1. Chromosomes and Gene Expression
  2. Microbiology and Infectious Disease
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ParB spreading on DNA requires cytidine triphosphate in vitro

  1. Adam SB Jalal
  2. Ngat T Tran
  3. Tung BK Le  Is a corresponding author
  1. Department of Molecular Microbiology, John Innes Centre, United Kingdom
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Cite this article as: eLife 2020;9:e53515 doi: 10.7554/eLife.53515

Abstract

In all living organisms, it is essential to transmit genetic information faithfully to the next generation. The SMC-ParAB-parS system is widely employed for chromosome segregation in bacteria. A DNA-binding protein ParB nucleates on parS sites and must associate with neighboring DNA, a process known as spreading, to enable efficient chromosome segregation. Despite its importance, how the initial few ParB molecules nucleating at parS sites recruit hundreds of further ParB to spread is not fully understood. Here, we reconstitute a parS-dependent ParB spreading event using purified proteins from Caulobacter crescentus and show that CTP is required for spreading. We further show that ParB spreading requires a closed DNA substrate, and a DNA-binding transcriptional regulator can act as a roadblock to attenuate spreading unidirectionally in vitro. Our biochemical reconstitutions recapitulate many observed in vivo properties of ParB and opens up avenues to investigate the interactions between ParB-parS with ParA and SMC.

Introduction

Faithful chromosome segregation is essential in all domains of life if daughter cells are each to inherit the full set of genetic information. The SMC-ParAB-parS complex is widely employed for chromosome segregation in bacteria (Donczew et al., 2016; Fogel and Waldor, 2006; Gruber and Errington, 2009; Harms et al., 2013; Ireton et al., 1994; Jakimowicz et al., 2002; Kawalek et al., 2018; Lin and Grossman, 1998; Livny et al., 2007; Mohl et al., 2001; Sullivan et al., 2009; Tran et al., 2018; Wang et al., 2017). The centromere parS is the first DNA locus to be segregated following chromosome replication (Lin and Grossman, 1998; Livny et al., 2007; Toro et al., 2008; Lagage et al., 2016). ParB specifically nucleates on parS before spreading outwards to the flanking DNA and bridges/cages DNA together to form a nucleoprotein network in vivo (Breier and Grossman, 2007; Murray et al., 2006; Taylor et al., 2015; Graham et al., 2014; Sanchez et al., 2015; Debaugny et al., 2018; Broedersz et al., 2014; Funnell, 2016). This nucleoprotein complex recruits SMC to disentangle and organize replicated DNA (Gruber and Errington, 2009; Sullivan et al., 2009; Wang et al., 2017; Tran et al., 2017; Minnen et al., 2011). ParB-parS also interacts with an ATPase ParA to power the segregation of replicated chromosomes (Lim et al., 2014; Vecchiarelli et al., 2014; Vecchiarelli et al., 2012; Hwang et al., 2013; Leonard et al., 2005). Engineered strains harboring a nucleation-competent but spreading-defective mutant of parB are either unviable (Mohl et al., 2001) or have elevated number of anucleate cells (Harms et al., 2013; Kawalek et al., 2018; Lin and Grossman, 1998; Lagage et al., 2016; Jecz et al., 2015; Attaiech et al., 2015; Yu et al., 2010; Lee and Grossman, 2006). Despite the importance of spreading for proper chromosome segregation, the mechanism by which a few parS-bound ParB can recruit hundreds more ParB molecules to the vicinity of parS to assemble a high-molecular-weight nucleoprotein complex is not fully understood.

Since the first report in 1995 (Lynch and Wang, 1995), ParB spreading has been observed in vivo by chromatin immunoprecipitation in multiple bacterial species (Tran et al., 2018; Lagage et al., 2016; Breier and Grossman, 2007; Murray et al., 2006; Graham et al., 2014; Rodionov et al., 1999). The nucleation of ParB on parS has also been demonstrated in vitro (Harms et al., 2013; Mohl et al., 2001; Breier and Grossman, 2007; Murray et al., 2006; Sanchez et al., 2015; Chen et al., 2015; Surtees and Funnell, 2001; Ah-Seng et al., 2013); however, parS-dependent ParB spreading has resisted biochemical reconstitution (Taylor et al., 2015; Graham et al., 2014; Fisher et al., 2017; Madariaga-Marcos et al., 2019). Unsuccessful attempts to reconstitute spreading in vitro suggests that additional factors might be missing. Recently, works by Soh et al. (2019) and Osorio-Valeriano et al. (2019) on Bacillus subtilis and Myxococcus xanthus ParB, respectively, showed that ParB binds and hydrolyzes cytidine triphosphate (CTP) to cytidine diphosphate (CDP), and that CTP modulates the binding affinity of ParB to parS (Osorio-Valeriano et al., 2019; Soh et al., 2019). A co-crystal structure of Bacillus ParB with CDP and that of a Myxococcus ParB-like protein (PadC) with CTP showed CTP binding promotes a new dimerization interface between N-terminal domains of ParB subunits (Osorio-Valeriano et al., 2019; Soh et al., 2019). Crucially, Soh et al. (2019) showed by single-molecule imaging and cross-linking assays that Bacillus ParB, in the presence of CTP, forms a self-loading protein clamp at parS and slides away to spread to neighboring DNA (Soh et al., 2019). While reproducing a key result from Easter and Gober (2002) that showed Caulobacter crescentus ParA-ATP dissociated pre-bound ParB from parS (Easter and Gober, 2002), we found that CTP could also modulate the nucleation of Caulobacter ParB on parS. Personal communication with Stephan Gruber and the recent work by Osorio-Valeriano et al. (2019) and Soh et al. (2019) encouraged us to take steps further to investigate the role of CTP for ParB spreading in Caulobacter crescentus.

Here, we reconstitute a parS-dependent ParB spreading on DNA in real-time, using a label-free purified protein from Caulobacter crescentus. Consistent with pioneering works by Soh et al. (2019) and Osorio-Valeriano et al. (2019), we confirm that CTP regulates ParB-DNA interaction. We further provide evidence that the accumulation of Caulobacter ParB requires a closed DNA substrate, and that a DNA-binding transcription factor, TetR, can act as a roadblock to attenuate ParB accumulation unidirectionally in vitro. Our real-time and label-free reconstitution has successfully recapitulated many well-known aspects of ParB behaviors in vivo and might open up avenues to investigate further the roles of the ParB-parS nucleoprotein complex in ensuring faithful chromosome segregation.

Results

CTP reduces the nucleation of Caulobacter ParB on parS

Easter and Gober (2002) reported that ATP-bound Caulobacter ParA dissociated ParB from parS (Easter and Gober, 2002); however, the authors did not control for the effect of ATP alone on ParB-parS binding. NTPs are highly negatively charged and could have affected protein-DNA interactions by binding non-specifically to the often positively charged DNA-binding domain. To determine if ATP or other NTP alone affects ParB-parS interaction, we attached a linear 20 bp biotinylated parS DNA to a streptavidin-coated probe to measure the bio-layer interference (BLI) (Figure 1A). BLI assay monitors wavelength shifts (responses) resulting from changes in the optical thickness of the probe surface during association or dissociation of the analyte (see Materials and Methods). We monitored in real-time interactions between immobilized parS DNA and purified Caulobacter ParB or a premix of ParB + NTP (Figure 1B). Consistent with previous reports (Tran et al., 2018; Figge et al., 2003), Caulobacter ParB bound site-specifically to parS but not to non-cognate DNA (Figure 1—figure supplement 1). In the presence of ATP, GTP, or UTP, we observed a small reduction in ParB-parS binding at steady state regardless of whether Mg2+ was included in binding buffer or not (Figure 1B and Figure 1—figure supplement 2), suggesting that Caulobacter ParB is slightly sensitive to highly negatively charged compounds or to counter-ions (Na+) in NTP solutions. However, we noted that CTP had a pronounced effect on ParB-parS interaction, specifically in the presence of Mg2+ (Figure 1B and Figure 1—figure supplement 2). An increasing concentration of CTP (but not CMP or CDP) gradually reduced the binding of ParB to parS (Figure 1—figure supplement 3). In contrast, neither CTP nor other NTPs affected the binding of another protein-DNA pair, for example, the well-characterized TetR-tetO interaction (Saenger et al., 2000; Figure 1C). On closer inspection, we noted that ParB + CTP slowly dissociated from parS even before the probe was returned to a protein-free buffer (a gradual downward slope between 30th and 150th sec, Figure 1B), suggesting that CTP facilitated ParB removal from a 20 bp parS DNA. To investigate further, we monitored the dissociation rates of pre-bound CTP-free ParB-parS complexes after probes were returned to a protein-free buffer with or without CTP, we found ParB dissociating ~seven times faster in buffer with CTP than in buffer only solution (Figure 1D). Given the short length of a 20 bp parS DNA duplex that has only sufficient room for nucleation, our results suggest that CTP might decrease ParB nucleation on parS or liberates pre-bound ParB from parS site.

Figure 1 with 3 supplements see all
CTP reduces the nucleation of Caulobacter ParB at parS.

(A) The domain architecture of ParB (dark green) and TetR (grey), and their respective DNA-binding sites parS and tetO. Convergent arrows below DNA-binding sites indicate that parS and tetO are palindromic. (B) Bio-layer interferometric (BLI) analysis of the interaction between a premix of 1 μM ParB-His6 dimer ± 1 mM NTP and a 20 bp DNA duplex containing parS. Biotinylated DNA fragments were immobilized onto the surface of a Streptavidin (SA)-coated probe (See Materials and methods). The BLI probe was dipped into a buffer only solution (0–30 s), then to a premix of protein ± NTP (30–150 s: association phase), and finally returned to a buffer only solution (150–270 s: dissociation phase). Sensorgrams were recorded over time. BLI analysis of the interaction between 1 μM TetR-His6 and a 20 bp parS probe was also recorded (a negative control). (C) BLI analysis of the interaction between a premix of 1 μM TetR-His6 ± 1 mM NTP and a 28 bp DNA duplex containing tetO. BLI analysis of the interaction between 1 mM CTP and a 28 bp tetO probe was also recorded (a negative control). (D) BLI analysis of the interaction between 1 μM Caulobacter ParB-His6 (without CTP) and a 20 bp parS DNA. For the dissociation phase, the probe was returned to a buffer only or buffer supplemented with 1 mM CTP. All buffers used for experiments in this figure contained Mg2+. Each BLI experiment was triplicated and a representative sensorgram was presented.

