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Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Bacterial chromosome segregation usually involves cytoskeletal ParA proteins, ATPases which can form dynamic filaments. In aerial hyphae of the mycelial bacterium Streptomyces coelicolor, ParA filaments extend over tens of microns and are responsible for segregation of dozens of chromosomes. We have identified a novel interaction partner of S. coelicolor ParA, ParJ. ParJ negatively regulates ParA polymerization in vitro and is important for efficient chromosome segregation in sporulating aerial hyphae. ParJ–EGFP formed foci along aerial hyphae even in the absence of ParA. ParJ, which is encoded by sco1662, turned out to be one of the five actinobacterial signature proteins, and another of the five is a ParJ paralogue. We hypothesize that polar growth, which is characteristic not only of streptomycetes, but even of simple Actinobacteria, may be interlinked with ParA polymer assembly and its specific regulation by ParJ.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

ParA homologues (such as Soj in B. subtilis) are part of the machinery responsible for the rapid movement of bacterial plasmids or chromosomal origin regions (oriC) towards cell poles soon after the start of replication (Gerdes et al., 2000; 2004; Hayes and Barillà, 2006; Toro and Shapiro, 2010). Soj also plays a role in the regulation of chromosome replication and control of sporulation (Marston and Errington, 1999; Quisel et al., 1999; Murray and Errington, 2008). ParA homologues generally act in collaboration with ParB, a protein encoded by the second gene of the same operon (Gerdes et al., 2000). ParB is also required for efficient plasmid and chromosome segregation, forming high-order nucleoprotein complexes at partitioning sites (parS) near oriC (Lin and Grossman, 1998; Jakimowicz et al., 2002; Leonard et al., 2004).

ParA homologues are Walker A cytoskeletal ATPases. Thermus thermophilus ParA forms a sandwich dimer (Leonard et al., 2005a) and can oligomerize into filaments in the presence of DNA. The weak ATPase activity of ParA proteins is elevated in the presence of ParB homologues (Easter and Gober, 2002; Barillàet al., 2005; Leonard et al., 2005a; Bouet et al., 2007). Filaments formed by the ParA homologue of plasmid pTP228 (ParF) are remodelled in the presence of its ParB homologue due to enhancement of ParF ATPase activity (Barillàet al., 2005; 2007). Polymerization and depolymerization of ParA underpin its dynamic localization pattern in the cell (Ebersbach and Gerdes, 2004; Lim et al., 2005; Fogel and Waldor, 2006; Ringgaard et al., 2009). Several ParA homologues (SopA from F plasmid and ParA from plasmid pB171, chromosomal ParA from B. subtilis and V. cholerae) have been reported to oscillate within the cell as a helical spiral. This dynamic localization can be disrupted by mutations in the nucleotide binding site or in parB (Marston and Errington, 1999; Quisel et al., 1999; Autret et al., 2001; Fogel and Waldor, 2006). ParA has been postulated to supply energy for the movement of ParB segregation complexes (Leonard et al., 2005b). Disassembly of the ParA polymer of plasmid pB171, triggered by interaction with the segregation complex, moves the plasmids along the cell (Ringgaard et al., 2009). Similarly, in V. cholerae a dynamic band of ParAI shifts the segregating ParBI/parS complex to the cell pole to anchor the chromosome there (Fogel and Waldor, 2006).

The mycelial growth mode of Streptomyces coelicolor and its multiple synchronous septation during sporulation provide new perspectives on the mechanisms of chromosome segregation. During vegetative growth, hyphae elongate by tip extension and branching, while occasional septa separate adjacent multigenomic compartments (Flärdh, 2003). Sporulation, triggered by nutrient depletion, starts with rapid growth of aerial hyphae, which involves intensive chromosome replication (Ruban-Ośmiałowska et al., 2006). In aerial hyphae 50 or more non-segregated chromosomes may be present in one long tip compartment. Conversion of multigenomic aerial hyphae into chains of unigenomic spores requires synchronous segregation of multiple chromosomes and synchronous regular placement of Z-rings along the compartment (McCormick et al., 1994; Flärdh and Buttner, 2009). In contrast to the movement of chromosomes towards poles in rod-shaped bacteria, dozens of Streptomyces chromosomes are condensed and uniformly aligned along the hyphal tip compartment, ensuring that each prespore receives a single copy.

