Bacillus subtilis pnpA gene product, polynucleotide phosphorylase (PNPase), is involved in double-strand break (DSB) repair via homologous recombination (HR) or non-homologous end-joining (NHEJ). RecN is among the first responders to localize at the DNA DSBs, with PNPase facilitating the formation of a discrete RecN focus per nucleoid. PNPase, which co-purifies with RecA and RecN, was able to degrade single-stranded (ss) DNA with a 3′ → 5′ polarity in the presence of Mn2+ and low inorganic phosphate (Pi) concentration, or to extend a 3′-OH end in the presence dNDP·Mn2+. Both PNPase activities were observed in evolutionarily distant bacteria (B. subtilis and Escherichia coli), suggesting conserved functions. The activity of PNPase was directed toward ssDNA degradation or polymerization by manipulating the Pi/dNDPs concentrations or the availability of RecA or RecN. In its dATP-bound form, RecN stimulates PNPase-mediated polymerization. ssDNA phosphorolysis catalyzed by PNPase is stimulated by RecA, but inhibited by SsbA. Our findings suggest that (i) the PNPase degradative and polymerizing activities might play a critical role in the transition from DSB sensing to end resection via HR and (ii) by blunting a 3′-tailed duplex DNA, in the absence of HR, B. subtilis PNPase might also contribute to repair via NHEJ.
DNA double-strand breaks (DSBs) are highly cytotoxic lesions that arise spontaneously during growth or following exposure to DNA damaging agents, such as H2O2. The rapid detection and subsequent repair of DNA DSBs is critical for the survival of all living organisms. One-ended DSBs are repaired by homologous recombination (HR), whereas the two-ended DSBs are repaired by either HR or non-homologous end joining (NHEJ). HR relies on the presence of a homologous duplex template, whereas NHEJ directly rejoins the two ends of the broken chromosomes by ligation (1–7). NHEJ is error prone and can operate in any phase of the cell cycle in eukaryotes, but in bacteria only when a single chromosome copy is present, or when HR is impaired; therefore, only a minor fraction of two-ended DSBs are expected to be repaired by NHEJ. Whereas HR is error free and constrained to the S and G2 stages in eukaryotes, but is by far the major DSB repair avenue in bacteria (1,3,8,9).
In eukaryotes, among the first responder to DSBs is the MRX(N) complex constituted by Mre11, Rad50 and Xrs2 (in budding yeast) or Nbs1 (in mammals) (4,10). Then, the ends are processed to form the invasive 3′ single-stranded DNA (ssDNA) tails. This processing reaction occurs in two steps, a limited end resection by the MRX(N) complex in concert with Sae2 (CtIP in mammals) (11–14) followed by a processive resection of the 5′-end by the Sgs1(BLM)–Top3(TopoIIIα)–Rmi1(RMI1)–Dna2(DNA2) or Sgs1(BLM)–Exo1(EXO1)–MRX(MRN) complex to form a long 3′ ssDNA tail (long-range end processing) in the presence of the single-strand binding RPA protein (13–20). The 3′ ssDNA tail serves as the substrate for assembly of a Rad51 nucleoprotein filament that searches for the homologous duplex (3,7,21). In bacteria, the presence of a DSB (site-specific or randomly induced) in rec+ cells re-localizes RecN from a diffuse distribution to a discrete focus per nucleoid to form a discrete repair center (RC) per nucleoid (9). In Bacillus subtilis, damage-induced RecN accumulation is readily detected 15–20 min after mitomycin C (MMC) treatment, followed by RecO, RecR and RecA co-localization with RecN-induced RC 10–15 min later (22). After damage recognition by RecN long tracts end resection to generate extensive 3′ single-stranded tails occurs via two routes. The AddAB (counterpart of RecBCD or AdnAB) helicase–nuclease complex or the RecJ ssDNA exonuclease, in concert with a RecQ-like helicase (RecQ or RecS simplified here as RecQ/RecS) and SsbA, process the DNA ends to generate a long 3′-tailed duplex bound by RecA (6,9,21,23). In the absence of processive end resection (in the addAB ΔrecJ background) several RecN foci, rather than a single RC, are present per nucleoid and RecA threads are not observed (24).
Previous studies revealed that (i) at the DSB-induced foci, both eukaryotic MRX(N) and bacterial RecN increase the local concentration of damaged DNA ends by bridging and tethering them, leading to rosette-like structures in vitro (4,25,26); (ii) Mre11 degrades 3′ ends (an Mn2+-dependent activity modulated by Rad50 and Xrs2) (11,12), whereas RecN lacks an end-resection activity (27); (iii) eukaryotic and bacterial Ku, the DNA end-binding subunit of the NHEJ machinery, act as negative regulators of long-range end resection by preventing both eukaryotic Exo1-dependent or bacterial AdnAB-dependent formation of long 3′ ssDNA tails (2,28–30); (iv) an Mn2+-dependent 3′ → 5′ ssDNA exonuclease activity, which co-purifies with RecN, was recently attributed to polynucleotide phosphorylase (PNPase) (27); and (v) the Mre11 or the PNPase activity is dispensable when enzymatic ‘ligatable’ breaks are generated (15,17,27).