CTP facilitates ParB accumulation on a closed DNA substrate

Next, we investigated the effect of CTP on ParB-DNA interaction by employing a longer 169 bp parS-containing DNA fragment that has been labeled at both 5’ ends with biotin (Figure 2A). Immobilizing a dual biotin-labeled DNA on a streptavidin-coated BLI probe created a DNA substrate where both ends were blocked (a closed DNA) (Onn and Koshland, 2011; Figure 2—figure supplement 1). We monitored the interactions between immobilized DNA and purified Caulobacter ParB in the presence or absence of NTP. In the absence of NTP, we observed the usual nucleation event on parS with 1 µM ParB protein (Figure 2A). We noted that the BLI signal was not as high as with a 20 bp parS probe (Figure 2A) due to a less efficient immobilization of a longer DNA fragment on the BLI probe. Premixing ATP, GTP, or UTP with ParB did not change the sensorgrams markedly (Figure 2A). However, the addition of CTP significantly increased the response by ~12 fold (Figure 2A and Figure 2—figure supplement 2A), suggesting that more ParB associated with a 169 bp parS probe at steady state than by nucleation at parS alone. We observed that DNA-bound ParB was salt sensitive and dissociated readily to the solution when the BLI probe was returned to a low-salt protein-free buffer without CTP (Figure 2A, dissociation phase). However, the dissociation of pre-bound ParB-CTP from DNA was slowed down by ~fivefold if the probe was returned to a buffer supplemented with CTP (Figure 2—figure supplement 2B). The effect on the BLI response was not seen if Mg2+ was omitted (Figure 2—figure supplement 2C), neither did we observe an equivalent increase in response when a 169 bp dual biotin-labeled DNA containing a scrambled parS was employed instead (Figure 2A). Furthermore, we observed that a nucleation-competent but spreading-defective Caulobacter ParB (R104A) (Tran et al., 2018) mutant did not respond to the addition of CTP to the same extent as ParB (WT) (Figure 2B). Our results suggest that CTP is required for the increase in parS-dependent ParB accumulation in vitro. Lastly, we performed BLI experiments for eight additional chromosomal ParB proteins from a diverse set of bacterial species and consistently observed the specific effect of CTP on enhancing ParB association with a closed DNA in vitro (Figure 2—figure supplement 3). It is most likely that ParB-CTP interaction with DNA is conserved among ParB orthologs.

Figure 2 with 3 supplements see all
CTP facilitates ParB association with a closed DNA substrate beyond nucleation.

(A) BLI analysis of the interaction between a premix of 1 μM Caulobacter ParB-His6 ± 1 mM NTP and a 169 bp dual biotin-labeled DNA containing a parS or a scrambled parS site. Interactions between a dual biotinylated DNA and streptavidin (SA)-coated probe created a DNA substrate where both ends were blocked (a closed DNA substrate) (see the schematic of the BLI probes above the sensorgram). (B) Interactions between a nucleation-competent but spreading-defective ParB (R104) variant with a 169 bp parS DNA fragment in the presence or absence of CTP were also recorded. Each BLI experiment was triplicated and a representative sensorgram was presented. (C) A schematic of the pull-down assay and immunoblot analysis of pulled-down Caulobacter ParB-His6. The length of bound DNA is ~2.8 kb. Beads were incubated with ParB protein for five minutes before being pulled down magnetically. All buffers used for experiments in this figure contained Mg2+.

To independently verify the BLI data, we performed an in vitro pull-down of purified His-tagged Caulobacter ParB (Figure 2C). Streptavidin-coated paramagnetic beads were incubated with 2.8 kb dual biotin-labeled DNA fragments containing either parS or scrambled parS sites. Again, a dual biotin-labeled DNA formed a closed substrate on the surface of the beads. DNA-coated beads were incubated with purified Caulobacter ParB either in the presence or absence of NTP before being pulled down magnetically. Pulled-down ParB was released from beads and their protein level was analyzed by an α-His6 immunoblot (Figure 2C). We found ~13–15 fold more pulled-down ParB when CTP was included (Figure 2C). No enrichment was observed when scrambled parS-coated beads were used, confirming that the extensive in vitro association of ParB with DNA is dependent on parS (Figure 2C). Also, consistent with the BLI experiments, no further enrichment of ParB was seen when ATP, GTP or UTP was included (Figure 2C). Furthermore, a ParB (R104A) variant was not enriched in our pull-down assay regardless of whether CTP was present or not (Figure 2C). Altogether, our results suggest that a parS-dependent accumulation of ParB on a closed DNA substrate requires CTP.

A closed DNA substrate is required for an increased ParB association with DNA

Next, we investigated whether a DNA substrate with a free end (an open DNA) can also support ParB accumulation in vitro. The 169 bp dual biotin-labeled DNA was designed with unique BamHI and EcoRI recognition sites flanking the parS site (Figure 3A). To generate an open end on DNA, we immerged the DNA-coated probes in buffers contained either BamHI or EcoRI (Figure 3A–C and Figure 3—figure supplement 1). Before restriction enzyme digestion, we again observed an enhanced ParB association with a closed DNA substrate in the presence of CTP (Figure 3A). After digestion by either BamHI or EcoRI, the inclusion of CTP had no effect on the BLI response, indicating that ParB did not accumulate on an open DNA substrate in vitro (Figure 3B–C). Our conclusion was further supported by results from an experiment in which we added BamHI after ParB + CTP and a closed DNA substrate were preincubated together for 120 s (Figure 3—figure supplement 2). Here, in the presence of BamHI, ParB binding to DNA reduced gradually over 30 min, while it was unaffected if heat-inactivated BamHI was employed instead (Figure 3—figure supplement 2). Lastly, consistent with BLI experiments, our pull-down assay also showed that ParB-CTP failed to accumulate when a 2.8 kb dual biotin-labeled DNA was linearized by HindIII digestion (Figure 3D).

Figure 3 with 2 supplements see all
A closed DNA substrate is required for an increased association of ParB with DNA.

(A) BLI analysis of the interaction between a premix of 1 μM Caulobacter ParB-His6 ± 1 mM CTP and a 169 bp dual biotin-labeled parS DNA. (B) Same as panel A but immobilized DNA fragments have been restricted with BamHI before BLI analysis. (C) Same as panel A but immobilized DNA fragments have been restricted with EcoRI before BLI analysis. Schematic of DNA fragments with the relative positions of parS and restriction enzyme recognition sites are shown above the sensorgram. Each BLI experiment was triplicated and a representative sensorgram was presented. (D) A schematic of the pull-down assay and immunoblot analysis of pulled-down Caulobacter ParB-His6. For lanes 3 and 4, DNA-bound beads were incubated with HindIII to linearize bound DNA. Samples in lanes 1–4 were loaded on the same gel, the immunoblot was spliced together for presentation purposes. All buffers used for experiments in this figure contained Mg2+.

Next, we wondered if a tight protein-DNA binding could cap the open end of DNA, thereby mimicking a closed DNA substrate and restoring ParB accumulation. To investigate this possibility, we constructed a 170 bp dual biotin-labeled DNA fragment that contains a single parS site, a tetO operator, and flanking restriction enzyme recognition sites for EcoRI and BamHI (Figure 4A). With this closed DNA substrate, we observed an enhanced ParB association with DNA in the presence of CTP (Figure 4A). Again, we generated a free DNA end via restriction enzyme digestion. Consistent with previous experiments with a restricted 169 bp DNA probe, the addition of ParB + CTP had no effect on the BLI response (Figure 4B). However, it can be partially rescued by incubating a BamHI-restricted DNA probe with a premix of ParB + CTP + TetR (Figure 4B). We reason that TetR binding at tetO capped the open DNA end, essentially generated a closed DNA substrate. Our conclusion was further supported by results from an experiment in which a premix of ParB + CTP + TetR was tested against an EcoRI-restricted DNA instead (Figure 4C). Here, we did not observe an enhanced association of ParB with DNA even when TetR was included, most likely because of a persistent open DNA end that could not be blocked by TetR-tetO binding (Figure 4C). Finally, we observed that the TetR could attenuate ParB-CTP accumulation on a closed DNA substrate (magenta line vs. orange line, Figure 4A). This blocking effect is specific to TetR-tetO binding since the addition of anhydrotetracycline (ahTc), a negative effector of TetR (Saenger et al., 2000), allowed ParB-CTP to regain its accumulation on DNA (Figure 4—figure supplement 1). Overall, the ability of a DNA-bound TetR to act as a roadblock in vitro is consistent with previous ChIP-seq data that showed DNA-binding proteins or RNA polymerases could block or attenuate ParB spreading unidirectionally in vivo (Breier and Grossman, 2007; Murray et al., 2006; Sanchez et al., 2015; Rodionov et al., 1999).

Figure 4 with 1 supplement see all
TetR-tetO binding restores ParB association with an open DNA substrate.

(A) BLI analysis of the interaction between a premix of 1 μM Caulobacter ParB-His6 ± 1 mM CTP ± 1 μM TetR-His6 and a 170 bp dual biotin-labeled DNA containing a parS site. (B) Same as panel A but immobilized DNA fragments have been restricted with BamHI before BLI analysis. (C) Same as panel A but immobilized DNA fragments have been restricted with EcoRI before BLI analysis. Schematic of DNA fragments together with the relative positions of parS, tetO, and restriction enzyme recognition sites are shown above the sensorgram. Each BLI experiment was triplicated and a representative sensorgram was presented. All buffers used for experiments in this figure contained Mg2+.

parS DNA increases the CTP binding and hydrolysis rate of Caulobacter ParB

Recently, Myxococcus and Bacillus ParB were shown to bind and hydrolyze CTP (Osorio-Valeriano et al., 2019; Soh et al., 2019). Our in vitro results so far also hint at CTP binding directly to Caulobacter ParB. By employing a membrane-spotting assay (DRaCALA), we showed that Caulobacter ParB binds to radiolabeled CTP in the presence of parS DNA (Figure 5A). An excess of unlabeled CTP, but no other NTPs, could compete with radioactive CTP for binding to Caulobacter ParB (Figure 5B), suggesting that Caulobacter ParB does not bind other NTPs. The CTP binding of ParB was reduced when a non-cognate DNA site (NBS) (Wu and Errington, 2004) was used instead of parS (Figure 5A). We also failed to detect CTP binding in our DRaCALA assay or by isothermal titration calorimetry when DNA was omitted. Nevertheless, we robustly detected CTP hydrolysis to CDP and inorganic phosphate when Caulobacter ParB and CTP were included, albeit at a very low rate of ~0.4 CTP molecules per ParB per hour (Figure 5C). A background level of inorganic phosphate was observed when Caulobacter ParB was incubated with ATP, GTP, or UTP (Figure 5C). Crucially, the addition of a 22 bp parS DNA, but not a non-cognate 22 bp NBS DNA, increased CTP turnover rate sevenfold to ~3 CTP molecules per ParB per hour (Figure 5C). Lastly, the CTP hydrolysis was reduced to the background in the nucleation-competent but spreading-defective ParB (R104A) variant (Figure 5C). Altogether, our data suggest that parS DNA stimulates Caulobacter ParB to bind and hydrolyze CTP.

parS DNA increases the CTP binding and hydrolysis rate by Caulobacter ParB.