ParAB proteins are important for proper distribution of chromosomes in aerial hyphae before septation. The parAB operon is developmentally regulated, with one of its two promoters strongly upregulated shortly before sporulation septation (Kim et al., 2000; Jakimowicz et al., 2006). Deletion of parA or parB results in frequent anucleate spores (about 24% or 15% respectively), but does not visibly affect colony growth or sporulation. Elimination of ParA also strongly affects sporulation septation. Before DNA segregation and septation, ParB binds to numerous parS sites near oriC to form large complexes, which are arrayed regularly along the aerial hyphal tip compartment, disassembling soon after septation is completed (Jakimowicz et al., 2002; 2005). ParA has a complex localization pattern, dependent on the growth stage. In vegetative hyphae and young aerial hyphae it is found at the tips, but during further aerial growth it forms extended helical filaments along apical compartments, as ParB complexes form (Jakimowicz et al., 2007). ParA mediates efficient assembly of ParB complexes in vivo and in vitro, and ParA ATPase activity is crucial for its dimerization and interactions with ParB, though not for ParA localization in vivo. We have postulated that ParA provides scaffolding for proper distribution of ParB complexes and consequently controls synchronized segregation of several dozens of chromosomes, possibly mediating a segregation and septation checkpoint.

ParB was the only known interaction partner of ParA until recent evidence that B. subtilis Soj interacts with replication initiator DnaA, controlling its activity (Murray and Errington, 2008). Here, we show that a novel interaction partner of Streptomyces ParA, ParJ, influences ParA filament stability. Remarkably, ParJ is one of a very small number of signature proteins confined to, but ubiquitous among, Actinobacteria, whose members include many organisms of medical, environmental and industrial importance (Gao et al., 2009).

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

ParA interacts with a conserved, actinobacteria-specific protein

Interesting properties of S. coelicolor ParA, which include formation of elongated filaments in aerial hyphae, encouraged us to search for its interaction partners, using a bacterial two-hybrid system (BTH) to screen an expression library of S. coelicolor DNA (a kind gift from Professor Justin Nodwell, McMaster University, Canada, unpublished). One of the positive clones contained sco1662, encoding a conserved hypothetical protein SCO1662 found exclusively within Actinobacteria. In S. coelicolor, sco1662 is located downstream of the glycerol utilization operon and was thought to be the last gene of the operon (gylX), with an unknown function (Smith and Chater, 1988). However, comparison of gene arrangement in different Streptomyces species revealed that the position of the sco1662 homologues downstream of the gyl operon is not conserved (Fig. S1), suggesting that SCO1662 is not associated with glycerol catabolism. Indeed, the sco1662 orthologues are highly unusual among conserved Streptomyces genes in their non-syntenous locations in the genomes of different species. The protein was named ParJ for the reasons that are given below.

The ParA–ParJ interaction was verified in vitro using purified ParA (Jakimowicz et al., 2007) (or 6His-ParA) and recombinant ParJ purified in two expression systems: 6His–ParJ and ParJ (36 kDa) cleaved from a glutathione S-transferase (GST) fusion (Fig. S2). In one approach, a strong, concentration-dependent interaction of 6His-ParA with ParJ (cleaved from GST) immobilized on a microtitre plate was detected with an anti-ParA antibody (Fig. 1). In a second approach, using surface plasmon resonance (SPR), ParA was immobilized on a CM5 chip, and tested for interaction with increasing concentrations of 6His–ParJ (empty flow cell served as negative control for unspecific binding to the chip surface). The SPR studies showed binding with a KD of about 30 nM (calculated from steady-state affinity and from kinetic parameters) (Fig. 2A). Additionally, the kinetic analysis indicated a very slow dissociation rate (kd = 8.3 × 10−4 1/s). Thus the interaction of ParJ and ParA was confirmed.

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Figure 1. Interaction of ParA with SCO1662 (ParJ). Binding of 6His-ParA (10–5000 ng) to ParJ (cleaved from GST), or BSA, bound to a microtitre ELISA plate was detected with anti-ParA antibody conjugated with alkaline phosphatase, whose activity was detected by measuring absorbance of reaction product at 405 nm. Inset: Bacterial two-hybrid analysis of interaction of T25-ParA with T18-ParJ. Intense colony colour indicates interaction of the analysed proteins.

