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A CTP-dependent gating mechanism enables ParB spreading on DNA

  1. Adam SB Jalal
  2. Ngat T Tran
  3. Clare EM Stevenson
  4. Afroze Chimthanawala
  5. Anjana Badrinarayanan
  6. David M Lawson
  7. Tung BK Le  Is a corresponding author
  1. Department of Molecular Microbiology, John Innes Centre, United Kingdom
  2. Department of Biochemistry and Metabolism, John Innes Centre, United Kingdom
  3. National Centre for Biological Sciences, Tata Institute of Fundamental Research, India
  4. SASTRA University, India
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Cite this article as: eLife 2021;10:e69676 doi: 10.7554/eLife.69676

Abstract

Proper chromosome segregation is essential in all living organisms. The ParA-ParB-parS system is widely employed for chromosome segregation in bacteria. Previously, we showed that Caulobacter crescentus ParB requires cytidine triphosphate to escape the nucleation site parS and spread by sliding to the neighboring DNA (Jalal et al., 2020). Here, we provide the structural basis for this transition from nucleation to spreading by solving co-crystal structures of a C-terminal domain truncated C. crescentus ParB with parS and with a CTP analog. Nucleating ParB is an open clamp, in which parS is captured at the DNA-binding domain (the DNA-gate). Upon binding CTP, the N-terminal domain (NTD) self-dimerizes to close the NTD-gate of the clamp. The DNA-gate also closes, thus driving parS into a compartment between the DNA-gate and the C-terminal domain. CTP hydrolysis and/or the release of hydrolytic products are likely associated with reopening of the gates to release DNA and recycle ParB. Overall, we suggest a CTP-operated gating mechanism that regulates ParB nucleation, spreading, and recycling.

Introduction

Proper chromosome segregation is essential in all domains of life. In most bacterial species, faithful chromosome segregation is mediated by the tripartite ParA-ParB-parS system (Donczew et al., 2016; Fogel and Waldor, 2006; Harms et al., 2013; Ireton et al., 1994; Jakimowicz et al., 2002; Jalal and Le, 2020a; Kawalek et al., 2018; Lin and Grossman, 1998; Livny et al., 2007; Mohl et al., 2001; Tran et al., 2018). ParB, a CTPase and DNA-binding protein, nucleates on parS before spreading to adjacent non-specific DNA to form a higher-order nucleoprotein complex (Breier and Grossman, 2007; Broedersz et al., 2014; Graham et al., 2014; Jalal and Le, 2020a; Murray et al., 2006; Rodionov et al., 1999; Sanchez et al., 2015; Taylor et al., 2015). The ParB-DNA nucleoprotein complex stimulates the ATPase activity of ParA, driving the movement of the parS locus (and subsequently, the whole chromosome) to the opposite pole of the cell (Hwang et al., 2013; Leonard et al., 2005; Lim et al., 2014; Taylor et al., 2021; Vecchiarelli et al., 2014; Vecchiarelli et al., 2012). ParB spreads by sliding along the DNA, in a manner that depends on the binding of a co-factor, cytidine triphosphate (CTP) (Balaguer F de et al., 2021; Jalal et al., 2020c; Osorio-Valeriano et al., 2019; Soh et al., 2019). A co-crystal structure of a C-terminal domain truncated Bacillus subtilis ParB (ParB∆CTD) together with CDP showed the nucleotide to be sandwiched between adjacent subunits, thus promoting their dimerization (Soh et al., 2019). A similar arrangement was seen in the co-crystal structure of an N-terminal domain (NTD) truncated version of the Myxococcus xanthus ParB homolog, PadC, bound to CTP (Osorio-Valeriano et al., 2019). Self-dimerization at the NTD of B. subtilis ParB creates a clamp-like molecule that enables DNA entrapment (Soh et al., 2019). Biochemical studies with M. xanthus and C. crescentus ParBs showed that CTP facilitates the dissociation of ParB from parS, thereby switching ParB from a nucleating mode to a sliding mode (Jalal et al., 2020c; Osorio-Valeriano et al., 2019). ParB can hydrolyze CTP to CDP and inorganic phosphate (Jalal et al., 2020c; Osorio-Valeriano et al., 2019; Soh et al., 2019); however, hydrolysis is not required for spreading since ParB in complex with a non-hydrolyzable CTP analog (CTPɣS) can still self-load and slide on DNA (Jalal et al., 2020c; Soh et al., 2019). Furthermore, M. xanthus PadC does not possess noticeable CTPase activity (Osorio-Valeriano et al., 2019). As such, the role of CTP hydrolysis in bacterial chromosome segregation is not yet clear.

Here, we solve co-crystal structures of a C-terminal domain truncated C. crescentus ParB in complex with either parS or CTPɣS to better understand the roles of CTP binding and hydrolysis. Consistent with the previous report (Soh et al., 2019), the NTDs of C. crescentus ParB also self-dimerize upon binding to nucleotides, thus closing a molecular gate at this domain (the NTD-gate). Furthermore, the two opposite DNA-binding domains (DBD) move closer together to close a second molecular gate (the DNA-gate). We provide evidence that the CTP-induced closure of the DNA-gate drives parS DNA from the DBD into a 20-amino-acid long compartment between the DNA-gate and the C-terminal domain, thus explaining how CTP binding enables ParB to escape the high-affinity parS site to spread while still entrapping DNA. Lastly, we identify and characterize a ParB ‘clamp-locked’ mutant that is defective in CTP hydrolysis but otherwise competent in gate closing, suggesting a possible role for CTP hydrolysis/release of hydrolytic products in the reopening ParB gates and in recycling ParB. Collectively, we suggest a CTP-operated gating mechanism that might regulate ParB nucleation, spreading, and recycling.

Results

Co-crystal structure of a C. crescentus ParB∆CTD-parS complex reveals an open conformation at the NTD

We sought to solve a co-crystal structure of C. crescentus ParB nucleating at parS. After screening several constructs with different lengths of ParB and parS, we obtained crystals of a 50 amino acid C-terminally truncated ParB in complex with a 22 bp parS DNA (Figure 1). This protein variant lacks the C-terminal domain (CTD) responsible for ParB dimerization (Figure 1A; Figge et al., 2003). Diffraction data for the ParB∆CTD-parS co-crystal were collected to 2.9 Å resolution, and the structure was solved by molecular replacement (see Materials and methods). The asymmetric unit contains four copies of ParB∆CTD and two copies of the parS DNA (Figure 1—figure supplement 1A,B).

Figure 1 with 2 supplements see all
Co-crystal structure of a C. crescentus ParB∆CTD-parS complex reveals an open conformation at the N-terminal domain (NTD).

(A) The domain architecture of C. crescentus ParB: the NTD (dark green), the central DNA-binding domain (DBD, dark green), the C-terminal domain (CTD, faded green), and a linker that connects the DBD and the CTD together. The ParB∆CTD variant that was used for crystallization lacks the CTD (faded green). (B, left panel) Co-crystal structure of two C. crescentus ParB∆CTD monomers (dark green and gray) bound to a 22 bp parS DNA. The nucleotide sequence of the 22 bp parS is shown below the co-crystal structure, the core parS sequence is highlighted in bold, and each parS half-site is denoted by an arrow. The position of residue L224 is also indicated. (Right panel) The structure of a ParB∆CTD subunit bound to a parS half site with key features highlighted. (C) Superimposition of C. crescentus ParB∆CTD subunits shows two different orientations of the NTD. The arrow above each subunit shows the direction each NTD is projecting towards. (D) A top-down view of the superimposition of ParB∆CTD subunits shows their NTDs orienting ~80° apart from each other.

Each ParB∆CTD subunit consists of an NTD (helices α1–α4 and sheets β1–β4) and a DBD (helices α5–α10) (Figure 1B). Each ParB∆CTD binds to a half parS site, but there is no protein-protein contact between the two adjacent subunits (Figure 1B). We previously reported a 2.4 Å co-crystal structure of the DBD of C. crescentus ParB bound to parS (Jalal et al., 2020b) and elucidated the molecular basis for specific parS recognition, hence we focus on the conformation of the NTD here instead. We observed that helices α3 and α4 are packed towards the DBD and are connected to the rest of the NTD via an α3–β4 loop (Figure 1B,C). While the DBD and helices α3–α4 are near identical between the two ParB∆CTD subunits (root-mean-square deviation [RMSD] = 0.19 Å, Figure 1C), the rest of the NTD, from α1 to β4, adopts notably different conformations in the two subunits (Figure 1C,D). Specifically, NTDs (α1–β4) from the two ParB∆CTD subunits are related by a rotation of approximately 80o due to changes in a flexible loop in between α3 and β4 (Figure 1D). Furthermore, by superimposing the C. crescentus ParB∆CTD-parS structure onto that of Helicobacter pylori (Chen et al., 2015), we observed that the NTDs of ParB from both species can adopt multiple alternative orientations (Figure 1—figure supplement 2). Taken together, these observations suggest that the ability of the NTD to adopt multiple open conformations is likely a general feature of nucleating ParB.

Co-crystal structure of a C. crescentus ParB∆CTD-CTPɣS complex reveals a closed conformation at the NTD

Next, to gain insight into the spreading state of ParB, we solved a 2.7 Å resolution structure of C. crescentus ParB∆CTD in complex with CTPɣS (see Materials and methods). At this resolution, it was not possible to assign the position of the ligand's sulfur atom. Indeed, the placement of the sulfur atom relative to the terminal phosphorus atom may vary from one ligand to the next in the crystal, leading to an averaging of the electron density. Hence, we modeled CTP, instead of CTPɣS, into the electron density (Figure 2 and Figure 2—figure supplement 1). The asymmetric unit contains two copies of ParB∆CTD, each with a CTPɣS molecule and a coordinated Mg2+ ion bound at the NTD (Figure 2A). In contrast to the open conformation of the ParB∆CTD-parS structure, nucleotide-bound NTDs from opposite subunits self-dimerize (with an interface area of 2111 Å2, as determined by PISA; Krissinel, 2015), thus adopting a closed conformation (Figure 2A). Multiple CTPɣS-contacting residues also directly contribute to the NTD self-dimerization interface (summarized in Figure 2—figure supplement 2), indicating a coupling between nucleotide binding and self-dimerization. Furthermore, the C. crescentus ParB∆CTD-CTPɣS structure is similar to that of the CDP-bound B. subtilis ParB∆CTD (RMSD = 1.48 Å) and the CTP-bound M. xanthus PadC∆NTD (RMSD = 2.23 Å) (Figure 2—figure supplement 3A), suggesting that the closed conformation at the NTD is structurally conserved in nucleotide-bound ParB/ParB-like proteins.

Figure 2 with 3 supplements see all
Co-crystal structure of a C. crescentus ParB∆CTD-CTPɣS complex reveals a closed conformation at the N-terminal domain (NTD).

(A, left panel) The front view of the co-crystal structure of C. crescentus ParB∆CTD (dark green and gray) bound to a non-hydrolyzable analog CTPɣS (orange) and Mg2+ ions (dark green and gray spheres). (Right panel) The top view of the C. crescentus ParB∆CTD-CTPɣS co-crystal structure. Note that helix α10 is not resolved in this structure due to a poor electron density in this region. (B) The nucleotide-binding pocket of C. crescentus ParB showing amino acid residues that contact the CTPɣS molecule and the coordinated Mg2+ ion. (C) Protein-ligand interaction map of CTPɣS bound to C. crescentus ParB∆CTD. Hydrogen bonds are shown as dashed green lines and hydrophobic interactions as red semi-circles. Nitrogen, oxygen, phosphate, and magnesium atoms are shown as blue, red, purple, and green filled circles, respectively.

Each CTPɣS molecule is sandwiched between helices α1, α2, α3 from one subunit and helix α3′ from the opposite subunit (Figure 2B). Ten amino acids form hydrogen-bonding contacts with three phosphate groups of CTPɣS, either directly or via the coordinated Mg2+ ion (Figure 2C). These phosphate-contacting residues are referred to as P-motifs 1–3, respectively (P for phosphate motif, Figure 2C). Four amino acids at helix α1 and the α1–β2 intervening loop provide hydrogen-bonding interactions to the cytosine ring, hence are termed the C-motif (C for cytosine motif, Figure 2C). Lastly, six additional residues contact the ribose moiety and/or the pyrimidine moiety via hydrophobic interactions (Figure 2C). Nucleotide-contacting residues in C. crescentus ParB and their corresponding amino acids in ParB/ParB-like homologs are summarized in Figure 2—figure supplement 2 and Figure 2—figure supplement 3B. The C-motif forms a snug fit to the pyrimidine moiety, thus is incompatible with larger purine moieties such as those from ATP or GTP. Hydrogen-bonding contacts from the G79 main chain and the S74 side chain to the amino group at position 4 of the cytosine moiety further distinguish CTP from UTP (Figure 2C). Taken all together, our structural data are consistent with the known specificity of C. crescentus ParB for CTP (Jalal et al., 2020c).

Conformational changes between the nucleating and the spreading state of C. crescentus ParB

A direct comparison of the C. crescentus ParB∆CTD-parS structure to the ParB∆CTD-CTPɣS structure further revealed the conformational changes upon nucleotide binding. In the nucleating state, as represented by the ParB∆CTD-parS structure, helices α3 and α4 from each subunit bundle together (32o angle between α3 and α4, Figure 3). However, in the spreading state, as represented by the ParB∆CTD-CTPɣS structure, α3 swings outwards by 101o to pack itself with α4′ from the opposing subunit (Figure 3). Nucleotide binding most likely facilitates this ‘swinging-out’ conformation since both α3 and the α3–α4 loop, that is, P-motif 3 make numerous contacts with the bound CTPɣS and the coordinated Mg2+ ion (Figure 2C). The reciprocal exchange of helices ensures that the packing in the α3–α4 protein core remains intact, while likely driving the conformational changes for the rest of the NTD as well as the DBD (Figure 4A). Indeed, residues 44–121 at the NTD rotate wholesale by 94o to dimerize with their counterpart from the opposing subunit (Figure 4A and Figure 4—figure supplement 1A). Also, residues 161–221 at the DBD rotate upward by 26o in a near rigid-body movement (Figure 4A and Figure 4—figure supplement 1A). As a result, the opposite DBDs are closer together in the spreading state (inter-domain distance = ~ 27 Å) than in the nucleating state (inter-domain distance = ~ 36 Å) (Figure 4—figure supplement 1B). By overlaying the CTPɣS-bound structure onto the parS DNA complex, it is clear that the DBDs in the spreading state clash severely with DNA, hence are no longer compatible with parS DNA binding (Figure 4B). Our structural data are therefore consistent with the previous finding that CTP decreases C. crescentus ParB nucleation on parS or liberates pre-bound ParB from parS site (Jalal et al., 2020c). Overall, we suggest that CTP binding stabilizes a conformation that is incompatible with DNA-binding and that this change might facilitate ParB escape from the high-affinity nucleation parS site.