(A–B) CTP binding as monitored by DRaCALA 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 alone. The starting concentration of proteins used in panel A was 25 μM. The same concentration of radioactive CTP, unlabeled CTP, and DNA was used in experiments shown in panels A and B. (C) A continuous monitoring of inorganic phosphate (Pi) released by recording absorbance at 360 nm overtime at 25°C. The rates of CTP hydrolysis were inferred from a Pi standard. The NTP hydrolysis of Caulobacter ParB was also monitored in the presence of ATP, GTP, or UTP, with a 22 bp parS DNA duplex or a non-cognate 22 bp NBS DNA duplex (a DNA-binding site of Noc protein Wu and Errington, 2004). The nucleation-competent but spreading-defective ParB (R104A) mutant did not hydrolyze CTP in the presence of parS DNA. All buffers used for experiments in this figure contained Mg2+.

ParB accumulation on DNA is unstable in the presence of a non-hydrolyzable analog CTPγS

Next, we investigated the role of CTP hydrolysis by following ParB-parS interaction in the presence of a non-hydrolyzable analog CTPγS (Figure 6A and Figure 6—figure supplement 1). ParB was preincubated with CTP or CTPγS for 1 to 60 min before binding to a 169 bp closed parS DNA substrate (Figure 6A). We observed that Caulobacter ParB could accumulate on DNA in the presence of CTPγS, but in contrast to when CTP was employed, a longer preincubation time between ParB and CTPγS gradually reduced ParB accumulation on DNA (Figure 6A). Our results suggest the possibility that CTPγS, in the absence of parS DNA, converts apo-ParB in solution to a nucleation-incompetent form over time. Our observation is reminiscent of a time-course experiment in which CTPγS efficiently promoted the engagement between N-terminal domains of Bacillus ParB in the absence of parS DNA (Soh et al., 2019). The engagement of N-terminal domains was shown to convert Bacillus ParB from an open to a closed protein clamp (Soh et al., 2019). If not already bound on DNA, the closed form of ParB presumably cannot nucleate/load onto parS due to its now inaccessible DNA-binding domain (Soh et al., 2019). We wondered if CTPγS also catalyzed the N-domain engagement in Caulobacter ParB in the absence of parS DNA. To investigate this possibility, we employed site-specific cross-linking of a purified Caulobacter ParB (Q35C C297S) variant by a sulfhydryl-to-sulfhydryl crosslinker bismaleimidoethane (BMOE) (Figure 6B). A lone cysteine residue on the native ParB was first mutated to serine to create a cysteine-less ParB (C297S) variant, then a glutamine to cysteine mutation was engineered at position 35 at the N-terminal domain to create ParB (Q35C C297S). Both ParB (C297S) and ParB (Q35C C297S) were competent at spreading in the presence of CTP (Figure 6—figure supplement 2A). The cross-linking of ParB (Q35C C297S) was also enhanced by CTP and CTPγS but not by other NTPs, consistent with the specific role of CTP in ParB spreading (Figure 6—figure supplement 2B). We performed a time-course cross-linking of ParB (Q35C C297S) + CTP or CTPγS in the absence of parS DNA (Figure 6B). As shown in Figure 6B, CTPγS was twice as efficient as CTP in promoting the cross-linked form between N-terminal domains of Caulobacter ParB. The rapid increase in the nucleation-incompetent closed form of Caulobacter ParB might explain the overall reduction in accumulation overtime when ParB was preincubated with CTPγS (Figure 6A).

Figure 6 with 2 supplements see all
ParB accumulation on DNA is unstable in the presence of a non-hydrolyzable analog CTPγS.

(A) (Left panel) BLI analysis of the interaction between a premix of 1 μM Caulobacter ParB-His6 ± 1 mM CTP or CTPγS and a 169 bp dual biotin-labeled parS DNA. Purified ParB was preincubated with CTP or CTPγS for 1, 5, 15, 30, or 60 min before BLI analysis. (Right panel) Quantification of ParB-DNA interaction at the end of each association phase (150th sec). Error bars represent SD from three replicates. (B) (Left panel) Time course of Caulobacter ParB (Q35C C297S) cross-linking with CTP or CTPγS in the absence of parS DNA. Purified ParB-His6 (Q35C C297S) (10 µM) were preincubated with 1 mM CTP or CTPγS for 1, 5, 10, 15, or 30 min before BMOE was added. X indicates a cross-linked form of ParB. (Right panel) Quantification of the cross-linked (X) fractions. Error bars represent SD from three replicates. (C) BLI analysis of the interaction between a premix of 1 μM Caulobacter ParB-His6 ± 1 mM CTP or CTPγS and a 169 bp dual biotin-labeled parS DNA. CTP or CTPγS was added to purified ParB, and the mixture was immediately used for BLI experiments. All buffers used for experiments in this figure contained Mg2+.

To further investigate the effect of CTPγS on ParB spreading on a longer time scale, we extended the association phase between 169 bp parS DNA and a freshly prepared premix of ParB + CTP or CTPγS from 120 s to 60 min (Figure 6C). In the presence of CTP, the reaction reached steady state after 120 s and remained stable for the duration of the association phase (Figure 6C). However, in the presence of CTPγS, ParB accumulation rapidly reached the maximal level after 200 s, then declined slowly over 60 min. We suggest that DNA-bound ParB-CTPγS complexes gradually dissociated from DNA into solution (possibly via a transient clamp opening or CTPγS leaving its weak binding pocket rather than by hydrolysis) but were not replenished by a new cycle of nucleation-spreading-dissociation because over time most ParB-CTPγS in solution was in a nucleation-incompetent closed form. Taken together, we suggest that CTP hydrolysis is not required for parS-bound ParB to escape the nucleation site to spread, but possibly contributes to maintaining the stability of spreading by recycling ParB.

Discussion

In this work, we report that a small molecule (CTP) is required to enable Caulobacter ParB proteins (as well as eight other chromosomal ParB proteins, Figure 2—figure supplement 3) to spread in vitro. Recently, Soh et al. (2019) observed that F-plasmid and P1-plasmid ParB proteins also bind and hydrolyze CTP (Soh et al., 2019). Hence, it is most likely that the effect of CTP on ParB spreading is universal among plasmid and chromosomal ParB orthologs. A classical mutant whose arginine-rich patch (G101ERRxR) has been mutated to alanine for example ParB (R104A) (Lin and Grossman, 1998) was not responsive to CTP, suggesting that CTP is bound to the N-terminal domain of Caulobacter ParB. Indeed, Soh et al. (2019) reported a co-crystal structure that showed CDP binding to the arginine-rich patch at the N-terminal domain of Bacillus ParB (CTP was hydrolyzed to CDP during crystallization) (Soh et al., 2019). Osorio-Valeriano et al. (2019) also showed a similar binding pocket of CTP at the N-terminal domain of Myxococcus ParB by hydrogen-deuterium exchange mass spectrometry (Osorio-Valeriano et al., 2019). Intriguingly, a co-crystal structure of a Helicobacter pylori ParB-parS complex, together with the in vitro magnetic-tweezer and single-molecule TIRF microscopy-based experiments with Bacillus ParB showed that the N-terminal domain can oligomerize to bridge DNA together without the need of an additional ligand (Taylor et al., 2015; Graham et al., 2014; Chen et al., 2015; Fisher et al., 2017). There might be two different modes of action of ParB on DNA: one for bridging DNA together (that does not require CTP) and another for the lateral spreading of ParB on DNA (that requires CTP). Investigating the relative contribution of these two different modes of action to chromosome segregation in vivo is an important challenge for the future.

The requirement of a DNA substrate with blocked ends for ParB accumulation in vitro is suggestive of a lateral ParB diffusion along the DNA that is ParB can escape by running off a free DNA end (Figure 7). Inside cells, the spreading and bridging/caging of ParB have been inferred from the compact foci of fluorescently labeled ParB (Harms et al., 2013; Lagage et al., 2016; Sanchez et al., 2015; Thanbichler and Shapiro, 2006; Kusiak et al., 2011; Lin et al., 1997; Glaser et al., 1997; Erdmann et al., 1999), presumably resulting from the concentration of fluorescent signal to a defined location in the cytoplasm. Nucleation-competent but spreading-defective ParB mutants formed no or very diffusive foci in vivo (Graham et al., 2014; Song et al., 2017). Recently, it has been observed that an artificially engineered double-strand break ~8 kb away from parS did not cause a dissolution of ParB foci in Caulobacter cells (Badrinarayanan et al., 2015). This result seemingly contradicted our findings that Caulobacter ParB spreading in vitro requires a closed DNA. However, we reason that the abundant DNA-bound transcription factors and RNA polymerases in vivo act as roadblocks to minimize ParB runoff. This barricading effect has been recapitulated in our experiments with TetR, a tight DNA-binding transcriptional regulator (Figure 4).

A model for Caulobacter ParB nucleation and spreading.

(A) Caulobacter ParB nucleation at parS. CTP (orange) reduces Caulobacter ParB (dark green) nucleation at parS (magenta box), presumably by inducing conformational changes that are incompatible with a site-specific parS binding (Soh et al., 2019). Only apo- or CDP-bound ParB can nucleate on parS. ParB hydrolyzes CTP at a faster rate in the presence of parS. The domain architecture of ParB is also shown: NTD: N-terminal domain, DBD: DNA-binding domain, and CTD: C-terminal domain. (B) Caulobacter ParB escapes from the nucleation site parS. Apo-ParB at parS binds CTP and slides laterally away from the nucleation site parS while still associating with DNA. (C) Caulobacter ParB sliding and spreading on DNA. CTP-bound ParBs diffuse from the nucleation site parS and can run off the free DNA end unless they are blocked by DNA-bound roadblocks such as transcriptional regulators for example TetR. CTP hydrolysis is not required for ParB to escape from the nucleation parS site but might contribute to ParB recycling. It is not yet known whether both CTP molecules on a ParB dimer are concertedly hydrolyzed/dissociated for ParB to escape from the chromosome or a heterodimer state of ParB with a single CTP bound also exists in vivo.