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Figure 2. Analysis of ParJ–ParA binding by SPR. Five hundred RU (response units) of ParA or ParAK44E immobilized on CM5 was tested for interaction with 6His–ParJ. A. ParA interaction with increasing concentrations of 6His–ParJ (as indicated). B. Comparison of 6His–ParJ binding to ParA and ParAK44E. (Inset) Bacterial two-hybrid analysis of interaction of T25-ParAK44E with T18-ParJ. Intense colour of colonies indicates interaction of analysed proteins.

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Deletion of parJ affects the formation of aerial hyphae, sporulation septation and chromosome partitioning into spores

Since ParA extended filament is important for efficient chromosome segregation and regular septation in sporulating aerial hyphae, we expected that the newly identified ParA partner might also have a specific function at this stage. The question regarding its role in development and its connection with ParA could be answered by the construction of DJ548, a parJ deletion mutant. The mutant showed normal colony morphology on MS solid medium, but its aerial growth and sporulation started somewhat earlier than in the wild-type, particularly on the rich medium R2YE, which generally slows development. Aerial hyphae, as observed by the white colony phenotype, developed after 44–48 h of growth, compared with 70–90 h for the wild-type strain (Fig. 3A). Introduction of a wild-type copy of parJ with its putative native promoter (297 bp of upstream DNA) into the chromosome of parJ deletion mutant, DJ548 (yielding strain DJ556) restored the normal rate of development (Fig. 3B). Developmental acceleration was independent of ParA, since the same phenotype was shown by a parJparA double deletion strain, DJ555 (Fig. 3A).

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Figure 3. Developmental acceleration of parJ deletion strains. A. Development of parJ deletion strains and their parents on R2YE agar. B. Complementation of parJ deletion on R2YE agar.

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Further studies of mutant strains DJ548 (parJ deletion) and DJ555 (parJparA double deletion) used fluorescent microscopy to analyse chromosome segregation and septation. DNA and cell wall staining (with DAPI and wheat germ agglutinin, WGA-Texas Red conjugate, respectively) revealed that 8% of prespore compartments were anucleate in the parJ deletion mutant (Fig. 4, Table 1). In addition, prespore compartments were more uneven in size, compared with wild-type, with about 18% being minicompartments (shorter than 0.8 µm, half of the normal length) (Table 1). Chains of ‘early’ spores of strain DJ548 (parJ deletion) were also more susceptible to cell wall staining with WGA than the wild-type, indicating that their cell wall was less fully mature (Fig. S3). However, the mature spores were as resistant to SDS, temperature and lysosyme treatment as the wild-type. The segregation and septation defects were both complemented in DJ556 (Table 1). Altogether, these results showed that ParJ is involved in sporulation-associated chromosome segregation and cell division.

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Figure 4. Segregation and septation defect of a parJ deletion strain. Images of hyphae showing cell walls staining (red) and DNA stained with DAPI (blue). A. M145 (wild-type). B. DJ548 (parJ deletion strain). C. DJ556 (complementation strain). Minicompartments (shorter than 0.8 µm) are indicated by grey arrowheads, approximately normal-sized anucleate compartments by red arrowheads. Scale bars 5 µm.

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Table 1.  Statistical analysis of segregation/septation defects in parJ mutant strains. Thumbnail image of

Irregular septation and a large fraction of minicompartments, and a somewhat more severe segregation defect, were previously observed in a parA deletion strain (Jakimowicz et al., 2007, Table 1). The parJparA double mutant, DJ555, had a phenotype intermediate between the two single mutants (anucleate prespore compartments 13%, minicompartments about 13%, see Table 1). Thus, the phenotypes were not additive – instead, parJ deletion partially suppressed the segregation and septation defects of the parA deletion. The effects of parA overexpression in the parJparA deletion background were also investigated by integrating pIJ6902parA, containing parA controlled by the inducible tipA promoter, into DJ555 (strain DJ557). Remarkably, parA overexpression partially complemented the irregular septation and segregation phenotype of parJ deletion, giving 1–2% of anucleate prespore compartments and 8–10% minicompartments, similar to the phenotype of parA overexpression in a parA single deletion background (DJ553). In summary, these genetic interactions reinforce the notion that the two proteins interact in vivo but also suggest that some parJ function(s) are independent of its partner protein ParA.

