Molecular Anatomy of ParA-ParA and ParA-ParB Interactions during Plasmid Partitioning

Background: Small homodimeric δ₂ (ParA) and ω₂ (ParB) and parS mediate pSM19035 partitioning.

Results: The δ₂ ATPase is a modular protein.

Conclusion: δ₂·δ₂ and δ₂·ω₂ interacting domains are juxtaposed.

Significance: ATP, nonspecific DNA, and ω₂ bound to parS induce multiple transitions on δ₂.

Abstract

Firmicutes multidrug resistance inc18 plasmids encode parS sites and two small homodimeric ParA-like (δ₂) and ParB-like (ω₂) proteins to ensure faithful segregation. Protein ω₂ binds to parS DNA, forming a short left-handed helix wrapped around the full parS, and interacts with δ₂. Protein δ₂ interacts with ω₂ and, in the ATP-bound form, binds to nonspecific DNA (nsDNA), forming small clusters. Here, we have mapped the ω₂·δ₂ and δ₂·δ₂ interacting domains in the δ₂ that are adjacent to but distinct from each other. The δ₂ nsDNA binding domain is essential for stimulation of ω₂·parS-mediated ATP hydrolysis. From the data presented here, we propose that δ₂ interacts with ATP, nsDNA, and with ω₂ bound to parS at near equimolar concentrations, facilitating a δ₂ structural transition. This δ₂ “activated” state overcomes its impediment in ATP hydrolysis, with the subsequent release of both of the proteins from nsDNA (plasmid unpairing).

Introduction

Faithful segregation of newly replicated genomes to daughter cells requires specialized partitioning machinery. In eukaryotic cells, the well known microtubule-based mitotic spindle apparatus, which guides the separating chromosomes to the different ends of the dividing cell, drives the chromosome segregation process (1, 2). In contrast, the mechanism of chromosome separation and plasmid partition in bacterial cells is not well understood. Three main types of systems have been described that ensure faithful partition of low copy number plasmids (reviewed in Refs. 3–6). Of these, the ParAB system is the most widespread in plasmids and is the only type present also in bacterial chromosomes (3–6). Unraveling the molecular basis of this plasmid partitioning system is therefore important to help us to understand chromosomal segregation.

The ParAB partition system requires three or more components: two trans-acting dimeric proteins (ParA and ParB), one or more cis-acting parS site(s), and, in most cases, the host's chromosomal DNA (3–6). The Walker box ParA ATPases can be subdivided into two distinct families based on their size or structure. The large (Lg)³ Walker box ATPases (e.g. Escherichia coli P1-ParA (Lg) or F-SopA (ParA-Lg) plasmids) bind and hydrolyze ATP, and then ParA-ADP specifically binds to its cognate site to regulate expression of the ParAB locus. When hydrolysis is delayed, ParA·ATP* binds nonspecific DNA (nsDNA), forming a carpet on the DNA in vitro (7–11). This gradient-like distribution of the ParA protein on the nucleoid is an essential step for faithful plasmid segregation in vivo (12). The small (Sm) Walker-box ParA ATPases, which lack the specific DNA binding domain, can be divided into two subgroups. The first group includes Streptococcus pyogenes pSM19035-δ₂ and Bacillus subtilis chromosome-encoded Soj (ParA-Sm), which in the ATP-bound form bind to nsDNA (13, 14). DNA binding by the small ParA ATPases (pSM19035-δ₂ and Soj) is a critical step for faithful segregation of plasmids and chromosomes (13, 15). In contrast, the small ParA ATPases of the second group (e.g. Salmonella newport TP228-ParF plasmid) form bundles in the absence of DNA (16, 17).

The structurally unrelated ParB centromere-binding proteins can also be classified into two groups. The first group includes large dimeric helix-turn-helix ParB proteins (e.g. P1-ParB, F-SopB, and chromosome-encoded Spo0J) that, upon binding to parS, spread over many kb, bridging, looping, and condensing the nsDNA (18–22). The second group includes the small dimeric ribbon-helix-helix ParB-like proteins (e.g. TP228-ParG and pSM19035-ω₂). Protein ω₂, upon site-specific binding to parS DNA, which comprise 7–10 contiguous iterons or heptads (Fig. 1A), assembles as a left-handed helix wrapped around the full parS site without significant spreading onto nsDNA and without distortion of the parS DNA (14, 23–25).

FIGURE 1.

Relevant genome organization, critical residues for δ₂ interaction with nsDNA, and structural comparison of ParA ATPases. A, genome organization of the duplicated part of plasmid pSM19035. The rep (involving copS, RNA III, repS, and the leading (oriS) and lagging (ssiA) replication origins) and segB loci (segB1 (ω, ϵ, and ζ) and segB2 (δ, ω, and parS sites, P_copS, P_δ, and P_ω)) are highlighted. The promoters (P), the mRNAs, and the genes of the relevant regions are shown and denoted as boxes, wavy lines, and rectangles, respectively. The upstream region of the promoters of the copS (P_copS), δ (P_δ), and ω (P_ω) genes, which constitute the cis-acting centromere-like parS sites, are enlarged. The variable number of contiguous 7-bp iterons (heptads) and their relative orientation are symbolized by arrows (→ or ←). The promoters repressed by ω₂ (double red spheres) are indicated. B, mapping of the nsDNA binding domain in δ₂. Electrostatic potential surface representation of δ₂ in the ATPγS·Mg²⁺-bound form (PDB code 2OZE) displayed using PyMOL. The surface charge of δ₂ is negative near the bottom of the U and positive at the tips of the arms of the U. The relevant α-helices involved in nsDNA binding (in green) map to the tips of the arms of its U-shaped structure. Relevant residues for nsDNA binding, residue Asp-60 (involved in ATP hydrolysis), and ATPγS·Mg²⁺ for one of the monomers are indicated, but they are repeated twice in the dimer structure. A lateral view highlighting the U-shaped form of δ₂ is shown. C, superimposition of full-length monomer structures of δ (in green) and T. thermophilus Soj (ParA-Sm) (in blue) (PDB code 2BEK), with P1-ParA (in yellow) (PDB code 3EZ6) and with TP228-ParF (ParA-Sm) (in orange) (PDB code 3K9H). The superimpositions were done using PyMOL.