Our results so far suggest three distinct stages of Caulobacter ParB-DNA interactions in vitro:

Stage 1: ParB nucleates on parS (Figure 7A). Results from experiments in Figure 1 indicate that CTP modulates ParB nucleation on a parS site. Soh et al. (2019) reported that CTP-bound ParB could form a closed protein clamp even in the absence of parS DNA, albeit not energetically favorable (Soh et al., 2019). The DNA-binding domain of a closed ParB clamp would be inaccessible to DNA, especially to a closed DNA substrate. It might be that only apo-ParB or a transiently formed CDP-bound ParB (from CTP hydrolysis) are able to nucleate on parS (Figure 7A). Supporting this interpretation, preincubation of Caulobacter ParB with a non-hydrolyzable analog CTPγS promoted efficiently the cross-linked form of ParB and subsequently reduced ParB nucleation and spreading on DNA (Figure 6). Initially, we were surprised by a weak CTP binding of Bacillus, Caulobacter, and Myxococcus ParBs (Osorio-Valeriano et al., 2019; Soh et al., 2019), however, this might be advantageous for the cells as a fraction of intracellular apo-ParB will remain to nucleate on parS.

Stage 2: Nucleated ParB escapes from parS (Figure 7B–C). We showed that Caulobacter ParB-parS complex binds CTP, and this facilitates ParB dissociation from parS (Figure 1D). It was reported that the DNA-binding domain in Bacillus ParB-CDP co-crystal structure is incompatible with parS binding (Soh et al., 2019) and this might enable ParB to escape from a high-affinity nucleation site to non-specific flanking DNA. Our observation of a low BLI response with an open DNA (Figure 3 and Figure 4) implies that ParB proteins dissociate off the free end well before the next ParB escapes from the parS nucleation site. We suggest that the transition from a parS-bound ParB to a spreading ParB might be the rate-limiting step.

Stage 3: ParB spreads or diffuses to non-specific DNA flanking parS. Our observation that Caulobacter ParB did not accumulate on an open DNA suggests that Caulobacter ParB diffuses laterally along the DNA. Similarly, cross-linking experiments on Bacillus ParB (Soh et al., 2019) proposed that the ParB-CTP complex forms a sliding clamp that moves along the DNA (Soh et al., 2019). From our experiments with CTPγS (Figure 6) and consistent with Soh et al. (2019), we suggest that the diffusive Caulobacter ParB along the DNA is CTP bound. The low CTP hydrolysis rate of Caulobacter ParB (~3 CTP molecules per ParB per hour) while ParB spreading could be observed by BLI within minutes also lends support to the interpretation that the diffusive spreading form of Caulobacter ParB is most likely CTP-bound (Figure 7B–C). For Bacillus and Caulobacter ParB, CTP hydrolysis is not required for ParB to escape from the nucleation site (Soh et al., 2019). Caulobacter ParB could still spread on a DNA when incubated with a non-hydrolyzable CTPγS (Figure 6), even though spreading, at least in vitro, was less stable over time in comparison to when CTP was employed (Figure 6C). As suggested for Bacillus ParB (Soh et al., 2019) and supported by our results with Caulobacter ParB, CTP hydrolysis might contribute to ParB recycling instead. It is possible that ParB recycling in vivo is achieved via several routes: (i) CTP dissociation from its weak binding pocket, (ii) CTP hydrolysis, (iii) a possible enhanced CTP dissociation/hydrolysis via collisions with DNA-bound roadblocks, or (iv) a transient clamp opening even when ParB is CTP bound. Additional work is required to investigate the dynamics of ParB clamp opening/closing, and whether both CTP molecules on opposite subunits of a ParB dimer are concertedly hydrolyzed/dissociated for ParB to escape from the chromosome or a heterodimer state of ParB with a single CTP bound exists in vivo.

Final perspectives

In this work, we showed the enhancing effect of CTP on Caulobacter ParB accumulation on DNA and further demonstrated that ParB spreading requires a closed DNA substrate and that a DNA-binding transcriptional regulator can act as a roadblock to attenuate spreading unidirectionally in vitro. Our real-time and label-free reconstitution of ParB spreading has successfully recapitulated many well-known aspects of ParB behaviors and is consistent with pioneering works by Soh et al. (2019) and Osorio-Valeriano et al. (2019). Beyond the biological significance of the findings, our label-free approaches to biochemical reconstitution obviate the often time-consuming and challenging task of site-specifically labeling proteins with fluorophores/chemical crosslinkers without affecting the function of proteins. Here, we have demonstrated the medium-throughput capability of our methodology by investigating the effect of CTP on the spreading of eight additional chromosomal ParB proteins. The ease and medium-throughput manner of our methodology is likely to facilitate future works by the community. Important areas of research in the future are to investigate (i) the effect of ParB-CTP spreading on the supercoiling state of the DNA and vice versa, and (ii) how ParA and SMC interact with ParB-CTP in vivo to organize and faithfully segregate replicated chromosomes to each daughter cell.

Materials and methods

Key resources table
Reagent type (species)
or resource
DesignationSource or referenceIdentifiersAdditional
information
Strain, strain background (Escherichia coli)Rosetta (DE3)MerckCat# 70954Electro-competent cells
Recombinant DNA reagentSee Supplementary file 1This paperSee Supplementary file 1
Sequence-based reagentSee Supplementary file 1This paperSee Supplementary file 1
AntibodyAnti-6xHis tag antibody (HRP) (Rabbit polyclonal)AbcamCat# ab1187Western blot (1:5,000)
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 (SA) biosensorsMolecular DevicesCat# 18–5019
Commercial assay or kitHisTrap High Performance columnGE HealthcareCat# GE17524801
Commercial assay or kitHiTrap Heparin High Performance columnGE HealthcareCat# GE17040601
Commercial assay or kitHiLoad 16/600 Superdex 75 pg columnGE HealthcareCat# GE28989333
Commercial assay or kitDynabeads MyOne Streptavidin C1ThermoFisherCat# 65001
Commercial assay or kitAmersham Protran supported Western blotting membranes, nitrocelluloseGE HealthcareCat# GE10600016pore size 0.45 μm, for DRaCALA assay
Peptide, recombinant proteinBamHI-HFNEBCat# R3136S20,000 units/mL
Peptide, recombinant proteinBamHINEBCat# R0136S20,000 units/mL
Peptide, recombinant proteinEcoRI-HFNEBCat# R3101S20,000 units/mL
peptide, recombinant proteinHindIII-HFNEBCat# R3104S20,000 units/mL
Peptide, recombinant proteinExonuclease VIINEBCat# M0379S10,000 units/mL
Peptide, recombinant proteinExonuclease T7NEBCat# M0263S10,000 units/mL
Chemical compound, drugBenzonase nucleaseSigma-AldrichCat# E1014
Chemical compound, drugNTPThermoFisherCat# R0481100 mM solution
Chemical compound, drugCTPγSJena BioscienceA gift from S. Gruber and
custom synthesis (purity ≥ 96%)
Chemical compound, drugCMP-PCPJena BioscienceCat# NU-254
Chemical compound, drugP32-α-CTPPerkin ElmerCat# BLU008H250UC3000 Ci/mmol, 10 mCi/ml, 250 µCi
Chemical compound, drugAnhydrotetracycline hydrochloride (ahTc)AbcamCat# ab145350Dissolved in ethanol
Chemical compound, drugBismaleimidoethane (BMOE)ThermoFisherCat# 22323Dissolved in DMSO
Software, algorithmBLItz ProMolecular DevicesCat# 50–0156Version 1.2 https://www.moleculardevices.com/
Software, algorithmRR Foundation for Statistical ComputingRRID: SCR_001905Version 3.2.4
Software, algorithmImage Studio LiteLI-COR BiosciencesRRID: SCR_013715Version 5.2
Software, algorithmExcel 2016MicrosoftRRID: SCR_016137

Protein overexpression and purification

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Full-length Caulobacter ParB (WT) and ParB (R104A) were purified as described previously (Tran et al., 2018). Briefly, pET21b::ParB-His6 (WT or R104A) was introduced into E. coli Rosetta (DE3) competent cells (Merck). A 10 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 to cool to 4°C before isopropyl-β-D-thiogalactopyranoside (IPTG) was added to a final concentration of 1 mM. The culture was shaken for 3 hr at 30°C before cells were harvested 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 (Sigma Aldrich), 1 mg of lysozyme (Sigma Aldrich), and an EDTA-free protease inhibitor tablet (Roche). The pelleted cells were then lyzed by sonication. The cell debris was removed via centrifugation at 28,000 g for 30 min, and the supernatant was filtered through a 0.45 µm sterile filter (Sartorius Stedim). The protein was then loaded into a 1 mL HisTrap column (GE Healthcare) that had been equilibrated with buffer A [100 mM Tris-HCl pH 8.0, 300 mM NaCl, 10 mM Imidazole, and 5% glycerol]. Protein was eluted from the column using an increasing (10 mM to 500 mM) imidazole gradient in the same buffer. ParB-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 equilibrated with 100 mM Tris-HCl pH 8.0, 25 mM NaCl, and 5% glycerol. Protein was eluted from the Heparin column using an increasing (25 mM to 1 M NaCl) salt gradient in the same buffer. ParB that was used for EnzChek Phosphate assay and DRaCALA was further polished via a gel-filtration column. To do so, purified ParB was concentrated by centrifugation in an Amicon Ultra-15 3 kDa cut-off spin filters (Merck) before being loaded into a Superdex 75 gel filtration column (GE Healthcare). The gel filtration column was pre-equilibrated with 100 mM Tris-HCl pH 8.0, 250 mM NaCl, and 1 mM MgCl2.

C-terminally His-tagged TetR (class B, from Tn10) were expressed from E. coli Rosetta (DE3) harboring a pET21b::TetR-His6 plasmid (Supplementary file 1). TetR-His6 were purified via a one-step Ni-affinity column using the exact buffers as employed for the purification of Caulobacter ParB-His6.