ParJ foci accompany ParA filaments in aerial hyphae

We investigated the cytological distribution of ParJ, using a strain DJ558 in which parJ was replaced by a parJegfp gene fusion in its chromosomal locus. ParJ–EGFP was almost fully functional as demonstrated by DNA and cell wall staining, which showed only a very slight segregation/septation defect in DJ558 (Table 1). Fluorescence microscopy revealed very weak EGFP fluorescence in DJ558 vegetative mycelium, but in some aerial hyphae ParJ–EGFP formed numerous small foci (Fig. 5A). We analysed 180 unseptated, possibly nascent aerial hyphae and 122 septated hyphae. About 40% of the unseptated hyphae showed no fluorescence, 34% contained only sparsely spaced and rather faint foci, and 25% contained intense and densely spaced foci. Intense foci were present in most (67%) of the septated hyphae foci. Thus, ParJ foci form in aerial hyphae at or close to the end of growth, before sporulation septation, and persist somewhat beyond the establishment of the septa.

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Figure 5. Localization of ParJ–EGFP foci in the aerial hyphae. A. Presence of ParJ–EGFP foci during septation. Images of aerial hyphae showing: left panel, ParJ–EGFP foci fluorescence (green), middle panel, cell wall staining with WGA-TexasRed (red), and right panel, merged fluorescence in DJ558 (M145parJ–egfp-apra); yellow arrow, early non-septated hyphae with faint ParJ foci; green arrow, non-septated hyphae with bright ParJ foci; red arrow, septated hyphae. Scale bars, 5 µm. B. Statistical analysis of intervals between ParJ–EGFP foci of DJ558 in septated and unseptated aerial hyphae calculated for about 200 foci.

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Foci in prespore compartments were slightly more densely packed than in non-septated hyphae, with the respective average inter-foci distances of 0.95 µm and 1.05 µm both being less than the length of a prespore compartment (about 1.3–1.4 µm) (Fig. 5B). Overall, 67% of prespore compartments had a single focus, about 26% contained two, and almost 5% three foci. A few compartments had more than three foci. Thus, the distribution of the foci is not closely related to the positions of septa.

To correlate the temporal patterns of subcellular localization of ParJ and ParA, we performed immunolocalization of ParA in strain DJ558 expressing parJ–egfp (Fig. 6A). In young aerial hyphae ParA localizes close to the tips, and later in aerial hyphal growth its signal extends towards the hyphal base, resembling a helical filament. Both faint and intense ParJ foci could be accompanied by partially or fully extended ParA filaments. This implies that ParJ foci form during the extension of ParA filaments, though the technical limitations of immunofluorescence impeded analysis of the dynamics of the two processes together in real time. Since ParA filaments were absent from septated hyphae, ParJ foci are longer-lived than ParA filaments.

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Figure 6. ParJ foci accompany ParA filaments in aerial hyphae. A. Coincidence of ParA and ParJ in aerial hyphae. Images of hyphae showing: left panel, ParA immunofluorescence (red); middle panel, ParJ–EGFP (green); and right panel, merged fluorescence. A, hyphae with ParA filament; J, hyphae with ParJ foci. Scale bars, 5 µm. B. Statistical analysis of intervals between ParJ–EGFP foci in both septated and unseptated (combined) aerial hyphae of DJ558 (‘wild-type’) and DJ559 (parA deletion).

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The formation of ParJ–EGFP foci was not much affected by the elimination of ParA. The intensity and average distances between foci formed in J3306 (parA deletion) were the same as in the wild-type, except that inter-foci distances varied slightly more (Fig. 6B). Thus, if there is any influence of ParA on ParJ distribution, it is small. In a reciprocal experiment, we did not observe any differences from the wild-type in ParA localization in parJ deletion strain DJ548 (data not shown).

Taken together, our results showed that ParJ forms foci in aerial hyphae at the time of extension of ParA filaments but the formation of foci does not depend on ParA. Transcription analysis (Fig. S4) showed that the developmental assembly of ParJ foci is underpinned by developmental induction of two parJ promoters, with parJ transcription apparently being negatively regulated by ParA at later time points.

ParJ induces ParA depolymerization in vitro

ParA polymerization and its ATPase activity are crucial for its function in chromosome segregation. Previously, it was found that ATP binding is required for ParA polymerization, while during disassembly of the filament ATP is presumably hydrolysed (Leonard et al., 2005b). To determine whether ParJ affects ParA polymerization in vitro, we monitored filament formation by ParA using pelleting assays, in which polymerized and unpolymerized proteins were separated into pellet and supernatant fractions, respectively, and further analysed by SDS/PAGE. Using this assay we found that in the presence of ATP Streptomyces ParA, like the other ParA homologues (but not requiring presence of DNA), formed higher-order complexes that would be consistent with filament formation: the protein was present only in the pellet fraction (Fig. 7A; B. Ditkowski, unpublished). However, in the presence of ATP and an equimolar amount (7 µM) of 6His–ParJ, ParA was retained in the supernatant fraction, suggesting that ParJ either inhibited ParA polymerization or induced depolymerization (Fig. 7A).