The interaction of the ParA and ParB components, which leads to proper separation of plasmid copies, has been extensively studied in plasmids of the γ-Proteobacteria class. These studies provide the foundation for filament- and non-filament-based modes of plasmid segregation. In the filament-based modes, ParA (Sm), when bound to ATP, assembles into filaments or bundles, and the partition complexes are mobilized by linear contractile filaments reminiscent of the spindle mechanism in eukaryotes (thread pushing or pulling model) (Fig. 2A) (16, 17). Alternatively, ParA (Sm), in the ATP-bound form, assembles by forming nucleoprotein filaments, and the partition complexes are mobilized by helical or linear filaments as a cargo (filament-pulling model) (Fig. 2B) (26, 27). In the non-filament-based models (diffusion-ratchet and DNA relay models), ParA dimers or small oligomers bind to the nucleoid (Fig. 2C) (9–12, 28, 29). In the diffusion-ratchet model, a propagating ParA (Lg) ATPase gradient is the driving force for movement of the partition complexes (9–11), whereas in the DNA relay model, the forces that drive segregation are generated by a ParA (Sm) gradient and the elastic forces within the DNA molecule (29). The segregation machinery of plasmids and bacteria of the Firmicutes phylum, which are evolutionarily separated by more than 1,500 million years from Proteobacteria (a genetic distance larger than the one between humans and plants), is less understood. It is likely that the characterization of the mechanism of active partitioning of low copy number plasmids of the Firmicutes phylum might contribute to the general understanding of ParAB-mediated plasmid and chromosome segregation in bacteria.

FIGURE 2.

Different models of ParAB-mediated plasmid segregation. A, plasmid segregation by the thread-pushing or -pulling model. A bundle structure is composed of a ParA dimer of dimers interacting unit via the IF1 domain (equivalent to residues 121–196 of δ₂, in brown) with another two sets of ParA₂ proteins rotated 90° interacting via a hypothetical IF2 domain (equivalent to residues 235–258 of δ₂) that overlaps with the defined nsDNA binding domain (residues 211–262, in green). The bundle may grow until it encounters a ParB₂-like protein (in yellow; N terminus in red) bound to parS DNA (in gray). For simplicity, only one ParB dimer bound to parS is shown. B, plasmid segregation by the nucleoprotein filament-pulling model. ParA₂ polymerizes onto nsDNA (in light blue) via their nsDNA binding domains (in green). Polymerization will require the interaction of the positively charged N-terminal and negatively charged C-terminal regions. The formed nucleoprotein filament then captures and tethers plasmids by subsequent binding and release steps (denoted by an arrow with an open head). C, plasmid segregation by the diffusion-ratchet-like or DNA relay models. These non-filament-based models were not further discriminated. ParB₂ binds to parS, forming a transient complex, and the N terminus of ParB₂ is unstructured (in red). ParA₂ interaction with nsDNA and with ParB₂ bound to parS facilitates a structural transition in ParB₂ (highlighted by the change of the unstructured N-terminal region). Then a ParA blob on the nsDNA captures, tethers, and moves the plasmid onto the nucleoid. ParA, which is asymmetrically bound to the nucleoid, chases parS-bound ParB·ParA. This chasing leads to directional movement of plasmid-bound ParB toward the ParA gradient with the nucleoid playing a passive (diffusion-ratchet) or active (DNA relay model) role. The models were constructed using PyMOL (PyMOL Molecular Graphics System, version 1.5.0.4, Schrödinger, LLC, New York), δ₂·ATPγS·Mg²⁺ (PDB 2OZE), and ω₂·miniparS (PDB 1IRQ, 2BNW, 2BNZ, and 2CAX) as templates.

Plasmids of the ubiquitous inc18 vancomycin-, methicillin-, or erythromycin-resistant family encode two small homodimeric ParA-like (δ₂) and ParB-like (ω₂) proteins and 2–6 parS sites in order to ensure faithful segregation (30). Plasmid pSM19035 is the best characterized member of this family. The pSM19035 parS sites overlap the promoter regions of the copS, δ, and ω genes (Fig. 1A) (25, 31–33). In vitro, ω₂ and ω₂ΔN19, which lacks the unstructured N-terminal region, transiently bind with high affinity and cooperatively (K_D(app) = 5 ± 1 nm) to parS DNA, forming short-lived partition complexes (14, 32).

The structure of δ₊₁₄ (having 14 extra N-terminal residues) bound to ATPγS and Mg²⁺ includes all 284 residues of the WT δ protein (15). ATP-bound δ₂⁴ possesses a U-shaped structure, with one arm and a part of the joining region representing a monomer (Fig. 1B). The Walker domains face the cleft, and the bottom of the U-shaped dimer is negatively charged, whereas the C-terminal upper tips are positively charged (Fig. 1B) (15). Previous mutagenesis data showed that the nsDNA binding domain lies at the tip of the arm (Fig. 1B) (14). Protein δ₂ shows a low but significant degree of sequence identity with the P1-ParA₂ (Lg) (∼24%) and the Soj (ParA-Sm) (∼22%) ATPases, but the sequence identity drops to <20% when compared with the TP228-ParF (ParA-Sm) ATPase. However, the quaternary structures of these four ParA ATPases, three from Gram-negative and one from Gram-positive bacteria, are conserved (Fig. 1C).

Protein δ₂ binds to nsDNA, forming short dynamic complexes. These complexes have a blob shape containing up to 5 ± 1 δ₂/blob rather than forming filaments or nucleoprotein filaments (15, 24). Likewise, free δ₂ in solution (i.e. in the absence of nsDNA) forms discrete blob-shaped structures containing 2–3 δ₂ molecules rather than long bundles (24).

In vitro experiments have shown that at substoichiometric ω₂/δ₂ molar ratios, the interaction of apo-δ₂ with the unstructured N-terminal end of ω₂ bound to parS DNA increases the binding affinity of ω₂ for parS DNA by ∼8-fold, leading to a long-lived complex (14). Protein δ₂ bound to nsDNA, upon interacting with a ω₂·parS complex, loses its association with nsDNA and relocalizes with ω₂·parS to form a ternary complex (parS·ω₂·δ₂), followed by formation of quinary complexes (parS·ω₂·δ₂·ω₂·parS) (24). In vivo, however, the interaction of a ternary (parS·ω₂·δ₂) complex with a dynamic δ₂·nsDNA complex leads to plasmid-nucleoid pairing (parS·ω₂·δ₂·nsDNA) (34). At equimolar ω₂/δ₂ molar ratios, however, ω₂ stimulates δ₂-mediated ATP hydrolysis, causing subsequent dislodging of δ₂ from paired complexes, via a poorly understood mechanism (15).

At present, the δ₂ regions involved in oligomerization and in the interaction with ω₂ are poorly defined, and they have been mapped in this work. We show here that the δ₂·ω₂ and δ₂·δ₂ interacting domains are adjacent to but distinct from each other and suggest some functional transitions that are crucial for the activation of the δ₂ ATPase activity.

Experimental Procedures

Strains and Plasmids

The B. subtilis strain and the plasmids used for segregation studies are indicated in Table 1. The plasmids used for overexpression of proteins were propagated in E. coli BL21(DE3) (pLysS) and are listed in Table 1.