Conformational changes between the nucleating and the spreading states of C. crescentus ParB.

Structures of C. crescentus ParB∆CTD in complex with parS (left panel) and with CTPɣS (right panel), with the pairs of helices (α3–α4, and α3′–α4′ for the opposite subunit) shown in light blue and dark blue, respectively. Below each structure, only the α3–α4, α3′–α4′ pairs, and the angles between these helices are shown.

Figure 4 with 1 supplement see all
The structure of a nucleotide-bound C. crescentus ParB∆CTD is incompatible with specific parS binding at the DNA-binding domain (DBD).

(A) Structural changes between C. crescentus ParB∆CTD-parS and ParB∆CTD-CTPɣS structures. Helices α3 and α4 are shown in light blue. The arrows next to the N-terminal domain (NTD) (residues 44–121) and the DBD (residues 161–221) show the direction that these domains rotate towards in the nucleotide-bound state. (B) Superimposing the C. crescentus ParB∆CTD-CTPɣS structure onto parS DNA shows DNA-recognition helices (α6 and α6′, magenta) positioning away from the two consecutive major grooves of parS, and helices α8–α9 and α8′–α9′ at the DBD (dashed box) clashing with parS DNA.

C. crescentus ParB entraps parS DNA in a compartment between the DBD and the CTD in a CTP-dependent manner

To verify the CTP-dependent closed conformation of ParB, we performed site-specific crosslinking of purified proteins using a sulfhydryl-to-sulfhydryl crosslinker bismaleimidoethane (BMOE) (Soh et al., 2019). Residues Q35, L224, and I304 at the NTD, DBD, and CTD, respectively (Figure 5A), were substituted individually to cysteine on an otherwise cysteine-less ParB (C297S) background (Jalal et al., 2020c), to create ParB variants where symmetry-related cysteines become covalently linked if they are within 8 Å of each other (Figure 5B). We observed that the crosslinking of both ParB (Q35C) and ParB (L224C) was enhanced ~2.5–3-fold in the presence of parS DNA and CTP (Figure 5B), consistent with CTP favoring a conformation when the NTD and the DBD are close together. In contrast, ParB (I304C) crosslinked independently of CTP or parS (Figure 5B), supporting the known role of the CTD as a primary dimerization domain (Figge et al., 2003; Fisher et al., 2017).

Figure 5 with 4 supplements see all
C. crescentus ParB entraps parS DNA in a compartment between the DNA-binding domain (DBD) and the C-terminal domain (CTD) in a cytidine triphosphate (CTP)-dependent manner.

(A) A schematic diagram of C. crescentus ParB showing the position of Q35 (at the N-terminal domain [NTD]), L224 (at the DBD), and I304 (at the CTD) that were substituted either individually or in combinations for cysteine. (B) Denaturing polyacrylamide gel analysis of bismaleimidoethane (BMOE) crosslinking products of 8 µM single-cysteine ParB (Q35C/L224C/I304C) variant ±0.5 µM 22 bp parS DNA ±1 mM CTP. X indicates a crosslinked form of ParB. Quantification of the crosslinked (X) fraction is shown below each representative gel image. Error bars represent SD from three replicates. (C, left panel) Denaturing polyacrylamide gel analysis of BMOE crosslinking products of 8 µM dual-cysteine ParB (Q35C I304C) variant ±0.5 µM DNA ±1 mM CTP. Different DNA were employed in crosslinking reactions: a linear 22 bp parS DNA (22 bp parS lin), a circular 3 kb parS plasmid (3 kb parS cir), and a circular 3 kb scrambled parS plasmid (3 kb nonS cir). The high molecular weight (HMW) smear near the top of the polyacrylamide gel is marked with a solid line and an asterisk (lane 7). When the crosslinking reaction was post-treated with a non-specific DNA nuclease, Benzonase, the HMW smear was no longer observed (dashed line and asterisk, lane 8). The polyacrylamide gel was also stained with a DNA dye, Sybr Green (SYBR), and only the top section of the gel is shown. Small 22 bp parS DNA duplex migrated out of the gel, thus was not observed near the top of the Sybr-stained gel. A schematic diagram of a dual-cysteine C. crescentus ParB dimer is also shown. (Right panel) Agarose gel analysis of BMOE crosslinking products. A subset of crosslinking reactions (lanes 6, 7, and 9–12) were loaded and resolved on 1% agarose gel. The gel was subsequently stained with Sybr Green for DNA. Shifted gel bands are marked with a solid line and an asterisk. (D) Same as panel (C) but another dual-cysteine variant, ParB (L224C I304C) was employed instead.

Figure 5—source data 1

Original files, annotation of the full raw gels, and data used to generate Figure 5.

https://cdn.elifesciences.org/articles/69676/elife-69676-fig5-data1-v1.zip

Previously, it was shown that B. subtilis ParB-CTP forms a protein clamp that entraps DNA (Soh et al., 2019); however, the location of DNA within the clamp is not yet clear. To locate such DNA-entrapping compartment, we employed a double crosslinking assay while taking advantage of the availability of crosslinkable cysteine residues in all three domains of C. crescentus ParB (Figure 5A). A C. crescentus ParB variant with crosslinkable NTD and CTD interfaces (Q35C I304C) was first constructed and purified (Figure 5C). ParB (Q35C I304C) could form high molecular weight (HMW) species near the top of the polyacrylamide gel in the presence of CTP, a 3 kb parS plasmid, and the crosslinker BMOE (lane 7, Figure 5C, left panel). The HMW smear on the polyacrylamide gel contained both protein and DNA as apparent from a dual staining with Coomassie and Sybr Green (Figure 5C, left panel). Slowly migrating DNA-stained bands were also observed when resolved on an agarose gel (Figure 5C, right panel). The HMW smear most likely contained DNA-protein catenates between a circular parS plasmid and a denatured but otherwise circularly crosslinked ParB (Q35C I304C) polypeptide. Indeed, a post-crosslinking treatment with Benzonase, a non-specific DNA nuclease (lane 8, Figure 5C, left panel) or the use of a linearized parS plasmid (lane 2 vs. lane 4, Figure 5—figure supplement 1) eliminated the HMW smear, presumably by unlinking the DNA-protein catenates. Lastly, the HMW smear was not observed when a plasmid containing a scrambled parS site was used (lane 10, Figure 5C, left panel) or when CTP was omitted from the crosslinking reaction (lane 6, Figure 5C, left panel), indicating that the DNA entrapment is dependent on parS and CTP. Collectively, these experiments demonstrate that as with the B. subtilis ParB homolog, C. crescentus ParB is also a CTP-dependent molecular clamp that can entrap parS DNA in between the NTD and the CTD.

Employing the same strategy, we further narrowed down the DNA-entrapping compartment by constructing a ParB (L224C I304C) variant in which both the DBD and the CTD are crosslinkable (Figure 5D). We found that crosslinked ParB (L224C I304C) also entrapped circular plasmid efficiently in a parS- and CTP-dependent manner, as judged by the appearance of the HMW smear near the top of the gel (lane 7, Figure 5D, left panel). By contrast, ParB (Q35C L224C) that has both the NTD and the DBD crosslinkable was unable to entrap DNA in any tested condition (Figure 5—figure supplement 2). We therefore hypothesized that ParB clamps entrap DNA within a compartment created by a 20-amino-acid linker in between the DBD and the CTD. To investigate further, we constructed a ParB (L224C I304C)-TEV variant, in which a TEV protease cleavage site was inserted within the DBD-CTD linker (Figure 5—figure supplement 3A). Again, ParB (L224C I304C)-TEV entrapped a circular parS plasmid efficiently in the presence of CTP (the HMW smear on lane 7, Figure 5—figure supplement 3A). However, a post-crosslinking treatment with TEV protease eliminated such HMW smear, presumably by creating a break in the polypeptide through which a circular plasmid could escape (lane 8, Figure 5—figure supplement 3A). We also extracted crosslinked ParB (L224C I304C) from gel slices that encompassed the HMW smear and electrophoresed the eluted proteins again on a denaturing gel to find a single band that migrated similarly to a double-crosslinked protein (lane 9, Figure 5—figure supplement 2B). Therefore, our results suggest that a ParB dimer, rather than ParB oligomers, is the major species that entraps DNA. Taken together, we suggest that C. crescentus ParB dimer functions as a molecular clamp that entraps parS-containing DNA within a DBD-CTD compartment upon CTP binding. This is also consistent with experiments that showed a premature and irreversible closing of ParB clamps, achieved either by an extended preincubation with CTPɣS (Jalal et al., 2020c and Figure 5—figure supplement 4B) or by pre-crosslinking a closed clamp form of ParB (Figure 5—figure supplement 4C), prevented nucleation at parS and DNA entrapment.

C. crescentus ParB (E102A) is a clamp-locked mutant that is defective in clamp reopening

Next, we investigated the potential role(s) of CTP hydrolysis. Hydrolysis is unlikely to be required for DNA entrapment and translocation since ParB in complex with CTPɣS can still self-load and slide on DNA (Jalal et al., 2020c; Soh et al., 2019). M. xanthus ParB (N172A) and B. subtilis ParB (N112S) mutants, which bind but cannot hydrolyze CTP, failed to form higher-order protein-DNA complexes inside the cells (Osorio-Valeriano et al., 2019; Soh et al., 2019). However, these ParB variants are already impaired in NTD self-dimerization (Soh et al., 2019), hence the mechanistic role of CTP hydrolysis is still unclear. We postulated that creation of a ParB variant defective in CTP hydrolysis but otherwise competent in NTD self-dimerization would enable us to investigate the possible role of CTP hydrolysis. To this end, we performed alanine scanning mutagenesis on the CTP-binding pocket of C. crescentus ParB (Figure 2C). Eleven purified ParB variants were assayed for CTP binding by a membrane-spotting assay (DRaCALA) (Figure 6A), and for CTP hydrolysis by measuring the release rate of inorganic phosphate (Figure 6B). Moreover, their propensity for NTD self-dimerization was also analyzed by crosslinking with BMOE (Figure 6C and Figure 6—figure supplement 1). Lastly, their ability to nucleate, slide, and entrap a closed parS DNA substrate was investigated by a biolayer interferometry (BLI) assay (Figure 6D and Figure 6—figure supplement 2A). Immobilizing a dual biotin-labeled DNA on a streptavidin-coated BLI surface created a closed DNA substrate that can be entrapped by ParB-CTP clamps (Figure 5—figure supplement 4A; Jalal et al., 2020c). The BLI assay monitors wavelength shifts resulting from changes in the optical thickness of the probe surface during the association/dissociation of ParB with a closed DNA substrate in real time (Figure 6—figure supplement 2).

Figure 6 with 2 supplements see all
Alanine scanning mutagenesis of the C. crescentus ParB cytidine triphosphate (CTP)-binding pocket reveals several classes of clamp mutants.

Eleven residues at C-motif and P-motifs 1–3 were individually substituted for alanine or glycine. (A) Membrane-spotting assay of ParB variants. CTP binding was monitored by membrane-spotting assay using radiolabeled CTP α-P32. The bulls-eye staining indicates CTP binding due to a more rapid immobilization of protein-ligand complexes compared to free ligands. All reactions contained various concentration of purified ParB, 5 nM radiolabeled CTP α-P32, 30 µM unlabeled CTP, and 1.5 µM 22 bp parS DNA. The bound fractions were quantified, and error bars represent SD from three replicates. All the reactions were spotted on the same membrane, the radiograph was rearranged solely for presentation purposes. (B) Inorganic phosphate release assay of ParB variants. The CTPase rates were measured at increasing concentration of CTP. All reactions contained 1 µM purified ParB variant, 0.5 µM 22 bp parS DNA, and an increasing concentration of CTP. (C) Bismaleimidoethane (BMOE) crosslinking assay of ParB variants. A second set of alanine scanning ParB variants, which harbor an additional Q35C substitution at the N-terminal domain (NTD), were also constructed and subsequently used in BMOE crosslinking experiments. Purified ParB variants (8 µM) were preincubated with 0.5 µM 22 bp parS DNA and an increasing concentration of CTP for 5 min before BMOE was added. Crosslinking products were resolved on a 12% denaturing polyacrylamide gel and the crosslinked fractions were quantified (see also Figure 6—figure supplement 1 for representation images). Error bars represent SD from three replicates. (D) Biolayer interferometry (BLI) assay of ParB variants. BLI analysis of the interaction between a premix of 1 µM ParB variant ± an increasing concentration of CTP and a 170 bp closed parS DNA substrate. See also Figure 5—figure supplement 4A for a schematic diagram of the BLI setup and Figure 6—figure supplement 2 for representative BLI sensorgrams. BLI signal at the end of the association phase (± SD from three replicates) was plotted against CTP concentrations.

Figure 6—source data 1

Original files, annotation of the full raw gels, and data used to generate Figure 6.

https://cdn.elifesciences.org/articles/69676/elife-69676-fig6-data1-v1.zip

Overall, we identified several distinct classes of ParB mutants:

  1. Class I: ParB (R60A), (R103A), (R104A), (R139A), (N136A), (G79S), and (S74A) did not bind or bound radiolabeled CTP only weakly (Figure 6A), thus also showed weak to no CTP hydrolysis (Figure 6B) or clamp-closing activity (Figure 6C,D).

  2. Class II: ParB (Q58A) and (E135A) that are competent in CTP-binding (Figure 6A), but defective in CTP hydrolysis (Figure 6B) and in entrapping a closed parS DNA substrate (Figure 6D). We noted that ParB (Q58A) and ParB (E135A) already had an elevated crosslinking efficiency even in the absence of CTP (Figure 6C). This premature clamp closing might have resulted in a less than wild-type level of DNA entrapment (Figure 6D).

  3. Class III: ParB (E102A) did not hydrolyze CTP (Figure 6B) but nevertheless bound CTP efficiently (Figure 6A) to self-dimerize at the NTD and to entrap DNA to the same level as ParB (WT) at all CTP concentrations (Figure 6C,D).