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Figure 7. Influence of ParJ on ParA polymerization. A. Pelleting assay. ParA (6.66 µM) polymerization in the presence of ATP and 6His–ParJ protein (7 µM), as indicated. P, pellet fraction/polymerized protein; S, soluble fraction. B. ParA polymerization in the presence and absence of 6His–ParJ detected by dynamic light scattering. The top panel shows that augmentation in polymer average size (nm) is not seen in the presence of 6His–ParJ. Bottom panel, changes in light-scattering intensity, expressed as kct s−1. C. ParA depolymerization induced by addition of 6His–ParJ, detected by dynamic light scattering. The time of addition of increasing concentrations of 6His–ParJ to polymerized ParA is marked with an arrow. Top panel, change in polymer average size (nm). Bottom panel, changes in light-scattering intensity, expressed as kct s−1.

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The influence of ParJ on ParA filament assembly/disassembly in real time was studied using dynamic light scattering (DLS), which assesses both abundance and size of particles. For a ParA solution (3 µM) in the presence of ATP, the recorded average intensity of light scattering increased up to about 2000 kilocounts/second (kct s−1) and the particle size up to 500 nm. In the presence of 6His–ParJ (1.5 µM) and ATP both parameters were reduced, to about 500 kct s−1 and to 100 nm respectively. Thus ATP-dependent polymer formation by ParA was inhibited by ParJ (Fig. 7B). Moreover, addition of 6His–ParJ (0.75–7 µM) to earlier polymerized ParA filaments led to an immediate, ParJ concentration-dependent, decrease of light scattering and particle size, indicating rapid polymer disassembly (Fig. 7C)

To find out if the observed ParA depolymerization effect could result from stabilization of the monomer form, we tested the interaction of ParJ with ParAK44E, which contains a mutation in the Walker A motif and can neither bind ATP nor dimerize (Jakimowicz et al., 2007). BTH and SPR studies showed that ParJ interacts with ParAK44E, albeit less strongly than with the wild-type (Fig. 2B). To test if ParJ affects ATPase activity of ParA we employed an assay using radioactive ATP. Surprisingly, we found that ParJ itself (cleaved from GST) exhibited ATPase activity (Fig. 8A, inset). Additionally, the rate of ATP hydrolysis for ParA, 6His–ParJ and mixture of the proteins was monitored in the quantitative assay by measurement of released phosphate (PiPer Phosphate Assay Kit, Molecular Probes). These experiments showed that 6His–ParJ also had ATPase activity (thus, activity was independent of the purification method) and was more efficient in hydrolysing ATP than ParA (Fig. 8A). When both proteins were mixed, the ATPase activity was similar to the summed activities of the two separate proteins (Fig. 8A), thus ParJ presumably does not enhance ParA activity. However, further studies will be required to elucidate in detail any reciprocal effects on ATPase activity. Since ParJ does not contain any recognizable ATP binding motif, and in order to exclude potential contamination with ATPase, we confirmed binding of ATP to ParJ in an independent UV-cross-linking experiment (Fig. 8B). The observed binding was Mg2+ dependent and was observed for both ParJ cleaved from GST and 6HisParJ.

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Figure 8. ATPase activity of ParJ. A. ATPase activity of 6His–ParJ, ParA and mixed 6His–ParJ and ParA proteins, all at 1 µM concentration. The control reactions (ATP hydrolysis, and internal protein control) were subtracted. Inset, hydrolysis of [γ-32P]-ATP by ParJ. B. UV cross-linking of ParJ binding [α-32P]-ATP. Following cross-linking, samples were analysed by SDS-polyacrylamide gel electrophoresis and the gel was exposed to a phosphorscreen. Top panel shows Coomassie blue-stained SDS-polyacrylamide gel of UV cross-linked samples, bottom panel – autoradiography image from the UV cross-linking experiment 1, ParJ (cleaved from GST) in the presence of MgCl2; 2, ParJ (cleaved from GST) in the absence of MgCl2; 3, 6His–ParJ in the presence of MgCl2; 4, negative control – ParB in the presence of MgCl2.