TABLE 1.

Strains and plasmids

^a Mid-copy number (7–9 copies/cell) pHP14 derivatives bearing the δ and ω genes (or their variants) under the control of their own promoters (P_δ (parS1) and P_ω (parS2); see Fig. 1).

The δ gene encodes two co-linear polypeptides, a 298-residues (δ₊₁₄) and a WT 284-residue product. The plasmid pCB746-borne WT δ gene was used for site-directed mutagenesis and discrete deletions from the N- and C-terminal regions.

The exposed and positively charged Lys residues at the C-terminal end of δ₂ were substituted by Ala (δ₂K242A, δ₂K259A/K260A) or Ser (δ₂K248S), and the negatively charged residue 211 was replaced by Ala (δ₂D211A), as described previously (34). Many of the initially designed C-terminal deletion mutants were insoluble. To overcome this inconvenience, δ₂ variants containing or lacking residues within random coil regions around the initially programmed C-terminal deletion were constructed. Finally, the codons at positions 255, 227, 197, and 164 of the WT δ gene were fused to His₆ codons, leading to a series of constructs truncated from the 3′-end.

Plasmid Copy Number and Plasmid Stability Test

The number of plasmid copies per cell was estimated by quantitative PCR. Normalization was done with two distinct chromosomal genes as described earlier (35). The number of plasmid-containing cells was determined by replica-plating onto chloramphenicol-supplemented LB plates. The frequency of plasmid loss was calculated as described previously (35).

Chemicals, Enzymes, Proteins, and DNA

All chemicals were pro-analysis grade and purchased from Roche Diagnostics (Mannheim, Germany). DNA restriction and modification enzymes and nucleotides were from New England Biolabs (Frankfurt, Germany) and Sigma-Aldrich (Madrid, Spain). Ultrapure acrylamide was purchased from Serva (Heidelberg, Germany). The ω₂ or δ₂ variants were purified as described for WT ω₂ and δ₂ proteins (15, 31, 33). pBC30-borne parS2 DNA (source of parS DNA) was purified as described (31). parS DNA is expressed as moles of DNA molecules, and this was estimated using a molar extinction coefficient of 6,500 m⁻¹ cm⁻¹ at 260 nm. The protein concentrations were determined by absorption at 280 nm using molar extinction coefficients of 2,980 m⁻¹ cm⁻¹ for ω₂ and ω₂ΔN19 and 38,850 m⁻¹ cm⁻¹ for δ₂ and its variants. Concentrations of all the proteins are expressed as moles of protein homodimers. All structural images were generated using the PyMOL Molecular Graphics System, version 1.5.0.4 (Schrödinger, LLC).

Protein Cross-linking

The cross-linking agent DSS was used to study protein-protein interactions as described previously (36). Cross-linking was performed by incubating parS DNA with δ₂ or its mutant variants in buffer B (50 mm Tris-HCl (pH 7.5), 50 mm NaCl, and 10 mm MgCl₂) supplemented or not with 1 mm ATP for 15 min at 37 °C. The assays were performed in the presence (+) or absence (−) of ω₂ prior to the addition of the indicated concentrations of DSS at various ω₂/δ₂ molar ratios. Alternatively, parS DNA was preincubated with one of the proteins (5 min, 37 °C), and the preformed complex was incubated with the second protein (5 min). Then DSS was added, and reactions were left for 10 min at 37 °C and then stopped by the addition of 10 μl of stop buffer C (50 mm Tris-HCl, pH 7.5, 400 mm glycine, 3% β-mercaptoethanol, 2% SDS, 10% glycerol) and subjected to SDS-PAGE. Cross-linked protein bands were excised from the gel and identified by tryptic digestion coupled to MALDI-TOF-TOF analysis as described (14).

Protein·DNA Complexes

For EMSA, gel-purified 423-bp α-³²P-labeled HindIII-KpnI parS DNA was incubated with various amounts of WT ω₂, WT δ₂ (or its variants), or both proteins together in buffer B containing or lacking 1 mm ATP or 1 mm ADP for 15 min at 37 °C in a 20-μl final volume. The reaction was stopped by the addition of loading buffer (1 mm EDTA, 0.1% bromphenol blue, and 0.1% xylene cyanol), and the samples were subjected to 4% or 6% PAGE. Gel electrophoresis was conducted using 1× TAE as running buffer, at 200 V, 4 °C, and the gels were dried prior to autoradiography.

To obtain K_D(app) values, the concentration of free DNA and protein·DNA complexes was densitometrically determined under non-saturating conditions from differently exposed autoradiographs of EMSA gels. Protein concentrations that transfer 50% of the labeled DNA into complexes are approximately equal to the K_D(app) under conditions where the DNA concentration is much lower than the K_D(app).

ATPase Activity Assays

The ATPase activity was assayed by thin layer chromatographic separation of the reaction products as described (15). The indicated proteins in the presence or absence of parS DNA were incubated for 180 min at 37 °C in buffer B containing 5 mm ATP in a 20-μl reaction mixture.

Results

The C-terminal Region of δ₂ Contributes to Plasmid Segregation

To investigate the contribution of the C-terminal region of δ₂ to faithful partitioning, a plasmid stability test was used. A set of plasmid-borne WT and mutant ω and δ genes, transcribed from their native promoters (P), termed P_δ (parS1) and P_ω (parS2) (Fig. 1A), were tested. Under the growth conditions used in this study, each cell contained, on average, ∼8 ± 1 plasmid copies, and the different mutations (see Table 1) did not affect plasmid copy number (data not shown).

As already observed, the ω and δ gene products, the presence of at least one parS site, ω₂·δ₂ interaction, ATP binding and hydrolysis, and the nsDNA binding domain of δ₂ were necessary and sufficient for stabilization of an otherwise unstable plasmid in B. subtilis cells (Table 2) (15, 37).

TABLE 2.

Effect of the different variants of the pSM19035 partitioning system in the faithful segregation of an unstable vector

B. subtilis cells containing the Par^- vector or its derivatives (8 ± 1 copies/cell) bearing the par locus of pSM19035 or its variants were grown in antibiotic-free LB medium at 30 °C, and the frequency of plasmid loss during exponential growth was measured after 100 generations. The genes were transcribed from their native promoters.

Par genes		Stability
ParB-like	ParA-like
		%
None	None	<1a
None	δ	<1a
ω	None	<1a
ω	δ	100a
ωΔN19	δ	<1a
ω	δK36A	<1a
ω	δD60A	<1a
ω	δΔN20	67.8
ω	δΔC255	29.4
ω	δΔC197	<1
ω	δΔC164	<1

^a This information was reported previously (15), but the experiments were reproduced here for comparison.