Upon a closer inspection of the BLI sensorgrams (Figure 6—figure supplement 2B and Figure 7), we noted that the entrapped ParB (E102A) did not noticeably dissociate from a closed DNA substrate when the probe was returned to a buffer-only solution (dissociation phase, koff = 8.0 × 10–4 ± 1.9 × 10–4 s–1, Figure 6—figure supplement 2B and Figure 7). By contrast, entrapped ParB (WT) dissociated approximately 15-fold faster into buffer (koff = 1.2 × 10–2 ± 3.7 × 10–4 s–1). Further experiments showed that DNA entrapment by ParB (E102A), unlike ParB (WT), is more tolerant to high-salt solution (up to 1 M NaCl, Figure 7A). Nevertheless, ParB (E102A)-CTP could not accumulate on a BamHI-restricted open DNA substrate (Figure 7B,C; Jalal et al., 2020c), suggesting that ParB (E102A)-CTP, similar to ParB (WT), also form a closed clamp that runs off an open DNA end. Collectively, our results suggest that parS DNA and CTP induced a stably closed clamp conformation of ParB (E102A) in vitro.

The DNA-entrapped ParB (E102A)-CTP clamp is resistant to high-salt conditions.

(A) Biolayer interferometry (BLI) analysis of the interaction between a premix of 1 µM C. crescentus ParB (WT) or ParB (E102A) + 1 mM cytidine triphosphate (CTP) and 170 bp dual biotin-labeled parS DNA. For the dissociation phase, the probe was returned to a low-salt buffer that contains 100 mM NaCl (solid black or red lines) or to a high-salt buffer that contains 1 M NaCl (dashed black or red lines). The schematic diagram of the BLI probe shows a closed parS DNA substrate due to the interactions between a dual biotin-labeled DNA and the streptavidin (SA)-coated probe surface. (B) BLI analysis of the interaction between a premix of 1 µM C. crescentus ParB (WT) or ParB (E102A) + 1 mM CTP (solid lines) or –1 mM CTP (dashed lines) and 170 bp dual biotin-labeled parS DNA. (C) Same as panel (B) but immobilized DNA fragments have been restricted with BamHI before BLI analysis.

To investigate the function of ParB (E102A) in vivo, we expressed a FLAG-tagged version of parB (E102A) from a vanillate-inducible promoter (Pvan) in a C. crescentus strain where the native parB was under the control of a xylose-inducible promoter (Pxyl) (Figure 8A). Cells were depleted of the native ParB by adding glucose for 4 hr, subsequently vanillate was added for another hour before cells were fixed with formaldehyde for ChIP-seq. Consistent with the previous report (Tran et al., 2018), the ChIP-seq profile of FLAG-ParB (WT) showed an ~10 kb region of enrichment above background with clearly defined peaks that correspond to the positions of parS sites (Figure 8A). By contrast, the ChIP-seq profile of FLAG-ParB (E102A) is significantly reduced in height but has an extra peak over the parB coding sequence (Figure 8A, asterisk). The instability of FLAG-ParB (E102A) in its native C. crescentus host, and hence the reduced protein level (Figure 8—figure supplement 1A), might explain the overall lower height of its ChIP-seq profile (Figure 8). The reason for an extra peak over parB in the ChIP-seq profile of ParB (E102A) is still, however, unknown. We also noted that expressing ParB (E102A) could not rescue cells with depleted ParB (WT) (Figure 8—figure supplement 2). Again, due to the caveat of a lower ParB (E102A) protein level in C. crescentus (Figure 8—figure supplement 1A), we could not reliably link the in vitro properties of ParB (E102A) to its behaviors in the native host.

Figure 8 with 3 supplements see all
ParB (E102A) occupies a more extended DNA region surrounding parS sites than ParB (WT) in a heterologous host (E. coli) but not in the native host (C. crescentus).

(A) ChIP-seq showed the distribution of FLAG-tagged ParB (WT) (black) and FLAG-ParB (E102A) (red) on C. crescentus chromosome between +4025 kb and +4042 kb. Underlying genes and parS sites are also shown below ChIP-seq profiles. An asterisk (*) indicates an extra peak over the parB coding sequence in the profile of FLAG-ParB (E102A). ChIP-seq signals were reported as the number of reads per base pair per million mapped reads (RPBPM). (B) ChIP-seq showed the distribution of CFP-tagged ParB (WT) (black) and CFP-ParB (E102A) (red) on an E. coli chromosome between +2885 kb and +2915 kb. C. crescentus parS sites 3 and 4 were engineered onto the E. coli chromosome at the ygcE locus. CFP-tagged ParB (WT/E102A) was expressed from a leaky lactose promoter (Plac, no IPTG was added) on a medium-copy-number plasmid. Shaded boxes show areas with more enrichment in the ChIP-seq profile of CFP-ParB (E102A) compared to that of CFP-ParB (WT).

To overcome the caveat of protein instability, we instead investigated the spreading of ParB (WT) vs. ParB (E102A) from parS by analyzing the C. crescentus ParB/parS system in Escherichia coli. E. coli does not possess a ParA/ParB homolog nor a parS-like sequence, thus it serves as a suitable heterologous host. C. crescentus parS sites 3 and 4 were engineered onto the E. coli chromosome at the ygcE locus (Figure 8B). CFP-tagged ParB (WT/E102A) was expressed from a leaky lactose promoter (Plac, no IPTG was added) on a medium-copy-number plasmid. CFP-ParB (WT/E102A) was produced at the same level, as judged by an immunoblot (Figure 8—figure supplement 1B). We observed by ChIP-seq that CFP-ParB (WT) in an E. coli host spreads asymmetrically ~5 kb around parS sites. By contrast, the shape of the ParB (E102A) distribution was clearly different from that of ParB (WT); the profile was further expanded to both neighboring sides of parS (covering in total ~26 kb) at the expense of the enrichment at parS itself (Figure 8B). The more excessive spreading of ParB (E102A) might suggest that this variant, in the absence of CTP hydrolysis, persisted and perhaps slid further away from the loading site parS in E. coli. The reduced enrichment of ParB (E102A) at parS itself (Figure 8B) might be due to reduced cytoplasmic ParB (E102A) available to re-nucleate at parS and/or due to stably entrapped ParB (E102A) sterically hindering further nucleation events. We also noted that the ChIP-seq profile of CFP-ParB (E102A) in E. coli is highly asymmetrical, with more enrichment in the 2905–2911 kb region than the 2885–2899 kb region (shaded areas, Figure 8B). The asymmetrical spreading is possibly due to an impediment in one direction by roadblocks such as RNA polymerases or DNA-bound proteins, which have been shown previously to be able to interfere with ParB spreading (Balaguer F de et al., 2021; Breier and Grossman, 2007; Jalal et al., 2020c; Murray et al., 2006; Rodionov et al., 1999; Soh et al., 2019).

Lastly, we quantified the fluorescence intensity of CFP-ParB (WT/E102A) foci inside cells and found a higher CFP signal for CFP-ParB (E102A) when compared with CFP-ParB (WT) (Figure 8—figure supplement 3). The higher intensity of the localizations could be due to more DNA-bound ParB (E102A) molecules surrounding the parS locus, which is consistent with the ChIP-seq observation showing CFP-ParB (E102A) occupying a more extended genomic area in E. coli. Altogether, at least in the heterologous E. coli host, the ‘clamp-locked’ phenotype of ParB (E102A) implies a possible role of CTP hydrolysis and/or the release of hydrolytic products in reopening wild-type ParB clamp to release DNA and to recycle ParB.

Discussion

In this study, we provide structural insights into the nucleating and sliding states of C. crescentus ParB. Nucleating ParB is an open clamp in which parS DNA is held tightly (nM affinity) at the DBD (Tran et al., 2018). The NTDs of nucleating ParB can adopt multiple alternative conformations, and crucially there is no contact between opposing NTDs. We liken this conformation of the NTD to that of an open gate (NTD-gate), through which parS DNA might gain access to the DBD (Figure 9). In the sliding state, CTP promotes the self-dimerization of the NTDs, thus closing the NTD-gate (Figure 9). Opposing DBDs also move approximately 10 Å closer together, bringing about a conformation that is DNA incompatible. Again, we liken this conformation of the DBDs to that of a closed gate (DNA-gate) (Figure 9). Overall, the DNA-gate closure explains how CTP binding might switch ParB from a nucleating to a sliding state.

A model for C. crescentus ParB nucleating, sliding, and recycling cycle.

ParB (dark green) consists of three domains: an N-terminal CTP-binding domain (NTD), a central parS DNA-binding domain (DBD), a C-terminal dimerization domain (CTD), and a 20 amino acid linker that connects the DBD and the CTD together. Nucleating ParB is an open clamp, in which parS DNA is captured at the DBD (the DNA-gate). Upon binding CTP (orange), the NTD self-dimerizes to close the NTD-gate of the clamp. CTP-binding and the exchange of helices α4 and α4′ (blue) stabilize this closed conformation. The DBD also move closer together to close the DNA-gate, potentially driving parS DNA into a compartment between the DNA-gate and the C-terminal domain. In the nucleotide-bound state, the DBD and the DNA-recognition helices (α6 and α6′, magenta) are incompatible with DNA binding. CTP hydrolysis and/or the release of hydrolytic products (CDP and inorganic phosphate Pi) may reopen the gates to release DNA. Substitutions that affect key steps in the CTP biding/hydrolysis cycle are also indicated on the schematic diagram.

Our data suggest that the closure of the two gates drives parS DNA into a compartment in between the DBD and the CTD. Previously, (Soh et al., 2019) compared the B. subtilis ParB∆CTD-CDP co-crystal structure to that of a H. pylori ParB∆CTD-parS complex and proposed that DNA must be entrapped in the DBD-CTD compartment (Soh et al., 2019). Here, the available structures of nucleating and sliding ParB from the same bacterial species enabled us to introduce a crosslinkable cysteine (L224C) at the DBD, and subsequently provided a direct evidence that the DBD-CTD compartment is the DNA-entrapping compartment. The linker that connects the DBD and the CTD together is not conserved in amino acid sequence among chromosomal ParB orthologs (Figure 2—figure supplement 2); however, we noted that the linker is invariably ~20 amino acid in length and positively charged lysines are over-represented (Figure 2—figure supplement 2). The biological significance of the linker length and its lysines, if any, is currently unknown. However, it is worth noting that a human PCNA clamp was proposed to recognize DNA via lysine-rich patches lining the clamp channel, and that these lysine residues help PCNA to slide by tracking the DNA backbone (De March et al., 2017). Investigating whether lysine residues in the DBD-CTD linker of ParB have a similar role is an important subject for the future.

If not already bound on DNA, the closed ParB clamp presumably cannot self-load onto parS owing to its inaccessible DBD. In this study, we showed that parS DNA promotes the CTP-dependent NTD-gate closure (Figure 5B), thus is likely a built-in mechanism to ensure gate closure results in a productive DNA entrapment. However, the molecular basis for the parS-enhanced gate closure remains unclear due to the lack of a crystal structure of C. crescentus apo-ParB, despite our extensive efforts.

CTP functions as a molecular latch that stabilizes the closure of the NTD-gate of ParB. Here, we provide evidence that CTP hydrolysis might contribute to reopening the closed NTD-gate. A previous structure of a B. subtilis ParB∆CTD-CDP complex also has its NTD-gate closed (CTP was hydrolyzed to CDP during the crystallization) (Soh et al., 2019), hence it is likely that both CTP hydrolysis and the subsequent release of hydrolytic products are necessary to reopen the gates. However, ParB has a weak to negligible affinity for CDP, hence the CDP-bound ParB species might be short-lived in solution and might not play a significant biological role. Once the clamp is reopened, entrapped DNA might escape via the same route that it first enters. Other well-characterized DNA clamps, for example, type II topoisomerases open their CTD to release trapped DNA. However, the CTDs of ParB are stably dimerized independently of parS and CTP (Figure 5B), hence we speculate that the CTD of ParB is likely to be impassable to the entrapped DNA. The released ParB clamp might re-nucleate on parS and bind CTP to close the gate, hence restarting the nucleation and sliding cycle. Such a recycling mechanism might provide a biological advantage since a ParB clamp once closed could otherwise become stably trapped on DNA and thus eventually diffuse too far from the parS locus, as evidenced by the ChIP-seq profile of the E102A variant (expressed in E. coli) that is defective in CTP hydrolysis (Figure 8B). However, how CTP hydrolysis contributes to the assembly of the centromere in C. crescentus is still unclear due to the caveat that ParB (E102A) is unstable in the native host.

The CTP-bound structure of a M. xanthus ParB-like protein, PadC, was solved to a high resolution (1.7 Å); however, PadC does not possess noticeable CTPase activity (Osorio-Valeriano et al., 2019). A co-crystal structure of B. subtilis ParB with CDP was also solved to a high resolution (1.8 Å) but represents a post-hydrolysis state instead. Lastly, our CTPɣS-bound C. crescentus ParB crystals diffracted to 2.7 Å, thus preventing water molecules, including a potential catalytic water, from being assigned with confidence. Therefore, the mechanism of CTP hydrolysis by a ParB CTPase remains unresolved. Nevertheless, based on our alanine scanning experiment (Figure 8), we speculate that Q58 (P-motif 1) and E102 (P-motif 2) might be involved in the catalytic mechanism of C. crescentus ParB. Supporting this view, we noted that an equivalent Q37 in B. subtilis ParB does not contact the hydrolytic product CDP, and this residue is not conserved in the catalytic-dead M. xanthus PadC (F308, which does not contact CTP, occupies this position in PadC instead) (Figure 2—figure supplement 3). E102 is also not conserved in M. xanthus PadC (F348 occupies this equivalent position) (Figure 2—figure supplement 3). Given that ParB is the founding member of a new CTPase protein family (Jalal et al., 2020c; Osorio-Valeriano et al., 2019; Soh et al., 2019), further studies are needed to fully understand the molecular mechanism of CTP hydrolysis so that the knowledge gained might be generalized to other CTPases.