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Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

An actinobacterial signature protein is a partner of the cytoskeletal protein ParA

We have identified a completely novel ParA interaction partner, ParJ, that is necessary for efficient chromosome partitioning during sporulation septation. A recent extensive phylogenomic analysis of 19 Actinobacteria revealed that sco1662 is one of only five signature genes for the phylum, while a second of the five is also a parJ paralogue (Gao et al., 2009). In S. coelicolor this protein, SCO1997 (which shows 25% identity and 43% similarity to ParJ), has been crystallized (Gao et al., 2009), revealing that it forms a trimer with a large internal cavity surrounded by three pronged, claw-like domains. The electrostatic surface potential of each claw is negative, generating an overall acidic surface in this area that might interact with other, positively charged, protein(s). The sequence similarity of ParJ to SCO1997 and modelling studies suggest that it may have the same trimer structure. Conserved lysines and arginines of the C-terminal region of ParA could provide an interaction interface with ParJ. Additionally, divalent cation binding was predicted for SCO1997, which is consistent with our finding of ParJ ATPase activity. Since the SCO1997 structure does not contain any known nucleotide binding site, further studies will be required to characterize the ATP binding motif. The structures of SCO1997 and ParJ do not match any known proteins, so elucidation of their function is likely to provide novel insights into physiological characteristics that are unique to Actinobacteria, and which might be investigated in the context of disease therapy (the loss of ParA from Corynebacterium and from Mycobacterium severely reduces growth rate: Donovan et al., 2010; Nisa et al., 2010; K. Ginda et al., manuscript in preparation).

ParJ is responsible for ParA disassembly

Previously, several lines of evidence, mainly from plasmid systems, have shown that the dynamics of ParA polymerization is affected by ParB proteins (Møller-Jensen et al., 2003; Garner et al., 2004; Barillàet al., 2005; Bouet et al., 2007), but our finding that ParJ brings about ParA filament disassembly seems to be the first report of a different regulator of ParA polymerization. All studied ParB homologues appear to enhance nucleotide hydrolysis by cognate ParAs (Easter and Gober, 2002; Leonard et al., 2005a; Bouet et al., 2007; Jakimowicz et al., 2007). Leonard et al. (2005a) suggested two possible mechanisms of ATPase activation of ParA by ParB. In the first model, favoured by Leonard et al., activation of the ATPase activity of ParA polymer subunits leads to destabilization of the polymer; alternatively, according to the second model, enhanced ATPase activity might result from promotion of nucleotide exchange in the ParA-ADP monomer, shifting the equilibrium to the ATP-bound, polymerization-proficient dimer. There are also several lines of evidence showing that dynamics of polymer formed by ParA homologues is affected by ParB proteins. For example, in the case of plasmid pTP288 ParF, addition of ParG (ParB homologue) resulted in extensive remodelling of the polymer in vitro (Barillàet al., 2005). Enhanced ParA filamentation was also reported for plasmid proteins SopA/SopB (Bouet et al., 2007) and suggested for the ParM–ParR system, where interaction with ParR stabilizes the ParM filament, facilitating its continued polymerization (Møller-Jensen et al., 2003; Garner et al., 2004).

Our data also showed ParJ interacted with non-dimerizing ParAK44E, which is not able to bind ATP, albeit less strongly than with the wild-type protein. Thus, ParJ-mediated inhibition of ParA polymer assembly could result from ParA monomer stabilization. However, we could not exclude ParJ-dependent induction of ATPase activity of ParA as an explanation for ParA polymer destabilization.

ParJ is required for efficient chromosome segregation and septation

Elongated ParA filaments formed during the formation of ladders of Z-rings in aerial hyphae, depolymerizing at the stage of synchronous septation (Jakimowicz et al., 2007). The timing of formation of ParJ foci, particularly their presence in septated hyphae, is consistent with their role during final ParA filament disassembly. Our failure to detect any change in ParA filament dynamics in the parJ deletion mutant may be due to limitations of the immunofluorescence technique. In a parB deletion background (data not shown) ParJ foci still formed but, surprisingly, seemed less distinct than in the wild-type, presumably either because of some alteration of ParA filaments in the absence of ParB, or because of disturbed regulation of ParJ levels. Analysis of a set of mutant strains provided evidence that the functions of both proteins during septation and segregation are interlinked. The segregation defect of the double parJparA deletion strain was less severe than that resulting from ParA elimination. This partial phenotype could be a reflection of an increase in the amount of ParJ available in the parA deletion mutant to interact with, and sequester, some other element of the segregation/cell division machinery. This would be overcome by deleting parJ. Moreover, the level of parJ transcript – and therefore probably of ParJ – is elevated in a parA deletion mutant (Fig. S4). We tentatively suggest that ParJ interacts with some other subsequent element of segregation/cell division machinery. Our preliminary BTH tests for interaction of ParJ with some known division proteins (SsgB, FtsZ, MreB) did not give any positive result, so further studies are required to find protein(s) or subcellular structures that interact with ParJ.