In the filament pulling model, the polymerization onto nsDNA as shown for pB171-ParA₂ (Sm) is essential (26). The structure and mutagenesis analysis of the symmetric δ₂ ATPase revealed that nsDNA binding domain occurs at the tip of the U (Fig. 1B) (14). If δ₂ polymerizes on nsDNA, it is likely that one of the external arms (e.g. the negatively charged C-terminal end) of δ₂ might be in close proximity with the opposite arm (e.g. the positively charged N-terminal end) of another δ₂ (Figs. 1B and 2B). To test this hypothesis, δ₂ variants lacking the N- (δΔN20) or C-terminal (δΔC255) region were constructed. Protein δ₂ΔC255 also lacks the putative IF2 domain predicted to be involved in bundle formation (Fig. 2A) (17).

When the δ gene was replaced by the δΔN20 variant, whose product lacks the first 20 residues, or by δΔC255, whose product lacks the last 29 residues (15), plasmid segregation was only marginally affected by ∼1.5- and 3-fold, respectively (Table 2). However, when the δ gene was replaced by the δΔC197 or δΔC164 genes, which not only lacked the negatively charged C-terminal domain but also lacked the nsDNA binding domain, faithful plasmid segregation was abolished (Table 2). These results suggest that the DNA binding domain is essential, but the N- and C-terminal ends are dispensable for accurate plasmid partitioning. These data were not consistent with the filament-based models (Fig. 2, A and B).

The δ Dimer of Dimer Interacting Domain Does Not Overlap with the nsDNA Binding Domain

To further evaluate the contribution of the C-terminal region to the different activities of δ (i.e. ATPase, nsDNA binding, and δ₂·ω₂ and δ₂·δ₂ interaction), the exposed and positively charged Lys residues at the C-terminal end of δ₂ (δK242A, δK248S, and δK259A/K260A) were substituted by Ala or Ser, the genes were cloned, and their products were overexpressed and purified (Fig. 3). To ascertain the importance of these basic residues for DNA binding, the exposed and negatively charged Asp-211 residue was replaced by Ala. The mutated gene was cloned, and its product (δD211A) was purified.

FIGURE 3.

The δ₂ nsDNA binding domain and the ω₂·δ₂ or δ₂·δ₂ interacting domain do not overlap. A, δ₂ variants form dimers in solution. Equal amounts of δ, δD211A, δDK242A, δDK248S, or δK259A/K260A (3 μg) were preincubated for 5 min at 37 °C in buffer B. Then increasing concentrations of DSS (0.01, 0.05, 0.1, 0.25, and 0.5 nm) were added, and the reaction was further incubated for 10 min. The products of the cross-linking reactions were separated by 15% SDS-PAGE. Running positions of the monomer (1), dimer (2), trimer (3), and tetramer (4) are indicated. B, ATPase activity of δ₂ protein and its variants (δ₂D211A, δ₂K242A, δ₂K248S, and δ₂K259A/K260A) alone (1 μm; white bars), in the presence of ω₂ (1.5 μm; gray bars), or in the presence of both ω₂ and parS DNA (1.5 μm and 25 nm, respectively; black bars). Values are the average of three or more independent experiments, and error bars indicate S.E. C and D, binding to nsDNA of the different δ₂ variants. The 423-bp α-³²P-labeled parS DNA (0.1 nm) was incubated with increasing amounts of δ₂ (0.009–0.3 μm), δ₂K248S and δ₂K242A (0.15 nm to 2.4 μm), and δ₂K259A/K260A (0.3–4.8 μm) for 15 min at 37 °C, in buffer B containing 1 mm ATP, and complexes were detected by EMSA and autoradiography.

In the presence of ATP, dimers and higher order oligomers were observed with δD211A, δK242A, or δK248S upon the addition of the DSS cross-linking agent, but δK259A/K260A only formed dimers (Fig. 3A) (14).

Proteins δ₂K242A, δ₂K248S, and δ₂K259A/K260A bound and hydrolyzed ATP at efficiencies comparable with the WT δ₂ or δ₂D211A (Fig. 3B). The addition of ω₂ bound to parS DNA significantly stimulated the δ₂-, δ₂D211A-, δ₂K242A-, or δ₂K248S-mediated ATP hydrolysis ∼4-fold (Fig. 3A). However, the stimulation was less evident (∼1.7-fold) in the case of the δ₂K259A/K260A variant.

Wild-type δ₂ binds nsDNA with a K_D(app) of 140 ± 31 nm, and δ₂D60A, which binds but does not hydrolyze ATP, binds nsDNA with a K_D(app) of 75 ± 19 nm (Table 3) (14, 15). ATPγS or ADP failed to promote δ₂ binding to nsDNA (Table 3). Removal of a negatively charged residue (δ₂D211A) from the nsDNA binding domain increased binding to nsDNA with K_D(app) of 25 ± 1.5 nm (Table 3). Removal of positively charged residues (δ₂K242A, δ₂K248S, or δ₂K259A/K260A) reduced binding to nsDNA as measured by EMSA (K_D(app) of >1.6 μm) (Fig. 3, C and D). However, electron microscopic analysis showed that the δ₂K242A mutant bound to the nsDNA with an efficiency comparable with WT δ₂, which suggests that the mutation does not affect the on rate of protein-nsDNA complex formation (34).⁵ It is likely that the δ₂K242A·nsDNA complex has a higher off rate than the WT δ₂·nsDNA complex (see below).

TABLE 3.

Preformed ω₂·parS complexes facilitate ternary complex formation with δ₂ variants

Protein	Nucleotide cofactor	Addition of ω2	KD(app)a nm
δ2b	None	No	>3,000
δ2b	ATPγS	No	>3,000
δ2b	ADP	No	>3,000
δ2b	ATP	No	140 ± 31
δ2	ATP	Yes	40 ± 4.2
δ2D60Ab	ATP	No	75 ± 19
δ2D60A	ATP	Yes	20 ± 5.3
δ2D211A	ATP	No	25 ± 1.5
δ2D211A	ATP	Yes	12 ± 0.9
δ2K242A	ATP	No	∼3,000
δ2K242A	ATP	Yes	50 ± 2.4
δ2K248S	ATP	No	∼1,700
δ2K248S	ATP	Yes	45 ± 6.8
δ2K259A/K260A	ATP	No	>3,000
δ2K259A/K260A	ATP	Yes	85 ± 9.1

^a A linear (α-³²P-labeled) 423-bp-long HindIII-KpnI parS2 DNA (0.1 nm) segment containing seven heptads ((→→←)₂←) (see Fig. 1) was incubated with increasing concentrations of the indicated protein for 15 min at 37 °C in buffer B containing or not containing 1 mm ATP, ATPγS, or ADP, and the K_D(app) was determined. The K_D(app) values (nm) are the average of at least three independent experiments and are within a 10% S.E.

