A, N19 or T29A (Table S3). This indicates that the integrity of the ATP binding site of ₂, DNA binding and the N-terminus of ₂ and parS DNA are essential for correct pSM19035 partitioning.

Dynamic movement of protein (-GFP)₂ depends on parS DNA and ₂
To gain insight into the molecular mechanism by which the fully functional (-GFP)₂ (see above) contributes to plasmid segregation, we imaged its cellular localization in the presence or absence of ₂. The GFP signal overlapped with that of DAPI-stained DNA in 90% of the cells of a B. subtilis strain containing a pCB578-borne -gfp gene transcribed from its own P (parS1) promoter (Figure 6). When ₂ was also present (pCB702) it repressed the -GFP synthesis by 70-fold compared to the absence of ₂ (15). The low (-GFP)₂ signal was no longer statically associated with the nucleoid but was dynamically located near the cell poles and/or associated with the nucleoid. Image deconvolution showed that in the presence of parS DNA, ₂ and (-GFP)₂ a spiral-like structure was formed within the cytosol of B. subtilis cells (Figure 6B). It is of interest that unlike (-GFP)₂ that only formed a spiral-like structure in presence of parS DNA and ₂ under auto-regulated conditions, spiral-shaped filaments formed by other ParA (pB171-ParA or F-SopA) neither required ParB (pB171-ParB or F-SopB) nor parS (pB171-parC or F-sopC) DNA (31,32).

View larger version (38K): [in this window] [in a new window] [Download PowerPoint slide]   Figure 6. Subcellular localization of (-GFP)2 in the presence or absence of 2 and parS DNA. (A) The top illustration shows the structure of pCB578-borne P (parS1) and -gfp gene in B. subtilis cells. The lower panels show the localization of (-GFP)2 fusion protein in the absence of protein 2. Bars indicate 2 µm. (B) The top illustration shows the structure of pCB702-borne P (parS1) and -gfp gene, P (parS2) and gene in B. subtilis cells, and the repression by protein 2 is indicated by ellipsoids. The lower panels show the spiral-like organization of (-GFP)2 fusion protein in the presence of parS DNA, (-GFP)2 and 2. Images captured in different optical planes were subjected to 2D deconvolution. Bars indicate 2 µm.

 
To vary the intracellular concentration of (-GFP)₂ independently of the ₂ concentration, we placed the -gfp gene under the transcriptional control of the LacI repressor and integrated a single copy of this construct into the amy locus of the B. subtilis genome (Figure 7A). In the absence of a plasmid-borne parS site and of gene , (-GFP)₂ was seen to co-localize at low IPTG concentration (10 µM) with the nucleoid in 90% of the cells (n = 300), suggesting that (-GFP)₂ binds non-specifically to DNA (Figure 7B and 7B') and the fluorescence signal of (-GFP)₂ was 2- to 3-fold higher than the one from cells bearing the pCB578-borne parS1 site and gene -gfp (Table S1). Under this condition, the presence of a parS region and ₂ led to (-GFP)₂ re-localization (‘oscillation’) near one cell pole in 60% of the cells (n = 300) that had one nucleoid, or co-localized with one nucleoid in 70% of the cells (n = 200) that had two nucleoids (Figure 7C and 7C'). Similar results were reported for the chromosomally encoded BsuSoj in the presence of BsuSpo0J and parS sites (25,33).

View larger version (60K): [in this window] [in a new window] [Download PowerPoint slide]   Figure 7. Subcellular localization of (-GFP)2 in B. subtilis cells in the presence or absence of parS and protein 2. (A) Illustrations showing the structure of the (-GFP)2 expression cassettes integrated into the B. subtilis chromosome, and plasmids that had or lacked a parS2 sequence and encoded or the N19 variant. (B–F) Images of cells with fluorescence from (-GFP)2 or (K36A-GFP)2 and (B' to F') images of the same cells stained with DAPI to show DNA. Cells contained (-GFP)2 (B), (-GFP)2, 2 and parS DNA (C), (-GFP)2, 2N19 and parS DNA (D), (K36A-GFP)2 (E) and (K36A-GFP)2, 2 and parS DNA (F) were taken from exponentially growing cultures in the presence of 10 µM IPTG. Scale bar (in C') is 2 µm.

 
When ₂ was replaced by ₂N19 (Figure 7D and 7D') or by ₂T29A (data not shown), (-GFP)₂ co-localized with the nucleoid both in the presence or absence of parS DNA. Since ₂N19 binds to parS as well as wt ₂, contrasting ₂T29A that does not specifically bind to parS DNA (18,19,21), this shows that not only the ability of ₂ to bind parS, but the N-terminal region of ₂ is also required to stimulate the redistribution of (-GFP)₂ from the nucleoid to the cell poles.

At levels comparable to (-GFP)₂, the (K36A-GFP)₂ signal was distributed throughout the cells regardless of the presence or absence of a parS sequence and/or gene (Figure 7E, 7E', 7F and 7F'). This indicates that the binding of ₂ to DNA and oscillation in the cell requires ATPase activity. Indeed, variant (K36A-GFP)₂ bound to parS or non-parS DNA with a 6- to 7-fold lower affinity than wt ₂ (Table S2).

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION SUPPLEMENTARY DATA REFERENCES

 
ATPase ₂, structure and properties
We showed that the ATPase ₂ from the Firmicute plasmid pSM19035 and EcoMinD and TthSoj from Gram-negative bacteria have similar monomer structures, but form dimers under different conditions. EcoMinD and TthSoj are monomers in solution with or without ADP•Mg²⁺, and binding of ATP•Mg²⁺ induces formation of structurally similar dimers (7,29). In contrast, forms a dimer regardless of the presence of ADP•Mg²⁺ or ATP•Mg²⁺ or absence of a nucleotide cofactor (Figure S2) that is structurally different from the above two dimers (Figure 2B).

The ATPase activity of ₂ is regulated by binding of the ₂•parS DNA complex to ₂. When the concentrations of ₂ were increased up to 1.4 : 1 ₂ : ₂ ratios the ATPase activity was stimulated but above this ₂ : ₂ ratio, the stimulation diminished when the concentration of ₂ is further increased up to a ratio of 4.2 : 1.

Dynamic assembly of (•ATP•Mg²⁺)₂ on ₂•parS
Our results show that pSM19035-borne parS sequence(s), ATP binding and hydrolysis by ₂, and the N-terminus and the DNA binding ability of ₂ are essential components of the genuine pSM19035 partition system, because the mutation or deletion of either one of these components abrogates plasmid segregation. Protein (-GFP)₂ localized within the nucleoid of B. subtilis cells, whereas catalytically inactive (K36A-GFP)₂ was distributed throughout the cytoplasm, arguing for ATP•Mg⁺²-dependent nucleoid localization (Figure 7E and F). Under native regulation, (-GFP•ATP•Mg⁺²)₂, ₂ and parS DNA formed spiral-like structures (Figure 6B) by a mechanism that depends on the integrity of the ATPase activity of (-GFP)₂. However, in presence of parS DNA, a moderate excess of (-GFP)₂ over ₂ resulted in relocation or oscillation of (-GFP)₂ from the nucleoid to the cell poles. It is striking that spiral-like structures were only observed in the presence of (-GFP)₂, parS DNA and ₂ under autoregulated conditions. Possibly, the excess (-GFP)₂ bound to the nucleoid maske