The hypothesis of a more complex ParJ function than solely regulation of ParA filament dynamics is supported by the accelerated development of parJ deletion colonies. Since the same effect was observed in the parA deletion background it suggests some additional regulatory function for ParJ. At present we have no strong candidates for ParJ targets in the potential regulatory circuit, although one possibility would be the parAB operon itself.

Concluding remarks

In summary, we have described the first regulator of ParA polymerization other than ParB. Taking into consideration the conservation of both ParJ and polar growth among Actinobacteria and their absence from other bacterial phyla, we suggest a polar growth-related regulatory function for ParJ. ParJ may contribute to special ParA filament dynamics required for polar growth. Immediate questions are whether the conserved ParJ paralogue SCO1997 is also involved in ParA dynamics, and whether there is some interplay between ParJ and SCO1997. Presumably, any such interplay would be found throughout the Actinobacteria. More broadly, it is likely that in other bacterial phyla similarly complex molecular mechanisms govern the formation of supramolecular structures by ParA.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

DNA manipulations, bacterial strains, media and growth conditions, and sample preparation for microscopy, were as described before (Kieser et al., 2000; Jakimowicz et al., 2007). Application of the BTH system (Karimova et al., 2000) and library screening are described in supplementary material (Supporting information text). PCR targeting was used for the construction of mutant strains (Table 2). The PCR targeting procedure was applied as described before (Gust et al., 2003; Jakimowicz et al., 2007); details of the construction are given in supplementary material (Supporting information text). Protein purification followed published procedures; details are provided in supplementary material (Supporting information text).

Table 2.  Strains used in this study.
StrainRelevant genotypeSource
E. coli
DH5αsupE44ΔlacU169(φ80lacZΔM15)hsdR17 recA1 endA1 gyrA96 thi-1 relA1Lab stock
BW25113/pIJ790K12 derivative: ΔaraBAD, ΔrhaBADλ-Red(gam,bet,exo), cat, araC, rep101tsGust et al. (2003)
ET12567/pUZ8002dam-13::Tn9, dcm cat tet hsd zjj-201::Tn10/tra neo RP4Kieser et al. (2000)
BTH101F-, cya-99, araD139, galE15, galK16, rpsL1 (Strr), hsdR2, mcrA1, mcrB1Karimova et al. (2000)
S. coelicolor
M145SCP1-, SCP2-Kieser et al. (2000)
J3306M145ΔparAJakimowicz et al. (2007)
DJ548M145Δsco1662::apraThis study
DJ553J3306pIJ6902(hygr)parAThis study
DJ555J3306ΔparJ::apraThis study
DJ556M145ΔparJ::apra, pIJ82parJThis study
DJ557J3306ΔparJ::apra pIJ6902(hygr)parAThis study
DJ558M145parJ–egfp-apraThis study
DJ559J3306parJ–egfp-apraThis study

Surface plasmon resonance analysis

For SPR analysis, untagged ParA protein (500 response units) was immobilized on the CM5 Sensor Chip in presence of ATP according to the manufacturer instructions. As a control, a flow channel without immobilized protein was used. SPR analysis was performed on a BiaCore 3000, by applying increasing concentrations of 6His–ParJ (as indicated) solutions in binding buffer (10 mM HEPES/KOH, pH 7.4, 150 mM NaCl, 50 µg ml−1 BSA, 5 mM MgCl2, 0.05% Tween 20). Binding was measured for 3 min at flow rate 20 µl min−1 at room temperature in SPR running buffer (10 mM HEPES/KOH pH 7.4, 150 mM NaCl, 0.05% Tween 20). The ParJ protein injections were followed by two to four injections of 1–5 M NaCl to remove all bound analyte from the sensor chip. The results were plotted as sensograms after subtraction of the background response signal obtained in a control experiment. The BIAevaluation version 4.1 program (Pharmacia Biosensor AB) was used for fitting the curves to calculate dissociation rate kd and dissociation constant KD. KD was additionally calculated from steady-state equilibrium, and was consistent with the results delivered by BIAevaluation software.