^b δ₂ binding to DNA was reported elsewhere (14, 15) and determined here again for a direct comparison. When indicated, parS DNA was preincubated with ω₂ (24 nm), and then increasing concentrations of the indicated δ₂ were added, and δ₂-mediated complex formation or ternary complex formation was analyzed. Samples were separated by 6% PAGE, and the formation of parS·ω₂·δ₂ or δ₂·parS·ω₂ ternary complexes was quantified as described under “Experimental Procedures.”

Deletion of the nsDNA Binding Domain of δ Does Not Affect Dimer-Dimer Interaction

The δ genes coding for the previously mentioned deletion variants from the C-terminal end (Table 1) were cloned, and their products were overexpressed and purified. Proteins δ₂ΔC255 (which lacks the negatively charged C-terminal end and part of the predicted IF2 (17)), δ₂ΔC227 (which also lacks the ATP-interacting residues), δ₂ΔC197 (which also lacks the nsDNA binding domain), and δ₂ΔC164 (which also lacks part of the hypothetical IF1 region) were analyzed under in vitro conditions (Fig. 4A).

FIGURE 4.

The δ DNA binding domain is not essential for interaction with ω₂ and for dimer-dimer interaction. A, constructed C-terminal deletions of δ₂. The Walker A, A′, and B boxes are highlighted in red, and the nsDNA binding domain is highlighted in green, and the hypothetical IF1 and IF2 regions are denoted by dotted lines. The ω₂·δ₂ and δ₂·δ₂ interacting domains mapped in this report are shown in purple. The position of the last residue present in each variant is indicated. B, proteins δ, δΔC255, δΔC227, δΔC197, or δΔC164 (3 μg) formed dimers and higher order oligomers in solution when incubated with the cross-linking reagent DSS (0.5 mm), as shown by 15% SDS-PAGE. C, ATPase activity of δ₂ and its variants (δ₂ΔC255, δ₂ΔC227, or δ₂ΔC197 and δ₂ΔC164) alone (1 μm; white bars), in the presence of ω₂ (1.5 μm; gray bars), or in the presence of both ω₂ and parS DNA (1.5 μm and 25 nm, respectively; black bars). D and E, interaction of δ₂ variants with ω₂ and parS DNA. In D, the 423-bp α-³²P-labeled HindIII-KpnI parS DNA (0.1 nm) was incubated with different amounts of ω₂ (0.75 and 6 nm), a fixed amount of δ₂, δ₂ΔC255, or δ₂ΔC227 (140 nm) or with a fixed amount of ω₂ (0.75 nm) and increasing concentrations of δ₂ (35–140 nm) for 15 min at 37 °C, in buffer containing 1 mm ATP. In E, α-³²P-labeled parS DNA was incubated with a fixed amount of δ₂ΔC197 or δ₂ΔC164 (140 nm) or a fixed amount of ω₂ (0.75 nm) and increasing concentrations of δ₂ΔC197 or δ₂ΔC164 (35–140 nm) for 15 min at 37 °C in buffer B containing 1 mm ATP. Protein·DNA complexes were detected by EMSA. The free DNA (FD) and the formed complexes are indicated. F, the δ DNA binding domain is not essential for interaction of ω₂ with parS DNA. The percentage of ω₂·parS DNA complex formed in the presence of limiting ω₂ concentrations (0.75 nm) and increasing concentration of δ₂ or C-terminal δ₂ variants is shown. Values are the average of three or more independent experiments.

As revealed in Fig. 4B, the δ₂ C-terminal deletion mutants (δΔC255, δΔC227, δΔC197, and δΔC164) formed dimers and high order oligomers upon the addition of DSS. These data were not consistent with the proposed C-terminal dimer-dimer interface domain (IF2), as deduced from the structure of ParF (ParA-Sm) polymers (17). Because δ₂ΔC197 could still oligomerize (Fig. 4B), the effect of δK259A/K260A on protein oligomerization might be indirect (e.g. via thermal unfolding etc.) (Fig. 3A).

The ATPase activity of the δ₂ variants (δ₂ΔC227, δ₂ΔC197, and δ₂ΔC164) was not stimulated by the addition of parS DNA or parS bound to ω₂ (Fig. 4C). Upon addition of ω₂ bound to parS DNA, the ATPase activity of δ₂ΔC255 was slightly stimulated (Fig. 4C). This is consistent with crystallographic data that show that three δ residues, Lys-238, Ser-240, and Tyr-265, which map to the C terminus, are also involved in ATP binding. Ser-240 forms hydrogen bonds with the adenosine, and Lys-238 and Tyr-265 form hydrogen bonds with the amino group of the two ATPγS molecules in the δ₂-ATPγS structure (see below) (15). Thus, all three residues have activities that are crucial for full δ₂-mediated ATP hydrolysis, and δ₂ΔC255, which only lacks Tyr-265, shows an intermediate phenotype (Fig. 4C).