Recently, an F-plasmid ParB was shown to form biomolecular condensates in vivo that might bridge distal ParBF dimers together (Guilhas et al., 2020; Walter et al., 2020). If and how CTP binding/hydrolysis and the flexibility of the NTD contribute to this process is unclear and will be an important challenge for future studies. It is equally important to better understand the in vivo interaction between ParB and ParA now that CTP is in the picture. Recent in vitro work with ParABF showed that two protomers of a single ParBF dimer interact with a ParAF dimer in the absence of CTP (Taylor et al., 2021). However, two ParBF protomers from two distinct dimers interact with a ParAF dimer in the presence of CTP and parS (Taylor et al., 2021). Which mode of action is dominant in vivo for a chromosomal ParABS systems and whether interacting with ParA further facilitates CTP hydrolysis by ParB are still unknown. Future works will provide important insights to better understand the mechanism of ParA-directed DNA segregation.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Escherichia coli)See Supplementary file 1AThis paperSee Supplementary file 1A
Strain, strain background (Caulobacter crescentus)See Supplementary file 1AThis paperSee Supplementary file 1A
Recombinant DNA reagentSee Supplementary file 1BThis paperSee Supplementary file 1B
Sequence-based reagentSee Supplementary file 1BThis paperSee Supplementary file 1B
AntibodyAnti-GFP antibody (HRP) (Rabbit polyclonal)AbcamCat# ab190584Western blot (1:5000)
AntibodyAnti-GFP Sepharose beadsAbcamCat# ab69314For ChIP-seq experiments
AntibodyAnti-FLAG antibody (HRP) (Mouse monoclonal)MerckCat# A8592Western blot (1:5000)
AntibodyAnti-FLAG M2 affinity agarose beadsMerckCat# A2220For ChIP-seq experiments
Commercial assay or kitAmersham Protran supported western blotting membranes, nitrocelluloseGE HealthcareCat# GE10600016Pore size 0.45 μm, for DRaCALA assay
Commercial assay or kitEnzChek Phosphate Assay KitThermoFisherCat# E6646
Commercial assay or kitGibson Assembly Master MixNEBCat# E2611S
Commercial assay or kitGateway BP Clonase II enzyme mixThermoFisherCat# 11789020
Commercial assay or kitDip-and-Read Streptavidin biosensors SAX2Sartorius UKCat# 18-5019
Commercial assay or kitHisTrap High Performance columnGE HealthcareCat# GE17524801
Commercial assay or kitHisTrap Heparin High Performance columnGE HealthcareCat# GE17040601
Commercial assay or kitHiLoad 16/600 Superdex 200 pg columnGE HealthcareCat# GE28989335
Commercial assay or kit0.5 mL Zeba spin desalting columnsThermoFisherCat# 89,8827K Da molecular weight cutoff
Peptide, recombinant proteinBamHI-HFNEBCat# R3136S20,000 units/mL
Peptide, recombinant proteinHindIII-HFNEBCat# R3104S20,000 units/mL
Chemical compound, drugBenzonase nucleaseMerckCat# E1014250 units/µL
Chemical compound, drugCTPThermoFisherCat# R0451100 mM solution
Chemical compound, drugCTPγSJena BioscienceCustom synthesis (purity ≥96%)
Chemical compound, drugP32-α-CTPPerkin ElmerCat# BLU008H250UC3,000 Ci/mmol, 10 mCi/mL, 250 µCi
Chemical compound, drugBismaleimidoethane (BMOE)ThermoFisherCat# 22323Dissolved in DMSO
Chemical compound, drugAcTEV proteaseThermoFisherCat# 1257501510 units/µL
Software, algorithmBLItz ProMolecular DevicesCat# 50-0156Version 1.2
Software, algorithmAIMLESSEvans and Murshudov, 2013http://www.ccp4.ac.uk/Version 0.7.4
Software, algorithmBUCCANEERCowtan, 2006http://www.ccp4.ac.uk/Version 1.6.10
Software, algorithmCootEmsley and Cowtan, 2004http://www.ccp4.ac.uk/Version 0.9.5
Software, algorithmCHAINSAWStein, 2008http://www.ccp4.ac.uk/Version 7.0.077
Software, algorithmDIALSWinter et al., 2018https://dials.github.ioVersion 3.1.0
Software, algorithmExcel 2016MicrosoftRRID:SCR_016137Version 16.0
Software, algorithmGraphPad Prism 8GraphPad SoftwareRRID:SCR_002798Version 8
Software, algorithmImageJNIHhttps://imagej.net/ RRID:SCR_003070Version 1.50
Software, algorithmImage Studio LiteLI-COR BiosciencesRRID:SCR_013715Version 5.2
Software, algorithmPISAKrissinel, 2015http://www.ccp4.ac.uk/pisa/Version 2.1.1
Software, algorithmMolProbityWilliams et al., 2018http://molprobity.biochem.duke.edu/Version 4.5
Software, algorithmPHASERMcCoy et al., 2007https://www.phenix-online.org/Version 2.8.2
Software, algorithmPyMOLThe PyMOL Molecular Graphics Systemhttps://pymol.org/2/Version 2.4.0
Software, algorithmRR Foundation for Statistical Computinghttps://www.r-project.org/Version 3.2.4
Software, algorithmREFMAC5Murshudov et al., 1997http://www.ccp4.ac.uk/Version 5.8.0258
Software, algorithmSCULPTORBunkóczi and Read, 2011http://www.ccp4.ac.uk/Version 0.0.3
Software, algorithmXDSKabsch, 2010https://xds.mr.mpg.de/Version Nov11-2017
Software, algorithmXIA2Winter, 2009https://xia2.github.io/index.htmlVersion 0.3.7.0

Strains, media, and growth conditions

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E. coli and C. crescentus were grown in LB and PYE, respectively. When appropriate, media were supplemented with antibiotics at the following concentrations (liquid/solid media for C. crescentus; liquid/solid media for E. coli [μg/mL]): carbenicillin (E. coli only: 50/100), chloramphenicol (1/2; 20/30), kanamycin (5/25; 30/50), and oxytetracycline (1/2; 12/12).

Plasmids and strains construction

Construction of pET21b::parB∆CTD-(his)6

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The coding sequence of a C-terminally truncated C. crescentus ParB (ParB∆CTD, lacking the last 50 amino acids) was amplified by PCR using primers NdeI-Ct-ParB-F and HindIII-Ct-ParB-R, and pET21b::parB-(his)6 (Lim et al., 2014) as template. The pET21b plasmid backbone was generated via a double digestion of pET21b::parB-(his)6 with NdeI and HindIII. The resulting backbone was subsequently gel-purified and assembled with the PCR-amplified fragment of parB∆CTD using a 2X Gibson master mix (NEB). Gibson assembly was possible owing to a 23 bp sequence shared between the NdeI-HindIII-cut pET21b backbone and the PCR fragment. These 23 bp regions were incorporated during the synthesis of primers NdeI-Ct-ParB-F and HindIII-Ct-ParB-R. The resulting plasmids were sequence verified by Sanger sequencing (Eurofins, Germany).

Construction of pET21b::parB-(his)6 (WT and mutants)

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DNA fragments containing mutated parB genes (parB*) were chemically synthesized (gBlocks, IDT). The NdeI-HindIII-cut pET21b plasmid backbone and parB* gBlocks fragments were assembled together using a 2X Gibson master mix (NEB). Gibson assembly was possible owing to a 23 bp sequence shared between the NdeI-HindIII-cut pET21b backbone and the gBlocks fragment. The resulting plasmids were sequenced verified by Sanger sequencing (Genewiz, UK).

pENTR::attL1-parB (WT/mutant)-attL2

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The coding sequences of C. crescentus ParB (WT/mutants) were amplified by PCR and Gibson assembled into plasmid pENTR (Invitrogen) so that parB is flanked by phage attachment sites attL1 and attL2, that is, Gateway cloning compatible. Correct mutations were verified by Sanger sequencing (Genewiz, UK).

pMT571-1xFLAG-DEST

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Plasmid pMT571 (Thanbichler et al., 2007) was first digested with NdeI and NheI. The plasmid backbone was gel-purified and eluted in 50 µL of water. The FLAG-attR1-ccdB-chloramphenicolR-attR2 cassette was amplified by PCR using primers P1952 and P1953, and pML477 as template. The resulting PCR fragment and the NdeI-NheI-cut pMT571 were assembled together using a 2X Gibson master mix (NEB). Gibson assembly was possible owing to a 23 bp sequence shared between the two DNA fragments. These 23 bp regions were incorporated during the primer design to amplify the FLAG-attR1-ccdB-chloramphenicolR-attR2 cassette. The resulting plasmid was sequence verified by Sanger sequencing (Eurofins, Germany).

pMT571-1xFLAG::parB (WT/mutants)

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The parB (WT/mutant) genes were recombined into a Gateway-compatible destination vector pMT571-1xFLAG-DEST via LR recombination reaction (Invitrogen). For LR recombination reactions: 1 µL of purified pENTR::attL1-parB (WT/mutant)-attL2 was incubated with 1 µL of the destination vector pMT571-1xFLAG-DEST, 1 µL of LR Clonase II master mix, and 2 µL of water in a total volume of 5 µL. The reaction was incubated for an hour at room temperature before being introduced into E. coli DH5α cells by heat-shock transformation. Cells were then plated out on LB agar+ tetracycline. Resulting colonies were restruck onto LB agar+ kanamycin and LB agar+ tetracycline. Only colonies that survived on LB+ tetracycline plates were subsequently used for culturing and plasmid extraction.

pKTN25::cfp-parB (WT/E102A)

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The coding sequence of ParB (WT/E102A) was amplified by PCR using primers P3392 and P3393, and pET21b::C. crescentus ParB (WT/E102A)-His6 as template. The resulting DNA was gel-purified and assembled with a BglII-EcoRI-cut pVCFPN-5 (Thanbichler et al., 2007) using a 2X Gibson master mix, to result in vectors where the cfp is fused to the 5′-end of parB (WT/E102A). Gibson assembly was possible owing to a 23 bp sequence shared between the BglII-EcoRI-cut pVCFPN-5 backbone and the PCR fragment. To create vectors for expressing ParB (WT/E102A) in E. coli, the cfp-parB (WT/E102A) segment was amplified by PCR using primers P3396 and P3397, and pVCFPN-5::parB (WT/E102A) as template. The resulting DNA was then assembled with a HindIII-ClaI-cut pKTN25 (Karimova et al., 1998) using a 2X Gibson master mix. Gibson assembly was possible owing to a 23 bp sequence shared between the HindIII-ClaI-cut pKTN25 backbone and the PCR fragment. Note that the double digestion with HindIII and ClaI removed the T25-encoding gene from the pKTN25 plasmid. The resulting vectors pKTN25::cfp-parB (WT/E102A) allow for the expression of CFP-tagged ParB (WT/E102A) from an IPTG-inducible lactose promoter (Plac).

Strains TLE1146 (AB1157 ygcE::260 bp parS::apramycinR)

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Lambda Red recombineering (Datsenko and Wanner, 2000) was used to insert a cassette consisting of 260 bp C. crescentus parS3-4 sites and an apramycin antibiotic resistance gene aac(3)IV at the ygcE locus on the E. coli chromosome. To generate the first half of the cassette, DNA containing parS3-4 sites was amplified by PCR using P1304 and P1305, and C. crescentus genomic DNA as template. To generate the second half of the cassette, DNA containing aac(3)IV was amplified by PCR using P1306 and P1307, and pIJ773 as template (Gust et al., 2003). The two resulting PCR products were gel-purified and joined together using a 2X Gibson master mix. The full-length 260 bp parS::apramycinR cassette was further amplified by PCR using P1304 and P1307. P1304 and P1307 also carry 49 bp homology to the left or the right of the insertion point at the ygcE locus. The resulting PCR product was gel-extracted and electroporated into an arabinose-induced E. coli AB1157/pKD46 cells. Colonies that formed on LB+ apramycin were restruck on LB+ apramycin and incubated at 42°C to cure of pKD46 plasmid. Finally, the correct insertion of the parS-apramycinR cassette was verified by PCR and Sanger sequencing.

Strains AB1157 + pKTN25::cfp-parB (WT/E102A)

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E. coli AB1157 cells were made competent chemically and were transformed with pKTN25-cfp-parB (WT/E102A) to result in strains TLE3077 and TLE3078, respectively.

Strains TLE1146 + pKTN25::cfp-parB (WT/E102A)

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E. coli TLE1146 cells were made competent chemically and were transformed with pKTN25-cfp-parB (WT/E102A) to result in strains TLE3079 and TLE3080, respectively.

Strains MT148 + pMT571-1xFLAG::parB (WT/mutants)

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Electro-competent C. crescentus CN15N cells were electroporated with pMT571-1xFLAG::ParB (WT/mutants) plasmid to allow for a single integration at the vanA locus. The correct integration was verified by PCR, and ΦCr30 phage lysate was prepared from this strain. Subsequently, van::Pvan-1xflag-parB (WT/mutant), marked by a tetracyclineR cassette, was transduced by phage ΦCr30 into MT148 (Thanbichler and Shapiro, 2006) to result in strains TLS3050-TLS3060.

Protein overexpression and purification

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Plasmid pET21b::parB∆CTD-(his)6 was introduced into E. coli Rosetta (DE3)-competent cells (Merck) by heat-shock transformation. 40 mL overnight culture was used to inoculate 4 L of LB medium + carbenicillin + chloramphenicol. Cells were grown at 37°C with shaking at 250 rpm to an OD600 of ~0.4. The culture was then left in the cold room to cool to 28°C before isopropyl-β-D-thiogalactopyranoside (IPTG) was added at a final concentration of 0.5 mM. The culture was shaken for an additional 3 hr at 30°C before cells were pelleted by centrifugation. Pelleted cells were resuspended in a buffer containing 100 mM Tris-HCl pH 8.0, 300 mM NaCl, 10 mM imidazole, 5% (v/v) glycerol, 1 µL of Benzonase nuclease (Merck), 5 mg of lysozyme (Merck), and an EDTA-free protease inhibitor tablet (Merck). Cells were further lyzed by sonification (10 cycles of 15 s with 10 s resting on ice in between each cycle). The cell debris was removed through centrifugation at 28,000 g for 30 min and the supernatant was filtered through a 0.45 µm sterile filter (Sartorius). The protein was then loaded into a 1 mL HisTrap column (GE Healthcare) that had been pre-equilibrated with buffer A (100 mM Tris-HCl pH 8.0, 300 mM NaCl, 10 mM imidazole, and 5% [v/v] glycerol). Protein was eluted from the column using an increasing (10–500 mM) imidazole gradient in the same buffer. ParB∆CTD-containing fractions were pooled and diluted to a conductivity of 16 mS/cm before being loaded onto a 1 mL Heparin HP column (GE Healthcare) that had been pre-equilibrated with 100 mM Tris-HCl pH 8.0, 25 mM NaCl, and 5% (v/v) glycerol. Protein was eluted from the Heparin column using an increasing (25 mM to 1 M NaCl) salt gradient in the same buffer. ParB∆CTD fractions were pooled and analyzed for purity by SDS-PAGE. Glycerol was then added to ParB∆CTD fractions to a final volume of 10% (v/v), followed by 10 mM EDTA and 1 mM DTT. The purified ParB∆CTD was subsequently aliquoted, snap frozen in liquid nitrogen, and stored at –80°C. ParB∆CTD that was used for X-ray crystallography was further polished via a gel-filtration column. To do so, purified ParB∆CTD was concentrated by centrifugation in an Amicon Ultra-15 3 kDa cutoff spin filters (Merck) before being loaded into a Superdex-200 gel filtration column (GE Healthcare). The gel filtration column was pre-equilibrated with buffer containing 10 mM Tris-HCl pH 8.0 and 250 mM NaCl. ParB∆CTD fractions were then pooled and analyzed for purity by SDS-PAGE.