ELISA

ParJ (or BSA) was bound to Greiner Microtiter and Maxisorp NUNC ELISA plates at 10 µg ml−1 in 100 µl of coating buffer (0.1 M NaHCO3 pH 9.2) and incubated at 4°C overnight. After rinsing with wash buffer (phosphate-buffered saline, 0.5% Tween 20) and blocking (phosphate-buffered saline, 0.5% Tween 20, 5% BSA), 10–5000 ng of 6His-ParA diluted in 100 µl of blocking buffer was added to each well and incubated at 30°C for 1 h. After washing, a rabbit ParA antiserum and alkaline phosphatase-conjugated goat anti-rabbit IgG antiserum were used for detection of bound ParA. The absorbance of the chromogenic conversion of substrate disodium p-nitrophenyl phosphate hexahydrate (1 mg ml−1) was read at 405 nm in an ELISA plate reader.

Pelleting assay

For pelleting assays, 6.66 µM ParA and 7 µM 6His–ParJ were incubated in reaction buffer in a total of 40 µl (50 mM Tris-HCl pH 7.0, 150 mM NaCl, 5 mM MgCl2) in the presence of ATP (2 mM) for 5 min at room temperature. After incubation at 30°C for 30 min. the samples were centrifuged at 20°C at 100 000 r.p.m. for 30 min in a TLA100 rotor (Beckman, ultra TLX). The supernatants were removed and mixed with gel-loading buffer. The pellets were resuspended in 15–20 µl of reaction buffer and mixed with gel-loading buffer. The whole pellet fraction and half of the supernatant fraction were subjected to SDS-PAGE and stained with Coomassie blue.

Dynamic light scattering

Polymerization of ParA in the presence of ParJ was measured by DLS (DynaPro Wyatt Technology). To observe the effect of ParJ on ParA polymerization 14 µl of protein mixture (3.26 µM ParA and 1.5 µM 6His–ParJ in 50 mM Tris-HCl pH 7.0, 150 mM NaCl, 5 mM MgCl2) was pre-incubated at room temperature for 5 min. Then ATP (2 mM) was added to the protein solution, which was placed in the quartz cuvette and incubated at 30°C in the DynaPro chamber. In a complementary approach 6His–ParJ (0.75, 1.5, 3, 7 µM) was added after 5 min of ParA incubation at 30°C in the chamber. Measurements were taken every 2 s.

ATPase activity assays

The ATPase activity of ParA and 6His–ParJ was assayed by the spectrophotometric detection of inorganic phosphate using a PiPer Phosphate Assay Kit (Molecular Probes) according to the manufacturer's instructions.

ATP binding assay

For the assay 6HisParJ and ParJ cleaved form GST were used. Prior to binding of ATP, ParJ was dialysed against 50 mM Tris (pH 7.0), 150 mM NaCl, 5 mM EDTA, 5% glycerol to remove bound Mg2+ and ATP and later against 50 mM Tris (pH 7.0), 150 mM NaCl, 1 mM EDTA, 5% glycerol. The reaction mixtures contained 6 µM protein, 2.5 µM [α-32P]-ATP (specific activity, 0.2 Ci mmol−1) in the 50 mM Tris (pH 7.0), 150 mM NaCl, 1 mM EDTA, 5% glycerol. Binding was performed for 30 min at room temperature followed by UV cross-linking (8 J) done on ice for 30 min using a UV Crosslinker (Hoefer Scientific Instruments). Reactions were terminated by addition of an equal volume of SDS sample buffer, and the samples were electrophoresed on a 10% SDS-poly-acrylamide gel. The gel was fixed, stained with Coomassie blue, dried and exposed to a phosphorscreen.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

We thank Justin Nodwell for providing S. coelicolor expression library in bacterial two-hybrid system, Jerzy Majka for help with BIAcore data analysis and Gabriella Kelemen for helpful discussion. This work was supported by the Ministry of Science and Higher Education (Grant N N301 285437). B.D. was supported by a scholarship of the President of Polish Academy of Sciences and British Council Young Investigator Programme.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
FilenameFormatSizeDescription
MMI_7409_sm_FigeS1-4_TableS1.pdf3017KSupporting info item

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