C-terminally His-tagged ParB (C297S) and ParB (Q35C C297S) were expressed from E. coli Rosetta (DE3) harboring a pET21b::ParB-His6 (C297S) or (Q35C C297S) plasmid (Supplementary file 1). ParB-His6 (C297S) and (Q35C C297S) were purified via a one-step Ni-affinity column using the exact buffers as employed for the purification of Caulobacter ParB-His6. ParB (Q35C C297S) stock solution was supplemented with TCEP (1 mM final concentration) before being flash-frozen in liquid nitrogen.

N-terminally His-tagged MBP-tagged ParB (orthologous proteins from various bacterial species) were expressed from E. coli Rosetta (DE3) harboring pET-His-MBP-TEV-DEST::ParB plasmids. His6-MBP-ParB proteins were purified via a one-step Ni-affinity column as described previously (Jalal et al., 2019).

Different batches of proteins were purified by A.S.B.J and N.T.T, and are consistent in all assays used in this work. Both biological (new sample preparations from a fresh stock aliquot) and technical (same sample preparation) replicates were performed for assays described in this study.

Construction of pET21b::ParB-His6 (C297S and Q35C C297S)

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DNA containing the codon-optimized coding sequence of ParB (C297S or Q35C C297S) was chemically synthesized (gBlocks dsDNA fragments, IDT). These gBlocks fragments and a NdeI-HindIII-digested pET21b backbone were assembled using a 2x Gibson master mix (NEB). Two and a half µL of each fragment at equimolar concentration was added to 5 µL 2x Gibson master mix (NEB), and the mixture was incubated at 50°C for 60 min. Five µL was used to transform chemically competent E. coli DH5α cells. Gibson assembly was possible due to a 23 bp sequence shared between the NdeI-HindIII-cut pET21b backbone and the gBlocks fragments. These 23 bp regions were incorporated during the synthesis of gBlocks fragments. The resulting plasmids were sequence verified by Sanger sequencing (Eurofins, Germany).

Construction of pET21b::TetR-His6

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DNA containing the coding sequence of TetR (class B, from Tn10) was chemically synthesized (gBlocks dsDNA fragments, IDT). This gBlocks fragment and a NdeI-HindIII-digested pET21b backbone were assembled together using a 2x Gibson master mix (NEB). Gibson assembly was possible due to a 23 bp sequence shared between the NdeI-HindIII-cut pET21b backbone and the gBlocks fragment. These 23 bp regions were incorporated during the synthesis of gBlocks fragments. The resulting plasmids were sequence verified by Sanger sequencing (Eurofins, Germany).

Construction of pENTR::ParB orthologs

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The coding sequences of ParB orthologs were chemically synthesized (gBlocks dsDNA fragments, IDT) and cloned into pENTR-D-TOPO backbone (Invitrogen) by Gibson assembly (NEB). The resulting plasmids were sequence verified by Sanger sequencing (Eurofins, Germany).

Construction of pET-His-MBP-TEV-DEST::ParB orthologs

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The parB genes were recombined into a Gateway-compatible destination vector pET-His-MBP-TEV-DEST (Jalal et al., 2019) via an LR recombination reaction (ThermoFisher). For LR recombination reactions: 1 µL of purified pENTR::parB (~100 ng/µL) was incubated with 1 µL of the destination vector pET-His-MBP-TEV-DEST (~100 ng/µL), 1 µL of LR Clonase II enzyme mix, and 2 µL of water in a total volume of 5 µL.

Construction of DNA substrates for BLI assays

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All DNA constructs (Supplementary file 1) were designed in VectorNTI (ThermoFisher) and were chemically synthesized (gBlocks dsDNA fragments, IDT). All linear DNA constructs were designed with M13F and M13R homologous regions at each end. To generate a dual biotin-labeled DNA substrate, PCR reactions were performed using a 2x GoTaq PCR master mix (Promega), biotin-labeled M13F and biotin-labeled M13R primers, and gBlocks fragments as template. PCR products were resolved by electrophoresis and gel purified.

Measurement of protein-DNA interaction by bio-layer interferometry (BLI)

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Bio-layer interferometry experiments were conducted using a BLItz system equipped with Dip-and-Read Streptavidin (SA) 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% 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 or without 1 μM TetR or NTPs at various concentrations) for 120 s or 60 min (Figure 6A). 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, 1 mM MgCl2, and 0.005% Tween 20] for 5 min to remove bound ParB.

For experiments where a closed DNA was cleaved to generate a free DNA end, DNA-coated tips were dipped into 300 μL of cutting solution [266 μL of water, 30 μL of 10x CutSmart buffer (NEB), and 4 μL of EcoRI-HF or BamHI-HF restriction enzyme (20,000 units/mL)] for 30 min at 37°C.

For experiments described in Figure 1—figure supplement 2, MgCl2 was omitted from all binding and protein storage buffers.

For experiments described in Figure 3—figure supplement 2, bio-layer interferometry assays were performed using 1x NEB 3.1 buffer [100 mM NaCl, 50 mM Tris-HCl pH 7.9, 10 mM MgCl2, 100 µg/ml BSA] instead of the low-salt binding buffer [100 mM NaCl, 10 mM Tris-HCl pH 8.0, 1 mM MgCl2, 0.005% Tween20]. After incubating 1 μM ParB and 1 mM CTP with a 169 bp parS-coated probe for 120 s, 4 μL of BamHI (20,000 units/mL) or heat-inactivated BamHI was added to a 300 μL reaction to start digesting bound DNA. The reaction was monitored for an additional 30 min. BamHI was inactivated by heat at 65°C for 30 min.

For experiments described in Figure 4—figure supplements 1, 3 μM of anhydrotetracycline (ahTc) was used to remove bound TetR from DNA. After incubating 1 μM ParB and 1 mM CTP ±1 μM TetR with a 170 bp parS-coated probe for 120 s, ahTc (dissolved in ethanol) or ethanol alone was added to a 300 μL reaction to the final concentration of 3 μM or 0.01%, respectively. The reaction was monitored for an additional 120 s.

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, the standard deviation of triplicated sensorgrams is less than ten percent, and a representative sensorgram was presented in each figure.

To verify that dual biotin-labeled DNA fragments formed a closed substrate on the surface of the BLI probe, we performed a double digestion with Exonuclease T7 and Exonuclease VII (NEB) (Figure 2—figure supplement 1). DNA-coated tips were dipped into 300 μL of cutting solution [266 μL of water, 30 μL of 10x RE buffer 4 (NEB), 2 μL of exonuclease T7 (10,000 units/mL) and 2 μL of exonuclease VII (10,000 units/mL)] for 30 min at 25°C. Tips were then cut off from the plastic adaptor (Figure 2—figure supplement 1) and immerged into a 1x GoTaq PCR master mix [25 μL water, 25 μL 2x GoTaq master mix, 0.5 μL of 100 μM M13F oligo, and 0.5 μL of 100 μM M13R oligo]. Ten cycles of PCR were performed, and the PCR products were resolved on 2% agarose gels (Figure 2—figure supplement 1).

NTP (stock concentration: 100 mM) used in BLI assays was purchased from ThermoFisher. CTPγS (stock concentration: 90 mM) was a generous gift from Stephan Gruber and Young-Min Soh. CTPγS was also custom-synthesized and re-purified to 96% purity (Jena Bioscience). Another non-hydrolyzable analog CMP-PCP (Jena Bioscience) was unsuitable for our assays as Caulobacter ParB does not bind CMP-PCP (Figure 6—figure supplement 1).

Construction of DNA substrates for pull-down assays

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A 260 bp DNA fragment containing Caulobacter parS sites (genomic position: 4034789–4035048) (Tran et al., 2018) or scrambled parS sites were chemically synthesized (gBlocks fragments, IDT). These DNA fragments were subsequently 5’ phosphorylated using T4 PNK enzyme (NEB), then cloned into a SmaI-cut pUC19 using T4 DNA ligase (NEB). The two resulting plasmids are pUC19::260bp-parS and pUC19::260bp-scrambled parS (Supplementary file 1). These plasmids were sequence verified by Sanger sequencing (Eurofins, Germany). To generate dual biotin-labeled DNA substrates, we performed PCR using a pair of biotinylated primers: around_pUC19_F and around_pUC19_R, and either pUC19::260bp-parS or pUC19::260bp-scrambled parS as a template. Phusion DNA polymerase (NEB) was employed for this round-the-horn PCR reaction. The resulting ~2.8 kb linear DNA fragments were gel-purified and eluted in 50 µL of autoclaved distilled water.

Pull-down assays

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Paramagnetic MyOne Streptavidin C1 Dyna beads (ThermoFisher) were used for pull-down assays. Thirty µL of beads were washed twice in 500 µL of high-salt wash buffer [100 mM Tris-HCl pH 8.0, 1 M NaCl, 1 mM MgCl2, and 0.005% Tween 20] and once in 100 µL binding buffer [100 mM Tris-HCl pH 8.0, 100 mM NaCl, 1 mM MgCl2, and 0.005% Tween 20] by repeating a cycle of resuspension and pull-down by magnetic attraction. Five µL of ~50 nM dual biotin-labeled DNA substrate was incubated with 30 µL of beads in 100 µL binding buffer for 30 min at room temperature. The reaction was occasionally mixed by pipetting up and down several times. Afterward, DNA-coated beads were washed once in 500 µL high-salt buffer [100 mM Tris-HCl pH 8.0, 1000 mM NaCl, 1 mM MgCl2, and 0.005% Tween 20] and once in 500 µL of binding buffer. Finally, DNA-coated beads were resuspended in 300 µL of binding buffer. Ninety-six µL of the resuspended beads were used for each pull-down assay. Four µL of Caulobacter ParB-His6 (WT) or (R104A) (stock concentration: 25 µM) were added to 96 µL of suspended beads. NTPs were either omitted or added to the suspended beads to the final concentration of 1 mM. The mixture was pipetted up and down several times and was left to incubate at room temperature for 5 min. Beads were then pulled down magnetically and unwanted supernatant discarded. DNA-coated beads (now with bound protein) were then washed once with 500 µL of binding buffer and once with 100 µL of the same buffer. The unwanted supernatant was discarded, and the left-over beads were resuspended in 30 µL of 1x SDS-PAGE sample buffer. Each experiment was triplicated, and a representative immunoblot was presented.