Other C-terminally His-tagged ParB mutants were purified using HIS-Select Cobalt gravity flow columns as described previously (Jalal et al., 2020b). Purified proteins were desalted using a PD-10 column (Merck), concentrated using an Amicon Ultra-4 10 kDa cutoff spin column (Merck), and stored at –80°C in a storage buffer (100 mM Tris-HCl pH 8.0, 300 mM NaCl, and 10% [v/v] glycerol). Purified ParB mutants that were used in BMOE crosslinking experiments were buffer-exchanged and stored in a storage buffer supplemented with TCEP instead (100 mM Tris-HCl pH 7.4, 300 mM NaCl, 10% [v/v] glycerol, and 1 mM TCEP).

Different batches of proteins were purified by ASBJ and NTT. Both biological (new sample preparations from a stock aliquot) and technical (same sample preparation) replicates were performed for assays in this study.

DNA preparation for crystallization, EnzChek phosphate release assay, and differential radical capillary action of ligand assay (DRaCALA)

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A 22 bp palindromic single-stranded DNA fragment (parS: GGATGTTTCACGTGAAACA TCC) (100 µM in 1 mM Tris-HCl pH 8.0, 5 mM NaCl buffer) was heated at 98°C for 5 min before being left to cool down to room temperature overnight to form 50 µM double-stranded parS DNA. The core sequence of parS is underlined.

Protein crystallization, structure determination, and refinement

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Crystallization screens for the C. crescentus ParB∆CTD-parS complex were set up in sitting-drop vapor diffusion format in MRC2 96-well crystallization plates with drops comprising 0.3 µL precipitant solution and 0.3 µL of protein-DNA complex, and incubated at 293 K. His-tagged ParB∆CTD (~10 mg/mL) was mixed with a 22 bp parS duplex DNA at a molar ratio of 2:1.2 (protein monomer:DNA) in buffer containing 10 mM Tris-HCl pH 8.0 and 250 mM NaCl. The ParB∆CTD-parS crystals grew in a solution containing 20.5% (w/v) PEG 3350, 260 mM magnesium formate, and 10% (v/v) glycerol. After optimization of an initial hit, suitable crystals were cryoprotected with 20% (v/v) glycerol and mounted in Litholoops (Molecular Dimensions) before flash-cooling by plunging into liquid nitrogen. X-ray data were recorded on beamline I04-1 at the Diamond Light Source (Oxfordshire, UK) using a Pilatus 6M-F hybrid photon counting detector (Dectris), with crystals maintained at 100 K by a Cryojet cryocooler (Oxford Instruments). Diffraction data were integrated and scaled using XDS (Kabsch, 2010) via the XIA2 expert system (Winter, 2009) then merged using AIMLESS (Evans and Murshudov, 2013). Data collection statistics are summarized in Table 1. The majority of the downstream analysis was performed through the CCP4i2 graphical user interface (Potterton et al., 2018).

Table 1
X-ray data collection and processing statistics.
StructureC. crescentus ParB∆CTD-parS complexC. crescentus ParB∆CTD CTPɣS complex
Data collection
 Diamond Light Source beamlineI04-1I03
 Wavelength (Å)0.9160.976
 DetectorPilatus 6M-FEiger2 XE 16M
 Resolution range (Å)72.96–2.90 (3.08–2.90)70.59–2.73 (2.86–2.73)
 Space groupP21P21
 Cell parameters (Å/°)a = 54.3, b = 172.9, c = 72.9, β = 90.5a = 69.5, b = 56.1, c = 71.4, β = 98.4
 Total no. of measured intensities198,135 (33888)92,266 (8473)
 Unique reflections29,654 (4775)14,516 (1756)
 Multiplicity6.7 (7.1)6.4 (4.8)
 Mean I/σ(I)8.7 (1.4)5.4 (1.2)
 Completeness (%)99.7 (100.0)98.8 (91.4)
Rmerge*0.135 (1.526)0.195 (1.210)
Rmeas0.146 (1.646)0.212 (1.357)
CC½0.997 (0.677)0.991 (0.825)
 Wilson B value (Å2)81.657.7
Refinement
 Resolution range (Å)72.96–2.90 (2.98–2.90)70.59–2.73 (2.80–2.73)
 Reflections: working/free§28155/146613824/678
Rwork0.240 (0.366)0.248 (0.371)
Rfree0.263 (0.369)0.284 (0.405)
 Ramachandran plot: favored/allowed/disallowed** (%)95.2/4.8/095.5/4.5/0
 R.m.s. bond distance deviation (Å)0.0050.002
 R.m.s. bond angle deviation (°)1.051.19
 Mean B factors: protein/DNA/other/ overall (Å2)98/74/-/9281/-/61/77
PDB accession code6T1F7BM8
  1. Values in parentheses are for the outer resolution shell.

  2. *

    Rmerge = ∑hkli |Ii(hkl)− ⟨I(hkl)⟩|/ ∑hkliIi(hkl).

  3. Rmeas = ∑hkl [N/(N− 1)]1/2 × ∑i |Ii(hkl)− ⟨I(hkl)⟩|/ ∑hkliIi(hkl), where Ii(hkl) is the ith observation of reflection hkl, ⟨I(hkl)⟩ is the weighted average intensity for all observations i of reflection hkl, and N is the number of observations of reflection hkl.

  4. CC½ is the correlation coefficient between symmetry equivalent intensities from random halves of the dataset.

  5. §

    The dataset was split into ‘working’ and ‘free’ sets consisting of 95% and 5% of the data, respectively. The free set was not used for refinement.

  6. The R-factors Rwork and Rfree are calculated as follows: R = ∑(| Fobs - Fcalc |)/∑| Fobs |, where Fobs and Fcalc are the observed and calculated structure factor amplitudes, respectively.

  7. **

    As calculated using MolProbity (Chen et al., 2010).

The ParB∆CTD-parS complex crystallized in space group P21 with cell parameters of a = 54.3, b = 172.9, c = 72.9 Å, and β = 90.5° (Table 1). Analysis of the likely composition of the asymmetric unit (ASU) suggested that it contains four copies of the ParB∆CTD monomer and two copies of the 22 bp parS DNA duplex, giving an estimated solvent content of ~47%.

Interrogation of the Protein Data Bank with the sequence of the C. crescentus ParB∆CTD revealed two suitable template structures for molecular replacement: apo-ParB∆CTD from Thermus thermophilus (Leonard et al., 2004) (PDB accession code: 1VZ0; 46% identity over 82% of the sequence) and Helicobacter pylori ParB∆CTD bound to parS DNA (Chen et al., 2015) (PDB accession code: 4UMK; 42% identity over 75% of the sequence). First, single subunits taken from these two entries were trimmed using SCULPTOR (Bunkóczi and Read, 2011) to retain the parts of the structure that aligned with the C. crescentus ParB∆CTD sequence, and then all side chains were truncated to Cβ atoms using CHAINSAW (Stein, 2008). Comparison of these templates revealed a completely different relationship between the NTD and the DBD. Thus, we prepared search templates based on the individual domains rather than the subunits. The pairs of templates for each domain were then aligned and used as ensemble search models in PHASER (McCoy et al., 2007). For the DNA component, an ideal B-form DNA duplex was generated in COOT (Emsley and Cowtan, 2004) from a 22 bp palindromic sequence of parS. A variety of protocols were attempted in PHASER (McCoy et al., 2007), the best result was obtained by searching for the two DNA duplexes first, followed by four copies of the DBD, giving a TFZ score of 10.5 at 4.5 Å resolution. We found that the placement of the DBDs with respect to the DNA duplexes was analogous to that seen in the H. pylori ParB∆CTD-parS complex. After several iterations of rebuilding in COOT and refining the model in REFMAC5 (Murshudov et al., 1997), it was possible to manually dock one copy of the NTD template (from 1VZ0) into weak and fragmented electron density such that it could be joined to one of the DBDs. A superposition of this more complete subunit onto the other three copies revealed that in only one of these did the NTD agree with the electron density. Inspection of the remaining unfilled electron density showed evidence for the last two missing NTDs, which were also added by manual docking of the domain template (from 1VZ0). For the final stages, TLS refinement was used with a single TLS domain defined for each protein chain and for each DNA strand. The statistics of the final refined model, including validation output from MolProbity (Chen et al., 2010), are summarized in Table 1.

Crystallization screens for the C. crescentus ParB∆CTD-CTPɣS complex crystal were also set up in sitting-drop vapor diffusion format in MRC2 96-well crystallization plates with drops comprising 0.3 µL precipitant solution and 0.3 µL of protein solution (~10 mg/mL) supplemented with 1 mM CTPɣS (Jena Biosciences) and 1 mM MgCl2, and incubated at 293 K. The ParB∆CTD-CTPɣS crystals grew in a solution containing 15% (w/v) PEG 3350, 0.26 M calcium acetate, 10% (v/v) glycerol, 1 mM CTPɣS, and 1 mM MgCl2. Suitable crystals were cryoprotected with 20% (v/v) glycerol and mounted in Litholoops (Molecular Dimensions) before flash-cooling by plunging into liquid nitrogen. X-ray data were recorded on beamline I03 at the Diamond Light Source (Oxfordshire, UK) using an Eiger2 XE 16M hybrid photon counting detector (Dectris), with crystals maintained at 100 K by a Cryojet cryocooler (Oxford Instruments). Diffraction data were integrated and scaled using DIALS (Winter et al., 2018) via the XIA2 expert system (Winter, 2009), then merged using AIMLESS (Evans and Murshudov, 2013). Data collection statistics are summarized in Table 1. The majority of the downstream analysis was performed through the CCP4i2 graphical user interface (Potterton et al., 2018).

The ParB∆CTD-CTPɣS complex crystallized in space group P21 with cell parameters of a = 69.5, b = 56.1, c = 71.4 Å, and β = 98.4° (Table 1). Analysis of the likely composition of the ASU suggested that it contains two copies of the ParB∆CTD monomer giving an estimated solvent content of ~50%. Molecular replacement templates were generated from the ParB∆CTD-parS complex solved above. Attempts to solve the structure in PHASER using individual subunits taken from the latter in both conformations did not yield any convincing solutions, suggesting that the subunits had adopted new conformations. Given that the two subunit conformations observed in the previous structure differed largely in the relative dispositions of DBD and NTDs, we reasoned that a better outcome might be achieved by searching for the DBD and NTD separately. This time PHASER successfully placed two copies of each domain in the ASU such that they could be reconnected to give two subunits in a new conformation. The result was subjected to 100 cycles of jelly-body refinement in REFMAC5 before rebuilding with BUCCANEER (Cowtan, 2006) to give a model in which 77% of the expected residues had been fitted into two chains and sequenced. The model was completed after further iterations of model editing in COOT and refinement with REFMAC5. In this case, TLS refinement was not used as this gave poorer validation results. The statistics of the final refined model, including validation output from MolProbity (Chen et al., 2010), are summarized in Table 1.

Measurement of protein-DNA interaction by BLI assay

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BLI experiments were conducted using a BLItz system equipped with High Precision Streptavidin 2.0 (SAX2) Biosensors (Molecular Devices). BLItz monitors wavelength shifts (nm) resulting from changes in the optical thickness of the sensor surface during association or dissociation of the analyte. All BLI experiments were performed at 22°C. The streptavidin biosensor was hydrated in a low-salt-binding buffer (100 mM Tris-HCl pH 8.0, 100 mM NaCl, 1 mM MgCl2, and 0.005% [v/v] Tween 20) for at least 10 min before each experiment. Biotinylated double-stranded DNA (dsDNA) was immobilized onto the surface of the SA biosensor through a cycle of baseline (30 s), association (120 s), and dissociation (120 s). Briefly, the tip of the biosensor was dipped into a binding buffer for 30 s to establish the baseline, then to 1 μM biotinylated dsDNA for 120 s, and finally to a low-salt-binding buffer for 120 s to allow for dissociation.

After the immobilization of DNA on the sensor, association reactions were monitored at 1 μM dimer concentration of ParB with an increasing concentration of CTP (0, 1, 5, 10, 50, 100, 500, 1000 µM) for 120 s. At the end of each binding step, the sensor was transferred into a protein-free binding buffer to follow the dissociation kinetics for 120 s. The sensor can be recycled by dipping in a high-salt buffer (100 mM Tris-HCl pH 8.0, 1000 mM NaCl, 10 mM EDTA, and 0.005% [v/v] Tween 20) for 5 min to remove bound ParB.

For the dissociation step in the BLI experiments in Figure 7A, the probe was returned to either a low-salt-binding buffer (100 mM Tris-HCl pH 8.0, 100 mM NaCl, 1 mM MgCl2, and 0.005% [v/v] Tween 20) for 30 s or a high-salt buffer (100 mM Tris-HCl pH 8.0, 1 M NaCl, 1 mM MgCl2, and 0.005% [v/v] Tween 20) for 30 s.

For experiments in Figure 7C, DNA-coated tips were dipped into 300 µL of restriction solution (266 µL of water, 30 µL of 10× buffer CutSmart [NEB], and 3 µL of BamHI-HF restriction enzyme [20,000 units/mL]) for 2 hr at 37°C. As a result, closed DNA on the BLI surface was cleaved to generate a free DNA end.

For experiments in Figure 5—figure supplement 4C, purified ParB (Q35C) was incubated with 1 mM CTPɣS in a binding buffer (100 mM Tris-HCl pH 7.4, 100 mM NaCl, 1 mM MgCl2) for 30 min before BMOE was added to 1 mM. DTT was then added to the final concentration of 1 mM to quench the reaction. Subsequently, crosslinked ParB (Q35C) was buffer-exchanged into a storage buffer (100 mM Tris-HCl pH 8.0, 300 mM NaCl, and 10% glycerol) using 0.5 mL Zeba desalting columns (ThermoFisher). BLITZ assays were performed using 5 μM dimer concentration of crosslinked ParB (Q35C) ± 1 mM CTP.