The δ₂·ω₂ Interacting Domain Does Not Map to the C-terminal End of δ₂

Protein ω₂ binds to parS, forming transient ω₂·parS complexes (K_D(app) of 5 ± 1 nm, half-life <1 min) (24, 32). The interaction of ω₂ bound to parS with δ₂ or apo-δ₂ led to longer-lived ω₂·parS complexes (K_D(app) 0.7 ± 0.1 nm, half-life ∼34 min) (14). It is likely that a poorly defined region of δ₂, upon interacting with the unstructured N-terminal domain of ω₂, facilitates structural transitions that enhance the stability of ω₂·parS complexes (14). To test whether ω₂ interacts with δ₂ through its C-terminal domain and whether such protein-protein interaction facilitates the binding of δ₂ or ω₂ to parS DNA, two different approaches were undertaken. First, in the presence of ATP, preformed ω₂·parS DNA complexes were incubated with limiting δ₂, δ₂K242A, δ₂K248S, or δ₂K259A/K260A concentrations. The preformed ω₂·parS complexes enhanced ternary complex formation with WT δ₂ (ω₂·parS·δ₂) to K_D(app) 40 ± 4.2 nm as compared with K_D(app) = 140 ± 31 nm with δ₂·nsDNA alone (Table 3). When WT δ₂ was replaced by δ₂K242A, δ₂K248S, or δ₂K259A/K260A, the preformed ω₂·parS complexes enhanced the formation of ternary complex from K_D(app) of >1600 nm to K_D(app) of 40–85 nm (Table 3). Second, in the presence of limiting ω₂ concentrations (0.75 nm), C-terminal deletion mutants, which lacked the DNA binding domain, cannot form ternary complexes but may help ω₂ to bind to parS DNA. In the presence of limiting ω₂, δ₂ΔC255, δ₂ΔC227, δ₂ΔC197, or δ₂ΔC164 enhanced ω₂·parS complex formation from K_D(app) of 5 ± 1 nm to K_D(app) of ≤0.75 nm (Fig. 4, D and E). The interaction of ω₂ with δ₂ΔC164 facilitated the formation of ω₂·parS complexes, but these protein·DNA complexes were found to be less stable in comparison with the WT complex (Fig. 4, E and F). These results suggest that the deletion mutants still interact with ω₂ and facilitate ω₂·parS complex formation. To compare these results, the data presented in Fig. 4, D and E, were quantified. As revealed in Fig. 4F, the concentration of δ₂ (or its mutant) sufficient to facilitate ω₂·parS DNA complex formation was significantly enhanced, with a K_D(app) of 30–50 nm, which is 3–5-fold below the K_D(app) for δ₂·nsDNA (K_D(app) of 140 ± 31 nm). It is likely, therefore, that the δ₂ variants interact with ω₂ with an efficiency similar to that of WT δ₂ (Fig. 4F). These data altogether support the conclusion that (i) interaction of the preformed ω₂·parS DNA complex with δ₂K242A, δ₂K248S, or δ₂K259A/K260A facilitates ternary complex formation; (ii) the C-terminal region of δ₂ (residues 164–284) is dispensable for interaction with the ω₂·parS DNA complexes; (iii) removal of the 120 C-terminal residues from δ₂ still enhances ω₂·parS complex formation, but the complexes formed are less stable (Fig. 4E); and (iv) the δ₂ concentration required for interaction with ω₂ bound to parS DNA (Fig. 4, D and E) or to facilitate ω₂·parS complex formation (Fig. 4F) is 3.5–4-fold lower than for interaction with nsDNA.

The Central Domain of δ₂ Interacts with ω₂

To identify the region(s) of δ₂ involved in the interaction with ω₂, both proteins were cross-linked in vitro by DSS in the presence of parS DNA, and the different protein bands were gel-purified (Figs. 5A (a–g) and 6A (a–f)), subjected to limited proteolysis, and analyzed by MALDI-TOF-TOF as described previously (14).

FIGURE 5.

A central domain of δ is required for ω₂·δ₂ interaction. A, DSS cross-linking assay showing the interaction between ω₂ (1 μg) and δ₂ (or δ₂ΔC227), at 0.5:1 and 1:1 ω₂/δ₂ ratios, in the presence of parS DNA. Cross-linked products were separated by 10–15% SDS-PAGE. Bands a–g were gel-purified and subjected to limited trypsin proteolysis, followed by MALDI-TOF-TOF identification of the polypeptides. B, the identified peptide sequences are shown in light (ω) or dark gray (δ) boxes, and the missing regions are shown in dotted lines. C, the relevant region of δ involved in the interaction with ω or with itself is highlighted in the crystal structure in purple (α4-α5-β3-β4-α6). In this structure, ATPγS·Mg²⁺ is shown in orange and yellow, and residue Asp-60 (Walker B′) is shown in green (PDB 2OZE, displayed using PyMOL). The experiments were performed more than three independent times.

FIGURE 6.

A central domain of δ is required for δ·δ and δ₂·δ₂ interaction. A, the δ₂·δ₂ interaction requires the presence of ω₂ and parS DNA. Preformed ω₂·parS DNA (ω₂ (3 μg), lanes 1–3) or δ₂·parS DNA complexes (lanes 4–6) were incubated with δ₂ or ω₂ as indicated (at a 3:1 ω₂/δ₂ ratio in lane 3 or at a 0.3:1 ω₂/δ₂ ratio in lane 6), and the specific interactions were analyzed using DSS. Cross-linked products were resolved by 10–15% SDS-PAGE. The stained protein bands e (∼40 kDa) and e′ (∼48 kDa) were gel-purified and subjected to limited trypsin proteolysis and MALDI-TOF-TOF analysis of the polypeptides. Tetrameric to hexameric complexes are marked (lanes 4–6). B, δ·δ interaction in the absence of nsDNA. The DSS-cross-linked product (lane 2) was separated by 10–15% SDS-PAGE. Band h was gel-purified and subjected to limited trypsin proteolysis, followed by MALDI-TOF-TOF identification of the polypeptides. C, the identified peptide sequences are shown in boxes, and the missing regions are shown in dotted lines. The N- and C-terminal regions are denoted.

With this technique, under high sequence coverage, the regions not involved in protein-protein interaction should be proteolyzed by trypsin, and the generated polypeptides should be detected with the expected molecular mass. On the other hand, if trypsin does not reach the covalently bound protein-protein regions, the expected polypeptide(s) should be absent. Alternatively, the interacting polypeptides, due to cross-linking among them, will have a higher molecular mass and therefore will also be absent. It is worth noting that our analysis was qualitative; hence, we considered a region involved in the protein-protein interaction if there was no detection of the polypeptide corresponding to that region, although it was present in the control (i.e. in the absence of the interacting partner).

First, ω₂-bound to parS DNA was analyzed. Protein bands a, b, and c in Fig. 5A corresponded to monomers, dimers, and tetramers of ω₂, respectively. In band a, the polypeptide corresponding to the β-strand (residues 22–31) of ω₂ (7.9 kDa, peptide coverage ∼89%), in the presence of parS DNA, was not observed (Fig. 5B). This finding is consistent with data showing that ω₂ interacts with parS DNA through this β-strand (25).

The cross-linked δ proteins in the presence of nsDNA, leading to bands d, e, f, and g, were gel-purified, subjected to limited tryptic digestion, and followed by MALDI-TOF-TOF analysis (Fig. 5, A and B). Here, δ polypeptides encompassing intervals 139–215 and 242–265 were not detected (Fig. 5B). Within the 139–215 interval, which lacks trypsin sites, the δ·δ interaction was previously mapped (15). Indeed, in the crystal structure, a δ dimer is stabilized by a hydrophobic surface patch of otherwise solvent-accessible surface area, augmented by two reciprocal intersubunit salt bridges formed between Arg-119 and Asp-189 of each monomer (15).

Within the 242–265 interval, which contains three trypsin sites, the nsDNA binding domain was previously mapped (14). The cross-linked ∼64-kDa protein in the absence of nsDNA (band h) was gel-purified, subjected to limited tryptic digestion, and followed by MALDI-TOF-TOF analysis (Fig. 6B). In the absence of nsDNA, a δ polypeptide encompassing the 242–247 interval was detected (Fig. 6C). However, these trypsin sites were not seen in δ₂ bound to nsDNA (Fig. 5B, band f). It is likely that the δ₂-δ₂ conformation adopted in the presence of nsDNA blocks trypsin's approach to these cognate sites following removal of the nsDNA.