Immunoblot analysis

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For immunoblot analysis, magnetic beads were resuspended directly in 1x SDS sample buffer, then heated to 42°C for 15 min before loading to 12% Novex Tris-Glycine SDS-PAGE gels (ThermoFisher). The eluted protein was resolved by electrophoresis at 150 V for 60 min. Resolved proteins were transferred to polyvinylidene fluoride membranes using the Trans-Blot Turbo Transfer System (BioRad) and probed with 1:5000 dilution of α-His6 HRP-conjugated antibody (Abcam). Blots were imaged and analyzed using an Amersham Imager 600 (GE Healthcare) and Image Studio Lite version 5.2 (LI-COR Biosciences). The band intensities were quantified for lanes 5 and 6 (Figure 2C), and the range of fold difference between replicates was reported.

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

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Purified Caulobacter ParB-His6 or TetR-His6 (final concentration: 25 µM) were incubated with 5 nM radiolabeled P32-α-CTP (Perkin Elmer), 30 µM of unlabeled cold CTP (Thermo Fisher), 1.5 μM of 22 bp parS or NBS DNA duplex in the reaction buffer [100 mM Tris pH 8.0, 100 mM NaCl, and 10 mM MgCl2] for 5 min at room temperature. For the NTP competition assay, the mixture was further supplemented with 500 µM of either unlabeled cold CTP, ATP, GTP, or UTP. Four μL of samples were spotted slowly onto a dry nitrocellulose membrane and air-dried. The nitrocellulose membrane was wrapped in cling film before being exposed to a phosphor screen (GE Healthcare) for two minutes. Each DRaCALA assay was triplicated, and a representative autoradiograph was shown.

DNA preparation for EnzCheck phosphate assay and DRaCALA

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A 22 bp palindromic single-stranded DNA fragment (parS: GGATGTTTCACGTGAAACATCC or NBS: GGATATTTCCCGGGAAATATCC) [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 or NBS DNA. The sequences of parS and NBS are underlined.

Measurement of NTPase activity by EnzCheck phosphate assay

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NTP hydrolysis was monitored using an EnzCheck Phosphate Assay Kit (Thermo Fisher). Samples (100 µL) containing a reaction buffer supplemented with 1 mM of NTP and 1 µM ParB (WT or R104A) were assayed in a Biotek EON plate reader at 25°C for 15 hr with readings every minute. The reaction buffer (1 mL) typically contained: 740 μL Ultrapure water, 50 μL 20x customized 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 (1 unit). Reactions with buffer only, buffer + protein only or buffer + NTP only were also included as controls. The plates were shaken at 280 rpm continuously for 15 hr at 25°C. The inorganic phosphate standard curve was also constructed according to the manual. Each assay was triplicated. The results were analyzed using R and the NTPase rates were calculated using a linear regression fitting in R.

In vitro crosslinking using a sulfhydryl-to-sulfhydryl crosslinker bismaleimidoethane (BMOE)

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A 50 µL mixture of 10 µM ParB (C297S) or (C297S Q35C) ± 1 mM NTP ± 0.5 µM parS dsDNA (22 bp) was assembled in a reaction buffer [10 mM Tris-HCl pH 7.4, 100 mM NaCl, and 1 mM MgCl2] and incubated for 15 min (Figure 6—figure supplement 2B) or for 1, 5, 10, 15, and 30 min (Figure 6B) 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 15 min before being loaded on 12% TruPAGE Tris-Glycine Precast gels (Sigma Aldrich). Each assay was triplicated. Gels were stained with InstantBlue solution (Expedeon) and band intensity was quantified using Image Studio Lite version 5.2 (LI-COR Biosciences). The crosslinked fractions were averaged, and their standard deviations calculated in Excel.

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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

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This work addresses a long-standing question of the mechanism by which ParB spreads onto DNA flanking the parS site. The paper presents a series of well-controlled and well-thought-out biochemical experiments that are some of the first to shed light on the molecular mechanism of CTP-dependent spreading by the ParB protein. Particularly, the authors biochemically differentiate parS binding versus ParB spreading on a variety of open or closed DNA substrates. There is a strong mechanistic focus, and the detailed biochemical dissection complements and enriches results described recently in other studies on CTP-dependent spreading by ParB.

Decision letter after peer review:

Thank you for submitting your article "ParB spreading on topologically closed DNA requires cytidine triphosphate in vitro" for consideration by eLife. Your article has been reviewed by three 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 the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This well-written manuscript "ParB Spreading on topologically closed DNA requires cytidine triphosphate in vitro" by Jalal et al. addresses the long-standing question of the mechanism by which ParB spreads onto DNA flanking the parS site. The authors present a series of biochemical experiments that are some of the first to shed light on the molecular mechanism of CTP-dependent spreading by the ParB protein. Particularly, the authors biochemically differentiate parS binding versus ParB spreading on a variety of topologically open or closed DNA substrates. This work immediately follow works published by the groups of Stephan Gruber (Nov. 2019) and Martin Thanbichler (Dec. 2019) which show that ParBs from B. subtilis and M. xanthus, respectively, are CTP-binding proteins, CTPases, and CTP is required for ParB spreading. Here the authors show that C. crescentus ParB also shares these activities. There is a mechanistic focus on ParB spreading in the manuscript, thus the perspective is slightly different and the detailed biochemical dissection complements and enriches the results described in the other two studies. Unique to this paper, the authors show how ParBs, from a number of bacteria, require a topologically closed DNA substrate for spreading to be detected in vitro, providing strong evidence that this is likely a shared requirement amongst all ParBs. Based off the DNA binding and release kinetics and CTPase activity of ParB, a model for CTP-dependent ParB spreading is provided.

Essential revisions:

1) Reviewer 1: The final sentences of the Introduction are exclusive to those that have an intimate knowledge of the field. The authors need to spell-out their rationale for this paper given the findings of Easter and Gober, 2002; Osorio-Valeriano et al., 2019; and Soh et al., 2019. What is confirmatory and what is new in this paper compared to these recent publications. I recommend the authors use many of the points they make in the cover letter to make a more reader-friendly end to the Introduction.

2) All three reviewers independently identified several issues with the model figure that are either misleading or potentially incorrect based off the data presented in the paper. Since this is one of the first papers describing a detailed molecular mechanism for CTP-dependent ParB spreading, the authors should be incredibly careful, and specific, as to what the data has shown thus far, and what remains speculative. Refer to the reviewer comments below for details regarding specific concerns:

Reviewer 1: Figure 6 – the model makes assumptions that should be addressed in the legend or main-text. (i) CTP binding sites are shown to always react in concert with each other. Is there justification for this? It is possible that one fires and then hangs on (or releases) CDP+Pi, while the other monomer of ParB remains bound to CTP. These heterodimer states are not even considered. (ii) The model suggests only the Apo form of ParB can bridge DNA. Indeed, ParB has only been shown (by many) to bridge DNA without CTP. But the authors have not formally shown whether or not CTP-ParB shares this bridging activity – in addition to being able to spread.

Reviewer 2: The authors have convincingly demonstrated that Caulobacter crescentus ParB hydrolyses CTP and that parS stimulates the hydrolysis seven fold. However, CTP hydrolysis is not contemplated in the model. The authors state that CTP hydrolysis is not required for ParB to escape from the nucleation site and report the experiment performed with CTPγS, which show ParB spreading. If so, then what is the role of CTP hydrolysis in the system? This is not entirely clear to me. Secondly, if there is no double-strand break in the DNA, so no open end, how do the diffusing ParB dimers come off the DNA? Is the CTP hydrolysis invoked at this stage of the process maybe? But at this point, the ParB molecules are not bound to parS, thus the CTPase activity would be very weak. Thirdly, does ParB dimer form a completely closed ring upon CTP binding? Is it known? Overall I think that the model would benefit from some clarifications.

– Reviewers 1 and 3 independently identified critical pieces of data that, upon reviewer consultation, was deemed as necessary for supporting the author's conclusions and therefore necessary for publication:

3) Reviewer 3: The authors document parS-dependence of the constrained (169 bp) substrates, but not of the 20 bp substrate. Is ParB binding to the 20 bp DNA in BLI dependent on parS? This is an important question in part because the authors imply that binding is specific by repeatedly referring to the ParB-parS complexes, and also because they discuss this as a "nucleation event" at parS. Testing specificity is an easy experiment – measure ParB binding to the tetO substrates used in Figure 1C. Does CTP promote dissociation without parS? By the models presented, this should be confirmed.

4) Reviewer 3: Figure 1 – The data that ATP, GTP or UTP have any effect are not convincing. First, the effects are very small and second, all other ParB binding properties are unaffected by these dNTPs. The TetR control might just signal that ParB is more sensitive to salt than TetR. Also, are the counterions with the dNTPs controlled for? Most of these are sold as sodium or lithium salts (this is not mentioned in the paper). If the DNA binding is not specific for parS, one possibility for example, is that the non-specific DNA binding properties of ParB are more salt sensitive than its site-specific DNA binding properties. If the authors believe the effects of ATP, GTP and UTP are significant, they need to show that all salt is properly controlled. This is not a difficult experiment. It is possible that nucleotide effects would affect parS and non-specific DNA binding differently, and salt would need to be considered. The authors may already have done these experiments and controls, just not reported them in the paper.

5) Reviewer 1 and reviewer 3: "Spreading" – The authors are measuring DNA binding by ParB, and interpreting that this is spreading. In essence, the authors have shown parS-dependence if CTP is present and if the DNA is topologically tethered. This does not necessarily equate to spreading, which implies that multiple molecules of ParB associate and then slide away. The change in BLI signal with the larger substrates is consistent with this view, but a secondary confirmation that the ParB-parS complexes are larger would add to the confidence of the result. The authors could also be more cautious in their use of the term spreading. For example, "CTP is required for the extensive parS-dependent ParB spreading in vitro". How extensive is this on a 169 bp substrate? Reviewers suggest that at least one of the three possible experiments suggested below should be performed to support the "spreading" argument. Again, not all experiments below are required:

Reviewer 1: An in vitro ChIP assay to show that ParB is indeed spreading to DNA flanking the parS site on the topologically closed DNA substrate on the beads would be a useful addition to the interpretation of the BLI data. This is not possible with the BLI DNA because of how small the DNA fragment is. However sonication of the 2.8kb fragment down to ~250bp fragments will provide ~11 fragments in the ChIP assay with ParB antibody. This can be compared to ChIP of apo-ParB which should be sitting only on the fragments containing the parS site.

