Uncoupling of nucleotide hydrolysis and polymerization in the ParA protein superfamily disrupts DNA segregation dynamics

Abstract

DNA segregation in bacteria is mediated most frequently by proteins of the ParA superfamily that transport DNA molecules attached via the segrosome nucleoprotein complex. Segregation is governed by a cycle of ATP-induced polymerization and subsequent depolymerization of the ParA factor. Here, we establish that hyperactive ATPase variants of the ParA homolog ParF display altered segrosome dynamics that block accurate DNA segregation. An arginine finger-like motif in the ParG centromere-binding factor augments ParF ATPase activity but is ineffective in stimulating nucleotide hydrolysis by the hyperactive proteins. Moreover, whereas polymerization of wild-type ParF is accelerated by ATP and inhibited by ADP, filamentation of the mutated proteins is blocked indiscriminately by nucleotides. The mutations affect a triplet of conserved residues that are situated neither in canonical nucleotide binding and hydrolysis motifs in the ParF tertiary structure nor at interfaces implicated in ParF polymerization. Instead the residues are involved in shaping the contours of the binding pocket so that nucleotide binding locks the mutant proteins into a configuration that is refractory to polymerization. Thus, the architecture of the pocket not only is crucial for optimal ATPase kinetics but also plays a key role in the polymerization dynamics of ParA proteins that drive DNA segregation ubiquitously in procaryotes.

FIGURE 1.

Mutations in a triplet of conserved residues induce hyperactive ATPase profiles in ParF. A, linear representation of ParF with Walker A, A′, and B motifs highlighted in red. Mutation of Gly-11 and Lys-K15 residues in the A box ablates ATP hydrolysis (15). Positions of Pro-104, Arg-169, and Gly-179 residues and their conservation in selected ParF relatives (30) are shown. B, ATPase assays of wild-type ParF and derivatives bearing P104A, R169A, or G179A mutations. ATP hydrolysis is plotted with proteins (4 μm) at 0–500 μm ATP concentrations. C, ATPase assays of wild-type ParF and derivatives bearing P104A, R169A, or G179A mutations. ATP hydrolysis is plotted as a function of protein concentration with ATP at 5 μm. The data shown in A and B are typical results of experiments performed at least in duplicate.

FIGURE 2.

Locations of residues Pro-104, Arg-169, and Gly-179 in the ParF crystal structure. A, ParF sandwich dimer bound to the nonhydrolyzable ATP analog AMPPCP (29). Monomers are shown in red and blue with locations of Pro-104, Arg-169, and Gly-179 highlighted on the former. B, close-up view of the Pro-104–Arg-169–Gly-179 triplet in monomeric ParF bound to ADP. C–E, wireframe representations in monomeric ParF bound to ADP of Pro-104 and Arg-169 with locations of the nucleotide and P-loop included, of Arg-169 and nearby residues Asp-187 and Glu-193, and of Gly-179 and Trp-47. The blue mesh represents the composite omit 2Fo-Fc map for the structure, contoured at 1σ.

FIGURE 3.

Fluorescence anisotropy studies of nucleotide binding by ParF and mutant proteins. A and B, anisotropy changes when MANT-ATP (0.9 μm) (A) or MANT-ATPγS (0.9 μm) (B) was titrated with increasing concentrations of wild-type ParF and ParF^H proteins. C, intrinsic tryptophan fluorescence measurements of increasing concentrations of ParF without exogenous nucleotide or in the presence of MANT-ATP or MANT-ATPγS (0.9 μm each). D, intrinsic tryptophan fluorescence measurements of increasing concentrations of ParF-P104A without exogenous nucleotide or in the presence of MANT-ATP or MANT-ATPγS (0.9 μm each). The average fluorescence anisotropy values for 10 measurements for each point are shown.

FIGURE 4.

ATPase stimulation and segregation defects of hyperactive ParF ATPase mutants. A, partition assays of the segregation probe vector pFH450 (32) and the vector possessing either the wild-type parFGH cassette (pFH547) (30) or the same cassette bearing mutations that produce ParF-P104A and ParF-R169A. B, ATPase activities of ParF^H mutants are not stimulated by ParG. Levels of ATP hydrolysis driven by ParF and mutated proteins as a function of ParG concentration are shown. ParF proteins were used at 0.5 μm. The data are expressed as fold stimulation of ATPase activity compared with basal activity without added ParG. The results in both panels are averages of at least three replicates.

FIGURE 5.

Polymerization kinetics of wild-type ParF and ParF^H. A, sedimentation assays in which proteins (4–8 μm) were incubated in the absence (−) or presence (2 mm) of nucleotides for 10 min at 30 °C, and the reactions were then centrifuged. In all, 100 and 33%, respectively, of the pellet (P) and supernatant (S) fractions were resolved on a 12% SDS gel and stained with Coomassie Blue. The percentages of ParF proteins detected in the pellet fractions are shown. B, polymerization of wild-type and mutated ParF proteins monitored by dynamic light scattering. Proteins (2 μm) were preincubated at 30 °C for 5 min, at which time ATP or ADP (500 μm) and MgCl₂ (5 mm) were added (arrows). The reactions were followed for a further 55 min. The data in both panels are representative examples of experiments performed at least in duplicate with standard deviations ± 10%.

FIGURE 6.

Nucleotide binding elicits different polymerization responses in wild-type ParF and ParF^H. Steps 1 and 2, ATP (red oval) induces ParF dimerization (step 1 and Ref. 29) that primes the protein for polymerization (step 2 and Ref. 15). Step 3, ParF^H mutations mimic the effect of ATP binding (step 3), so that the protein is more prone to autopolymerization (step 4). Steps 5 and 6, in contrast, nucleotide binding by ParF^H causes a conformational change (step 5) that locks the protein into a configuration that is refractory to polymerization (step 6).