All sensorgrams recorded during BLI experiments were analyzed using the BLItz analysis software (BLItz Pro version 1.2, Molecular Devices) and replotted in R for presentation. Each experiment was triplicated, standard deviations were calculated in Excel, and a representative sensorgram is presented in Figure 5—figure supplement 4, Figure 6—figure supplement 2, and Figure 7.

Differential radical capillary action of ligand assay (DRaCALA) or membrane-spotting assay

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Purified C. crescentus ParB-His6 (WT and mutants, at final concentrations of 0.7, 1.5, 3.1, 6.2, and 12.5 µM) were incubated with 5 nM radiolabeled P32-α-CTP (Perkin Elmer), 30 µM of unlabeled CTP (ThermoFisher), and 1.5 μM of 22 bp parS DNA duplex in the binding buffer (100 mM Tris pH 8.0, 100 mM NaCl, and 10 mM CaCl2) for 10 min at room temperature. 4 μL of samples were spotted slowly onto a nitrocellulose membrane and air-dried. The nitrocellulose membrane was wrapped in cling film before being exposed to a phosphor screen (GE Healthcare) for 2 min. Each DRaCALA assay was triplicated, and a representative autoradiograph was shown. Data were quantified using Multi-Gauge software 3.0 (Fujifilm), the bound fraction were quantified as described previously (Roelofs et al., 2011). Error bars represent standard deviations from triplicated experiments.

Measurement of CTPase activity by EnzChek phosphate release assay

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CTP hydrolysis was monitored using an EnzCheck Phosphate Assay Kit (ThermoFisher). Samples (100 µL) containing a reaction buffer supplemented with an increasing concentration of CTP (0, 1, 5, 10, 50, 100, 500, and 1000 µM), 0.5 µM of 22 bp parS DNA, and 1 µM ParB (WT or mutants) were assayed in a Biotek EON plate reader at 25°C for 8 hr with readings every minute. The reaction buffer (1 mL) typically contained 740 μL Ultrapure water, 50 μL 20× reaction buffer (100 mM Tris pH 8.0, 2 M NaCl, and 20 mM MgCl2), 200 μL MESG substrate solution, and 10 μL purine nucleoside phosphorylase enzyme (one unit). Reactions with buffer only or buffer + CTP + 22 bp parS DNA only were also included as controls. The plates were shaken at 280 rpm continuously for 8 hr at 25°C. The inorganic phosphate standard curve was also constructed according to the manual. The results were analyzed using Excel and the CTPase rates were calculated using a linear regression fitting in Excel. Error bars represent standard deviations from triplicated experiments.

In vitro crosslinking assay using a sulfhydryl-to-sulfhydryl crosslinker BMOE

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A 50 µL mixture of 8 µM ParB mutants (with residues at specific positions in the NTD, DBD, or CTD substituted to cysteine) ± CTP (0–1000 µM) ± 0.5 µM DNA (a 22 bp linear DNA or a 3 kb circular parS/scrambled parS plasmid) was assembled in a reaction buffer (10 mM Tris-HCl pH 7.4, 100 mM NaCl, and 1 mM MgCl2) and incubated for 5 min at room temperature. BMOE (1 mM final concentration from a 20 mM stock solution) was then added, and the reaction was quickly mixed by three pulses of vortexing. SDS-PAGE sample buffer containing 23 mM β-mercaptoethanol was then added immediately to quench the crosslinking reaction. Samples were heated to 50°C for 5 min before being loaded on 12% Novex WedgeWell Tris-Glycine gels (ThermoFisher). Protein bands were stained with an InstantBlue Coomassie solution (Abcam) and band intensity was quantified using Image Studio Lite version 5.2 (LI-COR Biosciences). The crosslinked fractions were averaged, and their standard deviations from triplicated experiments were calculated in Excel.

For the experiment described in lane 8 of Figure 5C,D and Figure 5—figure supplement 2, crosslinking reactions were performed as described above; however, the reaction was quenched using a quenching buffer (10 mM Tris-HCl pH 7.4, 100 mM NaCl, 1 mM MgCl2, and 2.3 mM β-mercaptoethanol) instead. Subsequently, 1 µL of a non-specific DNA nuclease (Benzonase, 250 units/µL, Merck) was added, and the mixture was incubated at room temperature for a further 10 min before SDS-PAGE sample buffer was added. Samples were heated to 50°C for 5 min before being loaded on 4–12% Novex WedgeWell Tris-Glycine gels (ThermoFisher).

For the experiments described in lane 8 of Figure 5—figure supplement 3A, crosslinking and quenching reactions were performed as described above before 1 µL of TEV protease (10 units/µL, ThermoFisher) was added. The mixture was incubated at room temperature for a further 30 min before SDS-PAGE sample buffer was added. Samples were heated to 50°C for 5 min before being loaded on 4–12% Novex WedgeWell Tris-Glycine gels.

For experiments described in lane 9 of Figure 5—figure supplement 3B, proteins were released from gel slices by a ‘crush & soak’ method. Briefly, 10 gel slices were cut out from unstained SDS-PAGE gels and transferred to a 2 mL Eppendorf tube. Gel slices were frozen in liquid nitrogen and were crushed using a plastic pestle. The resulting paste was soaked in 500 µL of soaking buffer (10 mM Tris-HCl pH 8, 100 mM NaCl, 1 mM MgCl2, and 1 µL of Benzonase [250 units/µL]), and the tube was incubated with rotation in a rotating wheel overnight. On the next day, the tube was centrifuged at 13,000 rpm for 5 min and the supernatant was transferred to a new 1.5 mL Eppendorf tube. The sample volume was reduced to ~50 µL using a SpeedVac vacuum concentrator before SDS-PAGE sample buffer was added in. The entire sample was loaded onto a single well of a 4–12% WedgeWell Tris-Glycine gel.

For experiments described in Figure 5—figure supplement 1, a circular parS-harboring plasmid was linearized at an unique HindIII site by HindIII-HF restriction enzyme. After restriction, the linearized DNA was extracted with phenol-chloroform and ethanol precipitated before being used in double-crosslinking experiments.

Polyacrylamide gels were submerged in an InstantBlue Coomassie solution (Abcam) to stain for protein or in a SYBR Green solution (ThermoFisher) to stain for DNA. Denatured samples were also loaded on 1% TAE agarose gels and electrophoresed at 120 V for 40 min at room temperature. Afterwards, agarose gels were submerged in a SYBR green solution to stain for DNA.

Chromatin immunoprecipitation with deep sequencing (ChIP-Seq)

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α-FLAG ChIP-seq experiments on formaldehyde-fixed C. crescentus cells, and the subsequent data analysis was performed exactly as reported previously (Tran et al., 2018).

For ChIP-seq experiments on fixed E. coli cells, cells harboring pKTN25-cfp-parB (WT) or pKTN25-cfp-parB (E102A) were grown in 50 mL LB at 30°C to mid exponential phase (OD600 ∼ 0.4, no IPTG was added). Subsequently, formaldehyde is added to a final concentration of 1% to fix the cells. All following steps are identical to ChIP-seq for C. crescentus, except that α-GFP antibody coupled to sepharose beads (Abcam) was used to immunoprecipitate CFP-tagged ParB–DNA complexes.

Each ChIP-seq experiment was duplicated using biological replicates. For a list of ChIP-seq experiments and their replicates in this study, see Supplementary file 1C.

Immunoblot analysis

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For western blot analysis, C. crescentus or E. coli cells were pelleted and resuspended directly in 1× SDS sample buffer, then heated to 95°C for 5 min before loading. Total protein was run on 12% Novex WedgeWell gels (ThermoFisher) at 150 V for separation. The same amount of total protein was loaded on each lane. Resolved proteins were transferred to PVDF membranes using the Trans-Blot Turbo Transfer System (BioRad) and probed with either a 1:5000 dilution of α-FLAG HRP-conjugated antibody (Merck) antibody or a 1:5000 dilution of α-GFP HRP-conjugated antibody (Abcam). Blots were imaged after incubation with SuperSignal West PICO PLUS Chemiluminescent Substrate (ThermoFisher) using an Amersham Imager 600 (GE Healthcare). Western blot experiments were duplicated using biological replicates.

Fluorescence microscopy and image analysis

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TLS3079 and TLS3080 were grown in M9 media supplemented with kanamycin (30 µg/mL) until OD600 ~ 0.1 prior to imaging. The expression of cfp-parB (WT/E102A) was induced with 0.25 mM IPTG in culture for 60 min before imaging. Imaging was performed using a wide-field epifluorescence microscope (Eclipse Ti-2E, Nikon) with a 63× oil immersion objective (NA 1.41), illumination from pE4000 light source, Hamamatsu Orca Flash 4.0 camera, and a motorized XY stage. Images were acquired using NIS-elements software (version 5.1). For imaging in the CFP channel, 435 nm excitation wavelength was used with 1 s exposure.

Images were analyzed using ImageJ, and plots were generated in GraphPad Prism 8.0. For extracting information on number of ParB foci per cell as well as intensity of ParB in foci, the following analysis pipeline was implemented: cell masks were generated in ImageJ using analyze particle function on thresholds applied to phase profiles. Separately, even background subtraction function was applied to fluorescence profiles, images were convolved (using ‘subtract background’ and ‘convolve’ functions in ImageJ), and regions of interest (ROIs) for foci were detected via an application of appropriate thresholds. The cell masks and ROIs thus detected were applied to the raw data (after background correction) to extract intensity information for each ROI as well as total cell fluorescence. ROI intensity was plotted as a ratio of intensity within a focus (intensityloc) normalized to total cell intensity (intensitytotal). Along with intensity measurement, number of foci per cell was also recorded. The pipeline was implemented in ImageJ using the following command:

n = roiManager("count");for (j = 0; j < n; j++){roiManager("Select", j);run("Analyze Particles...", "size = 3–10 circularity = 0.40–1.00 display summarize add");}

Data availability

The accession number for the sequencing data reported in this paper is GSE168968. Atomic coordinates for protein crystal structures reported in this paper were deposited in the RCSB Protein Data Bank with the accession number 6T1F and 7BM8. All of these sequencing and X-ray crystallography data are already open to the public. All other data generated or analyzed during this study are included in the manuscript.

The following data sets were generated
    1. Le TBK
    (2021) NCBI Gene Expression Omnibus
    ID GSE168968. A CTP-dependent gating mechanism enables ParB spreading on DNA in Caulobacter crescentus.
    1. Jalal ASB
    2. Pastrana CL
    3. Tran NT
    4. Stevenson CEM
    5. Lawson DM
    6. Moreno-Herrero F
    7. Le TBK
    (2020) RCSB Protein Data Bank
    ID 6T1F. Crystal structure of the C-terminally truncated chromosome-partitioning protein ParB from Caulobacter crescentus complexed to the centromeric parS site.
    1. Jalal ASB
    2. Pastrana CL
    3. Tran NT
    4. Stevenson CEM
    5. Lawson DM
    6. Moreno-Herrero F
    7. Le TBK
    (2021) RCSB Protein Data Bank
    ID 7BM8. Crystal structure of the C-terminally truncated chromosome-partitioning protein ParB from Caulobacter crescentus complexed with CTP-gamma-S.

References

    1. Kabsch W
    (2010) Xds
    Acta Crystallographica Section D Biological Crystallography 66:125–132.
    https://doi.org/10.1107/S0907444909047337

Decision letter

  1. Anthony G Vecchiarelli
    Reviewing Editor; University of Michigan, United States
  2. Gisela Storz
    Senior Editor; National Institute of Child Health and Human Development, United States
  3. Anthony G Vecchiarelli
    Reviewer; University of Michigan, United States

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Acceptance summary:

Bacterial ParB partition proteins have the novel property that they employ an unusual nucleotide cofactor for complex assembly at their specific DNA binding site, parS. The impact of this study is on our general understanding of this novel class of nucleotide-dependent processes, and the role that nucleotide-protein interactions play in DNA binding and bacterial physiology.

Decision letter after peer review:

Thank you for submitting your article "A CTP-dependent gating mechanism enables ParB spreading on DNA" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, including Anthony G Vecchiarelli as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Gisela Storz as the Senior Editor.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

All three reviewers agree that the work is exciting, high quality, will be of broad interest, and should be published in eLife. However, in the "Recommendations to the Authors" there are two control experiments that all three reviewers agree are simple to perform and required. Also, all three reviewers had concerns regarding the ChIP-seq data that need to be addressed prior to publication.

1) From Reviewer 1: A control experiment where the authors pre-treat ParB with cross-linker before addition of DNA substrate to show that premature and irreversible closing of the ring prevents interactions with parS as well as spreading/sliding. The pre-crosslinking might alter protein conformation in other ways (besides closing a gate) that might destroy the DNA binding site (so there are caveats). Nevertheless if crosslinking did not prevent DNA binding, that would affect the overall story.

2) From Reviewer 2: A linearized plasmid control. The authors use the binding to circular DNA as evidence that ParB is topologically constrained, compared to a 22 bp linear DNA substrate. But the latter might just be too short to make a stable (non-clamped) complex. The experiment is consistent with a clamped complex but does not formally show it. Or perhaps instead of Benzonase treatment, treat the complex with a restriction enzyme after the fact and show it disassembles. This data can be added as a supplemental figure so the authors don't need to repeat the original and redraw the figure. If not performed, then the language should reflect that the experiment does not prove topological constraints.

3) More critically, reviewers agree that the single MAJOR issue is the ChIP-seq data in Figure 8A. We all expected an expanded profile around parS sites with ParB[E102A] and that is clearly not happening. Also, the authors highlight the upstream extended signal that is amplified in the mutant and largely absent in WT, but this could just be a confounding interaction with the par operon or expression from the parB gene itself. All three reviewers appreciate the authors trying to bring their in vitro findings back into the cell, but it did not seem to work out as described in the text. Of note, the Gruber lab currently has a similar story on BioRxiv and their ChIP-seq data for their CTP-trap mutant of B.sub ParB shows an expansion of signal around the parS sites. As noted in our independent reviews, the authors need to re-interpret their ChIP-seq data, provide a better and more detailed explanation of these findings, and possibly tamper their claims.

Finally, please see the "Recommendations to the Authors" section from all three reviewers for suggestions to improve clarity and presentation before resubmission.

Reviewer #1 (Recommendations for the authors):

Ln 302-304 and Figure 8

The description and interpretation of the [E102A] ChIP-seq data, as having a more "extended profile", is not an entirely accurate. Figure 8A clearly shows that spread is only in one direction (upstream), and there is a massive signal around the parAB operon (particularly over the ParB gene) with [E102A] that is largely absent from the WT ChIP seq. This strong [E102A] signal over the parAB operon dissipates in both directions, and the spread from this specific location is described by the authors as [E102A]'s "extended profile". This result is striking but not discussed by the authors. The authors need to expand their description and interpretation of the ChIP data as it relates to this region, and speculate as to what they think this upstream signal around the parAB operon may represent.