Reviewer 1: Not as direct as the experiment suggested above, but perhaps more feasible for the authors – Move the tetO site closer to and farther from the parS site to show a corresponding shift in the BLI shift. A closer tetO with TetR shown produce a lower signal. A tetO site with TetR bound right beside the anchor shown produce a shift similar to that of the topologically closed DNA without tetO.

Reviewer 3: The experiments using restriction enzyme digestion to manipulate the 169 bp substrate are a convincing way to examine the role of ends. What happens if the restriction enzyme is added after ParB binds? If this is feasible in the BLI context (i.e. move the probe into wells with everything plus the RE), it would be a nice way to establish that ParB slides off an end (right now it is inferred).

https://doi.org/10.7554/eLife.53515.sa1

Author response

Summary:

This well-written manuscript "ParB Spreading on topologically closed DNA requires cytidine triphosphate in vitro" by Jalal et al. addresses the long-standing question of the mechanism by which ParB spreads onto DNA flanking the parS site. The authors present a series of biochemical experiments that are some of the first to shed light on the molecular mechanism of CTP-dependent spreading by the ParB protein. Particularly, the authors biochemically differentiate parS binding versus ParB spreading on a variety of topologically open or closed DNA substrates. This work immediately follow works published by the groups of Stephan Gruber (Nov. 2019) and Martin Thanbichler (Dec. 2019) which show that ParBs from B. subtilis and M. xanthus, respectively, are CTP-binding proteins, CTPases, and CTP is required for ParB spreading. Here the authors show that C. crescentus ParB also shares these activities. There is a mechanistic focus on ParB spreading in the manuscript, thus the perspective is slightly different and the detailed biochemical dissection complements and enriches the results described in the other two studies. Unique to this paper, the authors show how ParBs, from a number of bacteria, require a topologically closed DNA substrate for spreading to be detected in vitro, providing strong evidence that this is likely a shared requirement amongst all ParBs. Based off the DNA binding and release kinetics and CTPase activity of ParB, a model for CTP-dependent ParB spreading is provided.

During the revision of this manuscript, my colleague pointed out that the term “topologically open/closed” might cause unintended confusion. In the original manuscript, we used the conventional definition of topology i.e. how constituent parts are linked together. However, a strict mathematical definition of DNA topology implies a specific linking number. While the conventional definition of topology has been used extensively in the literature, we agree with them that this terminology might be misinterpreted to a supercoiling-dependent DNA-binding activity by ParB-CTP in the context of our manuscript. For that reason, we replaced the term “topologically open/closed DNA” with simply “open/closed DNA” throughout the manuscript. We added sentences to the Results section to clarify this terminology:

“Immobilizing a dual biotin-labeled DNA on a streptavidin-coated BLI probe created a DNA substrate where both ends were blocked (a closed DNA) (Onn and Koshland, 2011) (Figure 2—figure supplement 1).”

“Next, we investigated whether a DNA substrate with a free end (an open DNA) can also support ParB accumulation in vitro.”

Essential revisions:

1) Reviewer: The final sentences of the Introduction are exclusive to those that have an intimate knowledge of the field. The authors need to spell-out their rationale for this paper given the findings of Easter and Gober, 2002; Osorio-Valeriano et al., 2019; and Soh et al., 2019. What is confirmatory and what is new in this paper compared to these recent publications. I recommend the authors use many of the points they make in the cover letter to make a more reader-friendly end to the Introduction.

We agree and thank the reviewer for pointing this out. We have now extended the Introduction to make it more accessible to a broader audience.

2) All three reviewers independently identified several issues with the model figure that are either misleading or potentially incorrect based off the data presented in the paper. Since this is one of the first papers describing a detailed molecular mechanism for CTP-dependent ParB spreading, the authors should be incredibly careful, and specific, as to what the data has shown thus far, and what remains speculative. Refer to the reviewer comments below for details regarding specific concerns:

We agree and have either (i) toned down our interpretation of the data or (ii) performed further experiments to clarify the conclusion. Detailed responses to specific points are given below:

Reviewer 1: Figure 6 – the model makes assumptions that should be addressed in the legend or main-text. i) CTP binding sites are shown to always react in concert with each other. Is there justification for this?

We agree that there is no justification for this assumption given our current data. We have revised the Discussion and the legend of Figure 7 to make this clear. Specifically, we added the following sentence: “Additional work is also required to investigate whether both CTP molecules on opposite subunits of a ParB dimer are concertedly hydrolyzed/dissociated for ParB to escape from the chromosome or whether a heterodimer state of ParB with a single CTP bound exists in vivo.”

It is possible that one fires and then hangs on (or releases) CDP+Pi, while the other monomer of ParB remains bound to CTP. These heterodimer states are not even considered.

We agree that our current data do not rule out the absence/existence of the heterodimer state. We have now revised the Discussion and the legend of Figure 7 to make this clear to readers (see above).

ii) The model suggests only the Apo form of ParB can bridge DNA. Indeed, ParB has only been shown (by many) to bridge DNA without CTP. But, the authors have not formally shown whether or not CTP-ParB shares this bridging activity…in addition to being able to spread.

We indeed have not investigated the bridging activity of Caulobacter ParB in this manuscript. This is an important point and warrants an in-depth investigation in future studies. We have now removed the cartoon representation of bridging from the model in Figure 7C. We revised the Discussion and added the following sentence: “There might be two different modes of action of ParB on DNA: one for bridging DNA together (that does not require CTP) and another for the lateral spreading of ParB on DNA (that requires CTP). Investigating the relative contribution of these two different modes of action to chromosome segregation in vivo is an important challenge for the future”.

Reviewer 2: The authors have convincingly demonstrated that Caulobacter crescentus ParB hydrolyses CTP and that parS stimulates the hydrolysis seven fold. However, CTP hydrolysis is not contemplated in the model. The authors state that CTP hydrolysis is not required for ParB to escape from the nucleation site and report the experiment performed with CTPγS, which show ParB spreading. If so, then what is the role of CTP hydrolysis in the system? This is not entirely clear to me. Secondly, if there is no double-strand break in the DNA, so no open end, how do the diffusing ParB dimers come off the DNA? Is the CTP hydrolysis invoked at this stage of the process maybe? But at this point, the ParB molecules are not bound to parS, thus the CTPase activity would be very weak. Thirdly, does ParB dimer form a completely closed ring upon CTP binding? Is it known? Overall I think that the model would benefit from some clarifications.

The reviewer raised important points and we felt it is prudent to perform further experiments with CTPγS to clarify the model:

Experiment 1 (Figure 6A): We fortuitously discovered that a long preincubation of purified ParB and CTPγS (but not CTP) had a negative effect on ParB accumulation on a closed DNA substrate. To investigate further, ParB was preincubated with CTP or CTPγS for 1, 5, 15, 30, or 60 min before binding to a 169-bp closed DNA substrate (Figure 6A). We observed that Caulobacter ParB could accumulate on DNA in the presence of CTPγS, but in contrast to when CTP was employed, a longer preincubation time between ParB and CTPγS gradually reduced ParB accumulation on DNA (Figure 6A). Our results suggest the possibility that CTPγS, in the absence of parS DNA, gradually converts apo-ParB in solution to a nucleation-incompetent form over time. This observation is reminiscent of a time-course experiment in which CTPγS efficiently promoted the engagement between N-terminal domains of Bacillus ParB in the absence of parS DNA (Soh et al., 2019). The engagement of N-terminal domains was shown to convert Bacillus ParB from an open to a closed protein clamp (Soh et al., 2019). If not already bound on DNA, the closed form of ParB presumably cannot self-nucleate/load onto parS due to its now inaccessible DNA-binding domain (Soh et al., 2019).

Experiment 2 (Figure 6B): We wondered if CTPγS also catalyzes the N-domain engagement in Caulobacter ParB in the absence of parS DNA. To investigate this possibility, we employed site-specific cross-linking of a purified Caulobacter ParB (Q35C C297S) variant by a sulfhydryl-to-sulfhydryl crosslinker bismaleimidoethane (BMOE) (Figure 6B, same strategy as pioneered by the Gruber lab; Soh et al., 2019). We performed controls to confirm that Q35C and C297S mutations did not impair ParB activity in vitro (Figure 6—figure supplement 2). We then performed a time-course cross-linking of ParB (Q35C C297S) + CTP or CTPγS in the absence of parS DNA (Figure 6B). CTPγS was twice as efficient as CTP in promoting the cross-linked form between N-terminal domains of Caulobacter ParB (Figure 6B). The rapid increase in the nucleation-incompetent closed form of Caulobacter ParB might explain the overall reduction in spreading over time when ParB was preincubated with CTPγS (Figure 6A).

Experiment 3 (Figure 6C): To further investigate the effect of CTPγS on ParB spreading on a longer time scale, we extended the association phase between a 169-bp parS DNA and a freshly prepared premix of ParB + CTP or CTPγS from 120 sec to 60 min (Figure 6C). In the presence of CTP, the reaction reached a steady state after 120 sec and remained stable for the duration of the association phase (Figure 6C). However, in the presence of CTPγS, ParB accumulation rapidly reached the maximal level after 200 sec, then declined slowly over the course of 60 min. We reason that DNA-bound ParB-CTPγS complexes gradually dissociated from DNA into solution but were not replenished by a new cycle of nucleation-spreading-dissociation because over time most ParB-CTPγS in solution was in a nucleation-incompetent closed form. ParB-CTPγS could possibly escape the DNA via transient ring opening or CTPγS leaving its weak binding pocket rather than by hydrolysis.

Taken together, we suggest that CTP hydrolysis is not required for parS-bound ParB to escape the nucleation site parS to spread, but possibly contributes to the stability of spreading by recycling ParB.

We hope these new data shed further light on the possible role of CTP hydrolysis and answer some of the questions that the reviewer asked. Nevertheless, we have been cautious with the interpretation in the revised manuscript and only suggest that “CTP hydrolysis possibly contributes to ParB recycling”. We also refrained from saying “ParB forms a completely closed ring upon CTP binding”, since the dynamics of ring opening and closing (when CTP is bound) are not yet known. Specifically, we added the following sentences to the Discussion:

“It is possible that ParB recycling in vivo is achieved via several routes: (i) CTP dissociation from its weak binding pocket, (ii) CTP hydrolysis, (iii) a possible enhanced CTP dissociation/hydrolysis via collisions with DNA-bound roadblocks, or (iv) a transient clamp opening even when ParB is CTP bound. Additional work is required to investigate the dynamics of ParB clamp opening/closing, and whether both CTP molecules on opposite subunits of a ParB dimer are concertedly hydrolyzed/dissociated for ParB to escape from the chromosome or a heterodimer state of ParB with a single CTP bound exists in vivo.”