When bands d and f (∼32 and ∼64 kDa), which corresponded to δ monomers and dimers, respectively, were analyzed, six (band d) and seven (band f) discrete regions of δ were not observed. In these bands, the sequence coverage was 57.0 and 54.5%, respectively (Fig. 5B). Of these, only one discrete region, encompassing residues 1–7, was not observed in the ∼64-kDa protein (band f). The δ polypeptides encompassing residues 31–36 (Walker A), 67–71 (Walker A′), 116–119, 139–215, 240–265, and 275–284 were not detected in bands d and f.

In the presence of preformed ω₂·parS DNA complexes, at about stoichiometric ω₂/δ₂ ratios, δ and ω formed a new discrete band e (Fig. 5A, lanes 3 and 4). When the order of addition was altered or the relative molar ratio of ω₂/δ₂ was modified (3:1 or 0.3:1), bands e (as in Fig. 5A) and e′ were observed (Fig. 6A, lanes 3 and 6). Limited trypsin proteolysis and MALDI-TOF-TOF analysis of this gel-purified band e (∼40 kDa; Figs. 5A and 6A) showed that this novel polypeptide was composed of ω (7.9 kDa) and δ (32.6 kDa) cross-linked in a 1:1 stoichiometry with sequence coverage of 65.8% for δ and 81.3% for ω. The gel-purified band e′ (∼48 kDa) (Fig. 6A, lanes 3 and 6) was composed of two ω monomers and one δ.

When band e was analyzed, six discrete regions of δ and two discrete regions of ω were not observed (Fig. 5B). Polypeptides 1–10 and 47–52 (coverage >77%) of ω were missing. We propose that the unstructured N-terminal region of ω₂ bound to parS, upon interaction with δ₂ (Fig. 5B), undergoes a structural transition that might involve α-helix formation, as suggested by protein structure predictions (34). These ω·δ complexes might be stabilized by residues 47–52 of ω, because this region was also not detected when both proteins were incubated (Fig. 5B). When δ peptides missing in the gel-purified 40 kDa band e were compared with those of bands d (δ) and f (δ₂) (coverage >60%), the two discrete segments, encompassing residues 88–119 and 216–223, were also not detected, suggesting that these regions may correspond to the ω-interacting region (Fig. 5, B and C). Within the 88–119 missing interval, there is a potential trypsin cleavage site at position 116. Because we have not identified the cross-linked product, we cannot rule out the possibility that the 88–119 or the 88–116 region could be targeted by ω. Consistent with this, the region consisting of residues 88–116 or 88–119, comprising the α4–α6 interval, is surface-exposed in the δ₂ structure (Fig. 5C) (15).

Protein δ₂ΔC227 was incubated with ω₂·parS DNA, and then DSS was added. A novel band of ∼34 kDa (denoted as band e*), which was composed of one ω and one δΔC227 monomer, was observed at substoichiometric ω/δΔC227 ratios (Fig. 5A, lanes 7 and 8). In band e*, four discrete regions of δ and two of ω were not observed by MALDI-TOF-TOF analysis (Fig. 5B). When the peptides absent in this gel-purified 34 kDa band were compared with those absent in bands d and f, again the region detected previously with the WT protein (88–119 or 88–116 interval) was missing. The other potentially interacting region, the 216–223 interval, was present when δ₂ was replaced by δ₂ΔC227. Therefore residues 216–223, which were previously attributed to DNA binding, were not further evaluated. It is likely that the main ω·δ interacting domain maps to the central region of δ₂ or δ₂ΔC227 (residues 88–119; the α4, α5, β3, β4, and α6 interval) and is oriented opposite the face of the DNA binding domain (Figs. 5C and 7C). In contrast, using yeast or bacterial two-hybrid systems, it was suggested that the ParA·ParB interacting domain of the (F-SopA (ParA-Lg)) or the (pSM19035-δ₂) ParA ATPase maps to the C-terminal end (i.e. residues 198–284) (38, 39). Because the previous results suggested that δ₂ΔC197 still interacts with ω₂ bound to parS (Fig. 4E), it will be of significant interest to determine the F-SopA·SopB (ParA·ParB-Lg) interacting domain by other methodologies.

FIGURE 7.

Summary of the nsDNA, ω·δ, and δ₂·δ₂ binding motifs of δ₂ identified in this work. A, ribbon diagram of δ₂ with key motifs highlighted in green, cyan, purple, and brown. From left to right, δ₂ was rotated 90° to highlight the different motifs. Boxes of broken lines indicate the interacting regions. B and C, sequence alignment of relevant regions of the small ParA ATPases (δ₂, Thermus aquaticus Soj and plasmid TP228 ParF). B, nsDNA binding motif (α10 to part of α12; green). C, sequence alignment of the δ₂ region involved in δ₂·δ₂ (α3 to α4; cyan) and ω·δ (α4 to α6; purple) interaction derived from our cross-linking experiments (see Figs. 5A and 6A). Sequence alignments were generated using ClustalW and displayed using Jalview version 2.7.

The Central Region of δ₂ Is Involved in Dimer-Dimer Interface

To define the region(s) of δ₂ involved in dimer of dimer formation, δ₂ was preincubated with preformed ω₂·parS DNA complexes, the ternary complexes were treated with DSS, and the newly covalently bound protein was analyzed as described above (Fig. 5A, termed band g). A comparison of bands g (∼98 kDa, δ₂·δ) with bands d (∼32 kDa, δ) and f (∼64 kDa, δ₂) revealed that one discrete region, encompassing residues 68–119, was not detected in band g (Fig. 5B). This region constitutes the α3–α6 interval in the crystal structure and is an exposed surface distant from the DNA binding domain (Figs. 5 (B and C) and 7 (A and C)). However, the 68–119 interval partially overlaps the ω·δ interacting domain (residues 88–119; interval α4–α6) (Figs. 5B and 7C). The δ₂·δ interacting region is close to the ATP binding and hydrolysis domains (Walker A′ (residues 51–61) and Walker B (residues 142–147)) (Fig. 7, A and C).