Ln 316-317 and Figure 9

Given the flexibility of the N-terminal domain, do the authors think it possible that this domain flexibility plays a role in ParB's ability to not only close the ring, but also associate with other ParB dimers and/or with ParA? Given that the ParA interaction interface is on this flexible domain, I think the data here interfaces quite nicely with the BioRxiv preprint from the Mizuuchi group (already cited in the paper) that was also recently reviewed by eLife: https://www.biorxiv.org/content/10.1101/2021.01.24.427996v1. In particular: They find ParB can bind to ParA either using the two protomers of a single dimer or two protomers from distinct dimers. The former occurs in the absence of ligands, the latter upon addition of either CTP or parS, thus presumably corresponding to the state of ParB found in the cells near a parS site. The discussion would benefit from the authors putting their findings in the context of other ParB interactions shown to be required for chromosome segregation – ParA interaction and ParB dimer oligomerization.

Reviewer #2 (Recommendations for the authors):

1. The crosslinking at L224 supports the idea that DNA has left the DNA binding domain after clamp closure, consistent with the structural clashes observed. Where is L224 in the DNA binding region? The cartoon in Figure 5 places it at the C-terminal side of the DBD but it would help if the position of L224 was indicated on one of the structures, eg Figure 1B, in addition to the cartoons. Why is there a helix 10 in the structure (Figure 1) but not in the secondary structure of Figure 2-supp2? By my estimation L224 is in helix 10 but this is confusing.

2. Figure 8: The ChIPseq profile of E102A is confusing and warrants more explanation. The authors state it is "notably more extended than the profile of" wild-type ParB. The shaded area to which they refer does not center on a parS site. What is the explanation for this pattern? That the profile is more extended also predicts that the other parS peaks would be more extended, which does not appear to be the case. Why? If the peaks were normalized to the same height, would E102A peaks look broader?

3. I like the alanine scanning approach to target the residues that interact with CTP, in part because it is a more comprehensive survey of the CTP binding site, which is poorly understood compared to sites that bind ATP or GTP. This allowed them to identify E102A as a mutation that created a CTP dependent clamp with diminished dissociation rate because it could not hydrolyze CTP. But the analysis also created two other classes of mutants (pg 8). Although the authors would likely argue that their analysis is beyond the scope of the current study, it would add important insight to the story if the authors discussed or proposed explanations for the behavior of each class in the Discussion. For example, why do class II, which bind but do not hydrolyse CTP, behave differently than E102A? Based on their location could they be permanently closed?

4. Figure 5: There should be a linearized plasmid control. The authors use the binding to circular DNA as evidence that ParB is topologically constrained, compared to a 22 bp linear DNA substrate. But the latter might just be too short to make a stable (non-clamped) complex. The experiment is consistent with a clamped complex but does not formally show it. Or perhaps instead of Benzonase treatment, treat the complex with a restriction enzyme after the fact and show it disassembles.

Reviewer #3 (Recommendations for the authors):

The data on the recycling mechanism upon CTP hydrolysis presented in Figure 8A are not sufficiently convincing to conclude that without CTP hydrolysis (variant E102A) the clamped-ParB diffuse longer away from parS than wt ParB. Especially, the most important signal, which strongly extend the ParB signature over DNA on the left side, seems independent of parS loading; indeed, the peak is away from the first and weak parS site. It rather corresponds to the location of the parB gene, which suggests that the signal detected here does not correspond to sliding but rather to ParB synthesis. Therefore, this is against the author's conclusion that preventing CTP hydrolysis extend ParB sliding. In addition, the signal over the parB gene is increased 4-5 fold compare to wt while the parS-specific signals are decreased about 3-fold. This may prevent a direct comparison of the "spreading" signature. Also, if ParB (E102A) is trapped longer on DNA, one would expect that the ParB signal should be higher in between parS sites that are close together such as in between parS3-4-5-6. The authors should comment on this lack of increase, and discuss these in regard of the sliding model relatively to other assembly model.

https://doi.org/10.7554/eLife.69676.sa1

Author response

Essential revisions:

All three reviewers agree that the work is exciting, high quality, will be of broad interest, and should be published in eLife. However, in the "Recommendations to the Authors" there are two control experiments that all three reviewers agree are simple to perform and required. Also, all three reviewers had concerns regarding the ChIP-seq data that need to be addressed prior to publication.

1) From Reviewer 1: A control experiment where the authors pre-treat ParB with cross-linker BEFORE addition of DNA substrate to show that premature and irreversible closing of the ring prevents interactions with parS as well as spreading/sliding. The pre-crosslinking might alter protein conformation in other ways (besides closing a gate) that might destroy the DNA binding site (so there are caveats). Nevertheless if crosslinking did not prevent DNA binding, that would affect the overall story.

A premature closing of the ParB clamp, before being exposed to parS DNA, indeed prevents nucleation at parS as well as spreading/sliding. This observation was previously reported for C. crescentus ParB (Figure 6 of Jalal et al., eLife 2020). There, a prolonged (30-60min) pre-incubation of ParB (WT) with CTPɣS eliminated most of ParB binding to and sliding from parS. CTPɣS, even in the absence of parS DNA, gradually converted apo-ParB from an open to a closed protein clamp (Jalal et al., 2020; Soh et al., 2019). If not already bound on DNA, the now inaccessible DNA-binding domain of ParB cannot bind parS to nucleate and to subsequently slide to accumulate on DNA.

In the current manuscript, we have repeated the same experiment but with a crosslinkable ParB (Q35C), as requested by the reviewer, and observed the same behavior in the BLI assay (now reported in Figure 5—figure supplement 4B, 0 min vs. 30 min pre-incubation with CTPɣS).

We have also pre-crosslinked the closed-clamp form of ParB (Q35C) and subsequently used the crosslinked proteins in the same BLI assay (Figure 5—figure supplement 4B). To do so, purified ParB (Q35C) was incubated with CTPɣS for 30 min, then 1 mM BMOE was added to crosslink the N-terminal domains of ParB (Q35C) together. The crosslinking reaction was quenched with 1 mM DTT and ParB (Q35C) was purified away from excess BMOE and DTT using a Zeba buffer exchange column (See Materials and methods). Approx. 70% ParB (Q35C) was in a crosslinked form, consistent with a previous report (Jalal et al., 2020). Note that CTPɣS was required to convert apo-ParB (Q35C) to a closed clamp form before crosslinking. Without CTPɣS, most apo-ParB (E35C) is in an open clamp form (Jalal et al., 2020).

We have now added a sentence to the Results section to bring this important control to the attention of readers:

“This is also consistent with experiments that showed a premature and irreversible closing of ParB clamps, achieved either by an extended preincubation with CTPɣS (Jalal et al., 2020a and Figure 5—figure supplement 4B) or by pre-crosslinking a closed clamp form of ParB (Figure 5—figure supplement 4C), prevented nucleation at parS and DNA entrapment.”

2) From Reviewer 2: A linearized plasmid control. The authors use the binding to circular DNA as evidence that ParB is topologically constrained, compared to a 22 bp linear DNA substrate. But the latter might just be too short to make a stable (non-clamped) complex. The experiment is consistent with a clamped complex but does not formally show it. Or perhaps instead of Benzonase treatment, treat the complex with a restriction enzyme after the fact and show it disassembles. This data can be added as a supplemental figure so the authors don't need to repeat the original and redraw the figure. If not performed, then the language should reflect that the experiment does not prove topological constraints.

The reviewer raised important points and we have now performed additional controls with a linearized 3-kb parS DNA (reported in Figure 5—figure supplement 1).

We performed double BMOE crosslinking assays of dual-cysteine ParB variants + (circular/linearized) parS-containing plasmid DNA ± CTP. Purified ParB (Q35C I304C), ParB (L224C I304C), and ParB (Q35C L224C) were used for reactions in lanes 1-4, 5-8, and 9-12, respectively (Figure 5—figure supplement 1). Different DNA were employed in crosslinking reactions: a circular 3-kb parS plasmid (3 kb parS cir) or a 3-kb parS plasmid that had been linearized at an unique HindIII site by HindIII restriction enzyme (3 kb parS linear).

The high molecular weight (HMW) smear near the top of the polyacrylamide gel was observed only when ParB (Q35C I304C) or ParB (L224C I304C) was incubated with CTP and a circular parS plasmid (lanes 2 and 6, solid lines and asterisks, Figure 5—figure supplement 1), but not when a linearized parS plasmid was used (lanes 4 and 8, dashed lines and asterisks) or when CTP was omitted (lanes 1 and 5). ParB (Q35C L224C), as expected, did not produce a HMW smear even in the presence of CTP and a circular parS plasmid (see also Figure 5—figure supplement 2).

Overall, these results are consistent with a closed ParB clamp entrapping a topologically closed parS DNA, specifically within the DBD-CTD compartment. We have now revised the Results and Materials and Methods sections to describe these experiments.

3) More critically, reviewers agree that the single MAJOR issue is the ChIP-seq data in Figure 8A. We all expected an expanded profile around parS sites with ParB[E102A] and that is clearly not happening. Also, the authors highlight the upstream extended signal that is amplified in the mutant and largely absent in WT, but this could just be a confounding interaction with the par operon or expression from the parB gene itself. All three reviewers appreciate the authors trying to bring their in vitro findings back into the cell, but it did not seem to work out as described in the text. Of note, the Gruber lab currently has a similar story on BioRxiv and their ChIP-seq data for their CTP-trap mutant of B.sub ParB shows an expansion of signal around the parS sites. As noted in our independent reviews, the authors need to re-interpret their ChIP-seq data, provide a better and more detailed explanation of these findings, and possibly tamper their claims.

We agree entirely with all three reviewers and thank them for pointing this out. We have now revised the text and performed extra experiement to address the concern from the reviewers.

First, we have now described the ChIP-seq profile of ParB (E102A) in C. crescentus more accurately. Specifically, we removed this sentence “…the ChIP-seq profile of FLAG-ParB (E102A) … is notably more extended than the profile of FLAG-ParB (WT)…”. We highlighted the caveat associated with the instability of FLAG-ParB (E102A) protein in C. crescentus and cautioned against interpreting the roles of CTP hydrolysis in native C. crescentus host, both in the Results and the Discussion sections. We wrote:

“By contrast, the ChIP-seq profile of FLAG-ParB (E102A) is significantly reduced in height but has an extra peak over the parB coding sequence (Figure 8A, asterisk). […] Again, due to the caveat of a lower ParB (E102A) protein level in C. crescentus (Figure 8—figure supplement 1A), we could not reliably link the in vitro properties of ParB (E102A) to its behaviors in the native host.”

Second, to overcome the caveat of protein instability, we instead investigated the spreading of ParB (WT) vs. ParB (E102A) from parS by analysing the C. crescentus ParB/parS system in E. coli. E. coli does not possess a ParA/ParB homolog nor a parS-like sequence, thus it serves as a suitable heterologous host. C. crescentus parS sites 3 and 4 were engineered onto the E. coli chromosome at the ygcE locus (Figure 8B). CFP-tagged ParB (WT/E102A) was expressed from a leaky lactose promoter (Plac, no IPTG was added) on a medium-copy-number plasmid. CFP-ParB (WT/E102A) was produced at the same level, as judged by an immunoblot (Figure 8—figure supplement 1B).

We observed by ChIP-seq that CFP-ParB (WT) in an E. coli host spreads asymmetrically ~5 kb around parS sites. By contrast, the shape of the ParB (E102A) distribution was clearly different from that of ParB (WT); the profile was further expanded to both neighboring sides of parS (covering in total ~26 kb) at the expense of the enrichment at parS itself (Figure 8B). The more excessive spreading of ParB (E102A) might suggest that this variant, in the absence of CTP hydrolysis, persisted and perhaps slid further away from the loading site parS in E. coli. The reduced enrichment of ParB (E102A) at parS itself (Figure 8B) might be due to reduced cytoplasmic ParB (E102A) available to re-nucleate at parS and/or due to stably entrapped ParB (E102A) sterically hindering further nucleation events. We further noted that the ChIP-seq profile of CFP-ParB (E102A) in E. coli is highly asymmetrical, with more enrichment in the +2905-2911kb region than the +2885-2899kb region (shaded areas, Figure 8B). The asymmetrical spreading is possibly due to an impediment in one direction by roadblocks such as RNA polymerases or DNA-bound proteins, which have been shown previously to be able to interfere with ParB spreading (Balaguer et al., 2021; Breier and Grossman, 2007; Jalal et al., 2020; Murray et al., 2006; Rodionov et al., 1999; Soh et al., 2019). We also quantified the fluorescence intensity of CFP-ParB (WT/E102A) foci, and found a higher CFP signal for CFP-ParB (E102A) foci than that of CFP-ParB (WT) (Figure 8—figure supplement 3). This is also consistent with a scenario where CFP-ParB (E102A) occupies a more extended genomic area in our E. coli host. Altogether, at least in the heterologous E. coli host, the “clamp-locked” phenotype of ParB (E102A) implies a possible role of CTP hydrolysis and/or the release of hydrolytic products in re-opening wild-type ParB clamp to release DNA and to recycle ParB.

Lastly, we also noted that two preprints from the Gruber and the Thanbichler labs also reported the same phenomenon that the enrichment/height at parS itself is reduced but the enriched area around parS is expanded in the “clamp-locked” mutants in B. subtilis and M. xanthus (Antar et al., 2021; Osorio-Valeriano et al., 2021).

Finally, please see the "Recommendations to the Authors" section from all three reviewers for suggestions to improve clarity and presentation before resubmission.

Detailed responses to the specific points that reviewers have raised are given below:

Reviewer #1 (Recommendations for the authors):

Ln 302-304 and Figure 8

The description and interpretation of the [E102A] ChIP-seq data, as having a more "extended profile", is not an entirely accurate. Figure 8A clearly shows that spread is only in one direction (upstream), and there is a massive signal around the parAB operon (particularly over the ParB gene) with [E102A] that is largely absent from the WT ChIP seq. This strong [E102A] signal over the parAB operon dissipates in both directions, and the spread from this specific location is described by the authors as [E102A]'s "extended profile". This result is striking but not discussed by the authors. The authors need to expand their description and interpretation of the ChIP data as it relates to this region, and speculate as to what they think this upstream signal around the parAB operon may represent.