– Reviewers 1 and 3 independently identified critical pieces of data that, upon reviewer consultation, was deemed as necessary for supporting the author's conclusions and therefore necessary for publication. All three reviewers think the experiments suggested below can be done in 2 months:

3) Reviewer 3: The authors document parS-dependence of the constrained (169 bp) substrates, but not of the 20 bp substrate. Is ParB binding to the 20 bp DNA in BLI dependent on parS? This is an important question in part because the authors imply that binding is specific by repeatedly referring to the ParB-parS complexes, and also because they discuss this as a "nucleation event" at parS. Testing specificity is an easy experiment – measure ParB binding to the tetO substrates used in Figure 1C. Does CTP promote dissociation without parS? By the models presented, this should be confirmed.

Unlike Bacillus ParB, which has an additional non-specific DNA-binding activity at its C-terminal domain (Taylor et al., 2015, Graham et al., 2015, Fisher et al., 2019), the C-terminal domain of Caulobacter ParB lacks this non-specific DNA-binding activity (Tran et al., 2018). Therefore, Caulobacter ParB distinguishes well between parS and non-specific DNA via its central DNA-binding domain (Tran et al., 2018). In the context of this manuscript, we have performed the suggested experiment; we measured the binding of a range of concentrations of purified Caulobacter ParB with 20-bp parS or 28-bp tetO and between TetR and parS or tetO DNA (Figure 1—figure supplement 1). Consistent with the literature, Caulobacter ParB and TetR are specific to their cognate binding sites. We have added these results to Figure 1—figure supplement 1. Finally, because Caulobacter ParB does not bind non-parS DNA to begin with, we cannot test whether CTP promotes dissociation without parS.

4) Reviewer 3: Figure 1 – The data that ATP, GTP or UTP have any effect are not convincing. First, the effects are very small and second, all other ParB binding properties are unaffected by these dNTPs. The TetR control might just signal that ParB is more sensitive to salt than TetR. Also, are the counterions with the dNTPs controlled for? Most of these are sold as sodium or lithium salts (this is not mentioned in the paper). If the DNA binding is not specific for parS, one possibility for example, is that the non-specific DNA binding properties of ParB are more salt sensitive than its site-specific DNA binding properties. If the authors believe the effects of ATP, GTP and UTP are significant, they need to show that all salt is properly controlled. This is not a difficult experiment. It is possible that nucleotide effects would affect parS and non-specific DNA binding differently, and salt would need to be considered. The authors may already have done these experiments and controls, just not reported them in the paper.

We agree with the reviewer that the effect of ATP, GTP, and UTP was small and did not affect other binding properties of Caulobacter ParB. We further performed BMOE cross-linking experiment to investigate the effect of NTPs on the N-domain engagement of Caulobacter ParB, and indeed ATP, GTP, and UTP had a negligible effect (Figure 6—figure supplement 2B).

We have also performed additional experiment to investigate the sensitivity of ParB to salts/counterions (Na+)/other contaminants that came with the purchased NTP solutions. Specifically, we measured binding of ParB to a 20-bp parS DNA with/without NTPs in buffer lacking Mg2+. All NTPs (in buffer lacking Mg2+) decreased ParB-parS interaction very slightly (but reproducibly), suggesting that ParB might be slightly sensitive to salt/counterions/contaminants in the purchased NTP solutions. Again, given that the effect of ATP, GTP, or UTP (in buffer with or without Mg2+) in all different types of experiments (bio-layer interferometry/BMOE cross-linking/DRaCALA/NTP hydrolysis) was small, we toned down our interpretation and focused solely on the specific effect of CTP on Caulobacter ParB properties.

5) Reviewer 1 and reviewer 3: "Spreading" – The authors are measuring DNA binding by ParB and interpreting that this is spreading. In essence, the authors have shown parS-dependence if CTP is present and if the DNA is topologically tethered. This does not necessarily equate to spreading, which implies that multiple molecules of ParB associate and then slide away. The change in BLI signal with the larger substrates is consistent with this view, but a secondary confirmation that the ParB-parS complexes are larger would add to the confidence of the result. The authors could also be more cautious in their use of the term spreading. For example, "CTP is required for the extensive parS-dependent ParB spreading in vitro". How extensive is this on a 169 bp substrate?

These are excellent points and we agree with the reviewers. In the revised manuscript, we are more cautious in using the term “spreading”. Instead, we use the phrase “accumulation of ParB” whenever appropriate and only use “spreading” when data are available to back up this claim.

Reviewers suggest that at least one of the three possible experiments suggested below should be performed to support the "spreading" argument. Again, not all experiments below are required:

Clear instruction from the editor and reviewers here is much appreciated. Below are the results of the suggested experiments:

Reviewer 3: The experiments using restriction enzyme digestion to manipulate the 169 bp substrate are a convincing way to examine the role of ends. What happens if the restriction enzyme is added after ParB binds? If this is feasible in the BLI context (i.e. move the probe into wells with everything plus the RE), it would be a nice way to establish that ParB slides off an end (right now it is inferred).

We have now performed this experiment with BamHI (or heat-killed BamHI as a negative control) added after ParB-CTP was bound on a closed DNA substrate (Figure 3—figure supplement 2). After adding BamHI, ParB binding to DNA reduced gradually over the course of 30 minutes, while it was unaffected when an inactivated BamHI was used instead (Figure 3—figure supplement 2). This result backs up our interpretation that ParB slides off an open DNA end.

N.B. Experiments were performed at 22oC instead of at 37oC to keep ParB stable. Also, the binding of ParB-CTP to DNA was performed in NEB buffer 3.1 [100 mM NaCl, 50 mM Tris-HCl, 10 mM MgCl2, 100 µg/ml BSA, pH 7.9] instead of the low-salt binding buffer used throughout the manuscript [100 mM NaCl, 10 mM Tris-HCl, 1 mM MgCl2, 0.005% Tween20, pH 8.0]. NEB buffer 3.1 was used to ensure BamHI digested DNA as optimally as possible while keeping Caulobacter ParB stable. Due to the change in buffers, the BLI response of ParB-CTP with DNA was higher in buffer NEB 3.1 than in the usual binding buffer used throughout the manuscript. Materials and methods section has been updated to reflect this.

Reviewer: Not as direct as the experiment suggested above, but perhaps more feasible for the authors – Move the tetO site closer to and farther from the parS site to show a corresponding shift in the BLI shift. A closer tetO with TetR shown produce a lower signal. A tetO site with TetR bound right beside the anchor shown produce a shift similar to that of the topologically closed DNA without tetO.

We performed a variant of the suggested experiment (Figure 4—figure supplement 1): a premix of ParB, CTP, and TetR was allowed to bind to a closedDNA substrate and, after the system reached steady state, anhydrotetracycline was added to remove bound TetR from the DNA. After the roadblock (TetR) was removed by anhydrotetracycline, we observed that ParB binding to the closed DNA increased (dotted magenta line, Figure 4—figure supplement 1), consistent with the “spreading” phenomenon. We have performed a control showing that ParB-CTP binding to DNA is insensitive to added anhydrotetracycline in the absence of TetR (solid and dotted orange lines, Figure 4—figure supplement 1).

Reviewer 1: An in vitro ChIP assay to show that ParB is indeed spreading to DNA flanking the parS site on the topologically closed DNA substrate on the beads would be a useful addition to the interpretation of the BLI data. This is not possible with the BLI DNA because of how small the DNA fragment is. However sonication of the 2.8kb fragment down to ~250bp fragments will provide ~11 fragments in the ChIP assay with ParB antibody. This can be compared to ChIP of apo-ParB which should be sitting only on the fragments containing the parS site.

We attempted this experiment for the original manuscript but were not successful, most likely because fixing Caulobacter ParB to DNA (+/- CTP) in vitro with 1% formaldehyde inactivated most of ParB protein. We have not optimized the concentration of formaldehyde for fixation any further due to time constraints. In previous work, we were successful in performing a non-crosslinking version of in vitro ChIP-seq (IDAP-seq) using sonicated genomic DNA and purified ParB (Tran et al., 2018). However, this methodology is inappropriate in the presence of CTP since we now have ample evidence that ParB-CTP slides off the ends of sonicated DNA.

N.B.While assembling source data to upload to eLife, I found that some parts of Figure 2—figure supplement 3 were mislabeled in the original manuscript. Specifically, Morella thermoacetica His6-MBP-ParB should be Dechloromonas aromatica His6-MBP-ParB, and 0.01 mM CTP was used for Zymomonas mobilis and Xanthomonas campestris ParB instead of 1mM CTP used for all other ParB samples. To double check, we purified these eight proteins again using batch purification and redid the experiments for Figure 2—figure supplement 3. I apologize for the oversight. These two mislabellings have been corrected and the figure legend updated in the revised manuscript. The conclusion that “we performed BLI experiments for eight additional chromosomal ParB proteins from a diverse set of bacterial species and consistently observed the specific effect of CTP on enhancing ParB association with a closed DNA in vitro (Figure 2—figure supplement 3). It is most likely that ParB-CTP interaction with DNA is conserved among ParB orthologs”stands.

A higher concentration of CTP was inhibitory to Zymomonas and Xanthomonas ParB, presumably because most of these apo-ParB was converted to the closed clamp form before exposure to tested DNA.

https://doi.org/10.7554/eLife.53515.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
    Competing interests
    No competing interests declared
    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, Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7186-3976
  3. Tung BK Le

    Department of Molecular Microbiology, John Innes Centre, Norwich, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    For correspondence
    tung.le@jic.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4764-8851

Funding

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

  • Tung BK Le

Royal Society (UF140053)

  • Tung BK Le

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

  • Ngat T Tran
  • Tung BK Le

Royal Society (RG150448)

  • Adam SB Jalal
  • Tung BK Le

Biotechnology and Biological Sciences Research Council (BBS/E/J/000PR9778)

  • Tung BK Le

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

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. Received: November 11, 2019
  2. Accepted: February 19, 2020
  3. Accepted Manuscript published: February 20, 2020 (version 1)
  4. Version of Record published: March 3, 2020 (version 2)

Copyright

© 2020, 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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