Discussion

pSM19035 partitioning, which depends on the dynamic interaction between the δ₂ ATPase, ω₂, parS, and the host chromosome, is a multistep process with discrete functional and/or structural transitions necessary for faithful segregation. The small Walker box ATPase δ forms dimers that are stabilized by a hydrophobic patch that occupies about 2197 Ų/subunit of otherwise solvent-accessible surface and augmented by two reciprocal intersubunit salt bridges formed between Arg-119 and Asp-189 of each monomer (see Ref. 15). Protein δ₂, upon binding to nsDNA, forms discrete clusters on the nsDNA (4–6 δ₂/blob) (24), perhaps by interacting through the 68–119 region with other δ₂ molecules (see Fig. 7, A and C). In contrast, using a bacterial two-hybrid assay, the δ monomer-monomer interface was mapped to residues 104–105 (39), and using yeast two-hybrid and immunoprecipitation assays, the chromosome-encoded Pseudomonas aeruginosa ParA (Sm) monomer-monomer interface was mapped to an interval equivalent to residues 89–105 of δ₂ (40). To reconcile the apparent discrepancy between the crystal structure of δ₂·ATPγS·Mg²⁺ (15), our data (Fig. 5B), and those published by other groups (39, 40), we propose that the bacterial or yeast two-hybrid assays cannot discriminate between the monomer and the dimer interface and that the authors were mapping the dimer-dimer interface (see Fig. 7A, center).

The interaction of the N-terminal region of ω₂ (residues 1–10) with the central region of δ₂ (residues 88–119) leads to a discrete ternary (ω₂·parS·δ₂) complex (Figs. 5C and 7, A (left) and C). However, if parS DNA was omitted or ω₂ was replaced by ω₂ΔN19, such protein-protein interaction was not observed (14, 33). We propose that the unstructured N-terminal region of ω₂ bound to parS, upon interaction with δ₂, undergoes a structural transition with residues 1–9 folding into an α-helix, as predicted by different protein folding prediction programs (34). Such interaction, at stoichiometric ω₂ and δ₂ concentrations, may facilitate δ₂-mediated ATP hydrolysis and δ₂ disassembly from the paired (parS·ω₂·δ₂·nsDNA) complexes. In the presence of an excess of ω₂, the ATPase activity of δ₂ is inhibited (15), and the plasmid can be engaged in a new pairing complex with a δ₂ in the nucleoid.

While the functional characterization of the ω₂·δ₂ interacting domains was being carried out during this study, the interacting domain of chromosome-encoded P. aeruginosa ParA (Sm) with ParB (Lg) was published (40). These authors mapped the ParB·ParA interacting region to within an interval equivalent to residues 89–105 of δ₂, by yeast two-hybrid and immunoprecipitation assays (40). This is in good agreement with the ω·δ interacting region reported here (Fig. 5B). However, it was unexpected because equivalent regions in both small ParA ATPases share very little sequence identity (<15%). Furthermore, plasmid- and chromosome-encoded large ParB proteins, which are of the helix-turn-helix family, bind specifically and nonspecifically to DNA and spread over many kb, leading to bridging, looping, and condensing of the nsDNA (18–21). Conversely, the plasmid-encoded small ParB proteins (e.g. ω₂), which are ribbon-helix-helix proteins, specifically interact with parS without significant spreading (24). It is likely that the unrelated P. aeruginosa ParB (Lg) and ω₂ might expose an equivalent domain recognized by an equivalent region in the ParA (Sm) ATPase.

The results presented here are inconsistent with the ParAB filament-based (thread-pushing or -pulling (Fig. 2A) or filament-pulling (Fig. 2B)) models. In the thread-pushing or -pulling model, the central (IF1; see Fig. 4A) and the C-terminal regions (IF2) of the TP228-ParF (ParA-Sm) (Fig. 7B) should be involved in bundle formation, as depicted in Fig. 2A (16, 17). However, δ₂ΔC164, which lacks IF2 and part of IF1, forms dimers and small oligomers (Fig. 4B). We have shown by atomic force microscopy that upon ATP binding, δ₂ free in solution forms discrete blob-shaped structures containing 2–3 δ₂ molecules (24) rather than long bundles in the absence of nsDNA. It is likely that DNA-independent bundle formation (Fig. 2A) and δ₂ blobs on nsDNA (Fig. 2C) are two mutually exclusive states, and δ₂ bundles, if formed in vivo, might be for storage purposes, as proposed for RecA (41), rather than for plasmid segregation. Indeed, many of these large or small ParA proteins might form bundles (e.g. F-SopA), but such assembly is inhibited in the presence of nsDNA (42).

In the filament-pulling model, pB171-ParA₂ (Sm) binds and polymerizes on nsDNA, with the filaments pulling plasmids apart by depolymerization (Fig. 2B) (26, 27). If pB171-ParA₂ (Sm) interacts with and polymerizes on nsDNA, as depicted in Fig. 2B, the N- and C-terminal domains should play a crucial role in plasmid segregation. However, deletion of the N-terminal (δ₂ΔN20) or C-terminal (δ₂ΔC255) domain did not abrogate accurate plasmid segregation (Table 2). Alternatively, once the tip of U-shaped ParA₂ (Sm) binds nsDNA, ParA₂ (Sm) may polymerize through unknown regions. To accommodate our results, we have to assume that ParA₂ polymerization onto nsDNA leads to double filaments, because the dimer of dimer interface maps at the bottom of the U-shaped protein (Fig. 7A, center) and is free for interacting with a preformed filament. Protein δ₂ fails to form nucleoprotein filaments (15, 24). However, pseudofilaments on nsDNA were observed in the presence of δ₂ and a very large excess of ω₂ (>300-fold over its K_D(app)) (15). Dissection of these “(ω₂·δ₂)_n·nsDNA filaments” revealed that they were composed of discrete δ₂ blobs stabilized by the nonspecific interaction of ω₂ with nsDNA (34).

Our data support non-filament-based models and are compatible with the diffusion-ratchet or DNA relay models. Because these models diverged at later stages (11, 29) and we are analyzing the early ones, they are considered as a single model (Fig. 2C). From previous data and data presented here, we conclude that (i) the unstructured N-terminal region of ω₂ (denoted in red in Fig. 2C), which is dispensable for binding to parS DNA, is involved in the interaction with δ₂ (25, 33); (ii) δ₂, upon ATP binding, binds to nsDNA and forms discrete blob-shaped structures containing 5–6 δ₂/blob rather than filaments on the nsDNA in vitro (14, 24), and it is homogeneously distributed on the nucleoid in vivo (34); (iii) the interaction of δ₂ bound to nsDNA with ω₂ bound to parS DNA promotes a structural transition in both proteins (see Figs. 5C and 7A); (iv) this protein-protein complex captures and moves the plasmids from any cytosolic position toward the nucleoid (Fig. 2C) (14, 24, 34); and (v) at equimolar ratios, ω₂ bound to parS enhances δ₂-mediated ATP hydrolysis and directs the Brownian ratchet of the diffusing partition complexes (Fig. 2C).

Author Contributions

A. V. and J. C. A. designed the study; A. V. performed the experiments; A. V. and J. C. A. analyzed the data; J. C. A. wrote the manuscript. Both authors reviewed the results and approved the final version of the manuscript.