Please see my response to point 3 in the Essential Revisions for Authors.

Ln 316-317 and Figure 9

Given the flexibility of the N-terminal domain, do the authors think it possible that this domain flexibility plays a role in ParB's ability to not only close the ring, but also associate with other ParB dimers and/or with ParA? Given that the ParA interaction interface is on this flexible domain, I think the data here interfaces quite nicely with the BioRxiv preprint from the Mizuuchi group (already cited in the paper) that was also recently reviewed by eLife: https://www.biorxiv.org/content/10.1101/2021.01.24.427996v1. In particular: They find ParB can bind to ParA either using the two protomers of a single dimer or two protomers from distinct dimers. The former occurs in the absence of ligands, the latter upon addition of either CTP or parS, thus presumably corresponding to the state of ParB found in the cells near a parS site. The discussion would benefit from the authors putting their findings in the context of other ParB interactions shown to be required for chromosome segregation – ParA interaction and ParB dimer oligomerization.

These are excellent points and we have expanded the Discussion as followed:

“Recently, an F-plasmid ParB was shown to form biomolecular condensates in vivo that might bridge distal ParBF dimers together (Guilhas et al., 2020; Walter et al., 2020). […] Future work will provide important insights to better understand the mechanism of ParA-directed DNA segregation.”

We refrain from over-speculating here because we do not know enough and have not performed any experiments with phase separation or ParA in vivo. We, however, highlighted a few questions which we think are very important and hopefully they will motivate further investigation from multiple labs in our field.

Reviewer #2 (Recommendations for the authors):

1. The crosslinking at L224 supports the idea that DNA has left the DNA binding domain after clamp closure, consistent with the structural clashes observed. Where is L224 in the DNA binding region? The cartoon in Figure 5 places it at the C-terminal side of the DBD but it would help if the position of L224 was indicated on one of the structures, eg Figure 1B, in addition to the cartoons. Why is there a helix 10 in the structure (Figure 1) but not in the secondary structure of Figure 2-supp2? By my estimation L224 is in helix 10 but this is confusing.

Residue L224 locates in the loop that connects helix 9 and helix 10 of the DNA-binding domain (DBD). So the reviewer is correct that L224 positions towards the far C-terminal side of the DBD. We have now indicated the positions of L224 in Figure 1B.

The reviewer is correct that helix 10 can be seen in the ParB∆CTD-parS structure (Figure 1B) but not in the CTPɣS-bound structure. This is because of the poor electron density for helix 10 in the CTPɣS-bound structure, which prevented us from modeling in this last helix of the DBD. We have now mentioned this in the lenged of Figure 2 (that describes the CTPɣS-bound structure) to make this clear to readers.

2. Figure 8: The ChIPseq profile of E102A is confusing and warrants more explanation. The authors state it is "notably more extended than the profile of" wild-type ParB. The shaded area to which they refer does not center on a parS site. What is the explanation for this pattern? That the profile is more extended also predicts that the other parS peaks would be more extended, which does not appear to be the case. Why? If the peaks were normalized to the same height, would E102A peaks look broader?

Please see my response to point 3 in the Essential Revisions for Authors. Also, if the peaks were normalized to the same height, E102A peaks (in the ChIP-seq profiles for C. crescentus) are not broader either.

3. I like the alanine scanning approach to target the residues that interact with CTP, in part because it is a more comprehensive survey of the CTP binding site, which is poorly understood compared to sites that bind ATP or GTP. This allowed them to identify E102A as a mutation that created a CTP dependent clamp with diminished dissociation rate because it could not hydrolyze CTP. But the analysis also created two other classes of mutants (pg 8). Although the authors would likely argue that their analysis is beyond the scope of the current study, it would add important insight to the story if the authors discussed or proposed explanations for the behavior of each class in the Discussion. For example, why do class II, which bind but do not hydrolyse CTP, behave differently than E102A? Based on their location could they be permanently closed?

We noted that Class II mutants ParB (Q58A) and ParB (E135A) already had an elevated crosslinking efficiency even in the absence of CTP (Figure 6C). This premature clamp closing might indeed result in a less than wild-type level of DNA entrapment (Figure 6D). The reason the clamp closes prematurely for Class II mutants is not yet clear, and cannot easily be rationalized based on our current crystal structures, especially for the Q85A substitution. We wish to investiagate this class of mutants more closely in the near future. We hope the reviewer will understand.

4. Figure 5: There should be a linearized plasmid control. The authors use the binding to circular DNA as evidence that ParB is topologically constrained, compared to a 22 bp linear DNA substrate. But the latter might just be too short to make a stable (non-clamped) complex. The experiment is consistent with a clamped complex but does not formally show it. Or perhaps instead of Benzonase treatment, treat the complex with a restriction enzyme after the fact and show it disassembles.

Please see my response to point 2 in the Essential Revisions for Authors.

References:

Antar H, Soh Y-M, Zamuer S, Bock FP, Anchimiuk A, Rios PDL, Gruber S. 2021. Relief of ParB autoinhibition by parS DNA catalysis and ParB recycling by CTP hydrolysis promote bacterial centromere assembly. bioRxiv 2021.05.05.442573. doi:10.1101/2021.05.05.442573

Balaguer F de A, Aicart-Ramos C, Fisher GL, de Bragança S, Martin-Cuevas EM, Pastrana CL, Dillingham MS, Moreno-Herrero F. 2021. CTP promotes efficient ParB-dependent DNA condensation by facilitating one-dimensional diffusion from parS. eLife 10:e67554. doi:10.7554/eLife.67554

Breier AM, Grossman AD. 2007. Whole-genome analysis of the chromosome partitioning and sporulation protein Spo0J (ParB) reveals spreading and origin-distal sites on the Bacillus subtilis chromosome. Molecular Microbiology 64:703–718. doi:10.1111/j.1365-2958.2007.05690.x

Guilhas B, Walter J-C, Rech J, David G, Walliser NO, Palmeri J, Mathieu-Demaziere C, Parmeggiani A, Bouet J-Y, Le Gall A, Nollmann M. 2020. ATP-Driven Separation of Liquid Phase Condensates in Bacteria. Mol Cell 79:293-303.e4. doi:10.1016/j.molcel.2020.06.034

Jalal AS, Tran NT, Le TB. 2020. ParB spreading on DNA requires cytidine triphosphate in vitro. eLife 9:e53515. doi:10.7554/eLife.53515

Murray H, Ferreira H, Errington J. 2006. The bacterial chromosome segregation protein Spo0J spreads along DNA from parS nucleation sites. Molecular Microbiology 61:1352–1361. doi:10.1111/j.1365-2958.2006.05316.x

Osorio-Valeriano M, Altegoer F, Das CK, Steinchen W, Panis G, Connolley L, Giacomelli G, Feddersen H, Corrales-Guerrero L, Giammarinaro P, Hanßmann J, Bramkamp M, Viollier PH, Murray S, Schäfer LV, Bange G, Thanbichler M. 2021. The CTPase activity of ParB acts as a timing mechanism to control the dynamics and function of prokaryotic DNA partition complexes. bioRxiv 2021.05.05.442810. doi:10.1101/2021.05.05.442810

Rodionov O, Lobocka M, Yarmolinsky M. 1999. Silencing of genes flanking the P1 plasmid centromere. Science 283:546–549. doi:10.1126/science.283.5401.546

Sanchez A, Cattoni DI, Walter J-C, Rech J, Parmeggiani A, Nollmann M, Bouet J-Y. 2015. Stochastic Self-Assembly of ParB Proteins Builds the Bacterial DNA Segregation Apparatus. Cell Syst 1:163–173. doi:10.1016/j.cels.2015.07.013

Soh Y-M, Davidson IF, Zamuner S, Basquin J, Bock FP, Taschner M, Veening J-W, De Los Rios P, Peters J-M, Gruber S. 2019. Self-organization of parS centromeres by the ParB CTP hydrolase. Science 366:1129–1133. doi:10.1126/science.aay3965

Taylor JA, Seol Y, Neuman KC, Mizuuchi K. 2021. CTP and parS control ParB partition complex dynamics and ParA-ATPase activation for ParABS-mediated DNA partitioning. bioRxiv 2021.01.24.427996. doi:10.1101/2021.01.24.427996

Walter J-C, Rech J, Walliser N-O, Dorignac J, Geniet F, Palmeri J, Parmeggiani A, Bouet J-Y. 2020. Physical Modeling of a Sliding Clamp Mechanism for the Spreading of ParB at Short Genomic Distance from Bacterial Centromere Sites. iScience 23:101861. doi:10.1016/j.isci.2020.101861

https://doi.org/10.7554/eLife.69676.sa2

Article and author information

Author details

  1. Adam SB Jalal

    Department of Molecular Microbiology, John Innes Centre, Norwich, United Kingdom
    Contribution
    Formal analysis, Investigation, Methodology, Writing – original draft
    Competing interests
    None
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7794-8834
  2. Ngat T Tran

    Department of Molecular Microbiology, John Innes Centre, Norwich, United Kingdom
    Contribution
    Formal analysis, Investigation
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7186-3976
  3. Clare EM Stevenson

    Department of Biochemistry and Metabolism, John Innes Centre, Norwich, United Kingdom
    Contribution
    Investigation
    Competing interests
    None
  4. Afroze Chimthanawala

    1. National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India
    2. SASTRA University, Thanjavur, Tamil Nadu, India
    Contribution
    Formal analysis, Investigation, Writing – review and editing
    Competing interests
    None
  5. Anjana Badrinarayanan

    National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India
    Contribution
    Formal analysis, Investigation, Supervision, Writing – review and editing
    Competing interests
    None
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5520-2134
  6. David M Lawson

    Department of Biochemistry and Metabolism, John Innes Centre, Norwich, United Kingdom
    Contribution
    Formal analysis, Investigation, Methodology, Supervision, Writing – original draft
    Competing interests
    None
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7637-4303
  7. Tung BK Le

    Department of Molecular Microbiology, John Innes Centre, Norwich, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Supervision, Visualization, Writing – original draft, Writing – review and editing
    For correspondence
    tung.le@jic.ac.uk
    Competing interests
    None
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4764-8851

Funding

Royal Society (URF\R\201020)

  • Tung B K Le

Royal Society (RG150448)

  • Adam S B Jalal

Biotechnology and Biological Sciences Research Council (BB/P018165/1)

  • Tung B K Le

Biotechnology and Biological Sciences Research Council (BBS/E/J/000PR9791)

  • Ngat T Tran

Wellcome Trust (221776/Z/20/Z)

  • Tung B K Le

Science and Engineering Research Board (2019/003321)

  • Anjana Badrinarayanan

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This study was funded by the Royal Society University Research Fellowship (URF\R\201020), BBSRC grant (BB/P018165/1), and a Wellcome Trust grant (221776/Z/20/Z, to TBKL), and a DST-SERB CRG grant 2019/003321 (to AB). ASBJ’s PhD studentship was funded by the Royal Society (RG150448), and NTT was funded by the BBSRC grant-in-add (BBS/E/J/000PR9791 to the John Innes Centre). We thank Diamond Light Source for access to beamlines I04-1 and I03 under proposals MX13467 and MX18565 with support from the European Community’s Seventh Framework Program (FP7/2007-2013) under Grant Agreement 283570 (BioStruct-X). We thank Stephan Gruber, Martin Thanbichler, and Manuel Osorio-Valeriano for sharing unpublished results.

Senior Editor

  1. Gisela Storz, National Institute of Child Health and Human Development, United States

Reviewing Editor

  1. Anthony G Vecchiarelli, University of Michigan, United States

Reviewer

  1. Anthony G Vecchiarelli, University of Michigan, United States

Publication history

  1. Preprint posted: May 5, 2021 (view preprint)
  2. Received: May 11, 2021
  3. Accepted: August 3, 2021
  4. Version of Record published: August 16, 2021 (version 1)

Copyright

© 2021, Jalal et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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    Lizhi He et al.
    Research Article Updated

    The YAP and TAZ paralogs are transcriptional co-activators recruited to target sites by TEAD proteins. Here, we show that YAP and TAZ are also recruited by JUNB (a member of the AP-1 family) and STAT3, key transcription factors that mediate an epigenetic switch linking inflammation to cellular transformation. YAP and TAZ directly interact with JUNB and STAT3 via a WW domain important for transformation, and they stimulate transcriptional activation by AP-1 proteins. JUNB, STAT3, and TEAD co-localize at virtually all YAP/TAZ target sites, yet many target sites only contain individual AP-1, TEAD, or STAT3 motifs. This observation and differences in relative crosslinking efficiencies of JUNB, TEAD, and STAT3 at YAP/TAZ target sites suggest that YAP/TAZ is recruited by different forms of an AP-1/STAT3/TEAD complex depending on the recruiting motif. The different classes of YAP/TAZ target sites are associated with largely non-overlapping genes with distinct functions. A small minority of target sites are YAP- or TAZ-specific, and they are associated with different sequence motifs and gene classes from shared YAP/TAZ target sites. Genes containing either the AP-1 or TEAD class of YAP/TAZ sites are associated with poor survival of breast cancer patients with the triple-negative form of the disease.

    1. Chromosomes and Gene Expression
    2. Genetics and Genomics
    Natalia Petrenko, Kevin Struhl
    Research Article Updated

    The preinitiation complex (PIC) for transcriptional initiation by RNA polymerase (Pol) II is composed of general transcription factors that are highly conserved. However, analysis of ChIP-seq datasets reveals kinetic and compositional differences in the transcriptional initiation process among eukaryotic species. In yeast, Mediator associates strongly with activator proteins bound to enhancers, but it transiently associates with promoters in a form that lacks the kinase module. In contrast, in human, mouse, and fly cells, Mediator with its kinase module stably associates with promoters, but not with activator-binding sites. This suggests that yeast and metazoans differ in the nature of the dynamic bridge of Mediator between activators and Pol II and the composition of a stable inactive PIC-like entity. As in yeast, occupancies of TATA-binding protein (TBP) and TBP-associated factors (Tafs) at mammalian promoters are not strictly correlated. This suggests that within PICs, TFIID is not a monolithic entity, and multiple forms of TBP affect initiation at different classes of genes. TFIID in flies, but not yeast and mammals, interacts strongly at regions downstream of the initiation site, consistent with the importance of downstream promoter elements in that species. Lastly, Taf7 and the mammalian-specific Med26 subunit of Mediator also interact near the Pol II pause region downstream of the PIC, but only in subsets of genes and often not together. Species-specific differences in PIC structure and function are likely to affect how activators and repressors affect transcriptional activity.