PNPase (encoded by pnpA in B. subtilis or pnp in Escherichia coli) is a non-essential multifunctional protein, responsible for Mg2+- and inorganic phosphate (Pi)-dependent 3′ → 5′ processive phosphorolytic degradation of RNA, and template-independent polymerization of rNDPs into RNA (31–36). PNPase, which is generally thought of as an enzyme dedicated to RNA metabolism (37,38), in the presence of either Mn2+ or Fe3+ ions may catalyze template-independent polymerization of dNDPs into ssDNA (39–41). Genetic evidences implicates pnpA (pnp) in the repair of H2O2 damage both in B. subtilis and E. coli (27,42), and this effect might be direct because in pnpA cells there are no significant difference in the relative amounts of transcripts of genes involved in DNA repair and/or repair-by-recombination (27). In vivo analyses revealed that a null B. subtilis pnpA (ΔpnpA) mutation is epistatic to ΔrecN or Δku mutations, which by themselves are non-epistatic, whereas ΔpnpA is non-epistatic to addA or ΔrecJ (27). In vitro B. subtilis PNPase (PNPaseBsu), which co-purifies with RecN, has an Mn2+-dependent 3′ → 5′ exodeoxyribonuclease activity (27). This raises new questions as to whether this multifaceted enzyme is directly implicated in DNA metabolism during DSB repair in bacteria and suggests an early involvement of PNPase after a DSB.
To shed light on the molecular mechanism underlying the transition from RecN DSB sensing to the generation of the recombinogenic 3′-ssDNA ends, we characterized the PNPase enzymes from B. subtilis and E. coli (PNPaseBsu and PNPaseEco), respectively, and the effects of RecN, RecA and SsbA on PNPaseBsu enzymatic activities. E. coli and B. subtilis are separated by more than 1.5 billion years, a time divergence larger than the one between humans and plants. It is thus likely that any conserved PNPase activity should be present in most bacteria.
MATERIALS AND METHODS
Bacterial strains and plasmids
All B. subtilis strains used are isogenic with the rec+ control (BG214) trpC2 metB5 amyE sigB37 xre1 attSPβ attICEBs1. B. subtilis recN:yfp DNA (BG1089) was used to transform the isogenic derivative ΔpnpA (BG993) (27), rendering recN:yfp ΔpnpA (BG1091). The 5′-end of the B. subtilis pnpA gene was fused to the gfp gene leading to gfp:pnpA fusion (pCB800). Linearized pCB800 DNA was used to transform rec+ (BG214) and its isogenic ΔpnpA derivative (BG993) into the amy locus, rendering BG1167 (rec+gfp:pnpA) and BG1169 (ΔpnpA gfp:pnpA). The gfp:pnpA fusion failed to complement the ΔpnpA mutation of strain BG1169. The BG214 strain or E. coli BL21(DE3) strain were used for protein overexpression.
Plasmid pAZ101 and pGM742 have been previously described (43,44). Plasmids pCB890-borne pnpAR399D R400D and pCB891-borne pnpAD493G encode a modified pnpA gene that was constructed by site-directed mutagenesis: CGT of wild type (wt) codons 399 and 400, coding for Arg, was exchanged against GAT that codes for Asp in the expressed protein variant PNPaseBsuR399D R400D equivalent to PNPaseEcoR398D and R399D and GAC of wt codon 493, coding for Asp, was exchanged against GGC that codes for Gly in the expressed protein variant PNPaseBsuD493G equivalent to PNPaseEcoD492G (see 45).
Reagents and protein purification
All chemicals used were reagent grade, dithiothreitol (DTT), rNTPs, dNTP, rNDPs, dNDPs and ddTTP were purchased from Sigma or Promega and [γ-32P]ATP from Amersham Bioscience and then Perkin Elmer. The ssDNA and dsDNA concentrations were measured using extinction coefficients of 1.54 × 10−4 and 8780 M−1 cm−1 at 260 nm, respectively. The 60-nt long ssDNA used were: 5′-AGCT(A)56-3′ (oligo-A56), 5′-AGGCCTCCCGGGATCCGCTAGCAGGAGGAAATTCACCATGGGTAAAGGAGAAGAACTTTT-3′ (oligo-A) and 5′- CTCCTATTATGCTCAACTTAAATGACCTACTCTATAAAGCTATAGTACTGCTATCTAATC-3′ (oligo-F). The 20-nt long oligos complementary to oligo-A at positions 1–20 (oligo-B) and at positions 21–40 (oligo-C) or non-complementary with oligo-A (5′-AGTGACTTTAGTACAGTACC, oligo-E), and the 30-nt long ssDNA complementary to oligo-A at positions 40–60 with a 10-nt overhang (5′-ATGAACCTACGAAAAGTTCTTCTCCTTTAC-3′ (oligo-D), 10 µM each, were annealed to oligo-A (5 µM) in 50 mM Tris–HCl (pH 7.5), 50 mM NaCl, by heating for 5 min at 95°C and slow cooling at room temperature. The 60-nt long oligo-F ssDNA was extended using terminal transferase (Roche) and ddTTP; to reduce to a negligible level the proportion of the oligo with a 3′-OH end, two sequential extension reactions were performed, to render a 61-nt long oligo-F′ ssDNA. The 107-, 105- and 102-bp dsDNA fragments with a 3′ protruding, blunt or 5′ protruding end, respectively, were obtained by PCR amplification of the multiple cloning site of pGM742 with with primers FG953 (CCTGTTTGATGGTGGTTCGGTACG) and FG125 (GACCGCTTCTGCGTTCTGATT), cleaved by either KpnI (3′-protruding), SmaI (blunt) or BamHI (5′-protruding) restriction enzyme, and gel purified. The dsDNA fragments were incubated with 10 nM PNPaseEco or PNPaseBsu in buffer A containing 100 µM dADP for the time indicated at 26°C. The 60- or 61-nt long ssDNA and the dsDNAs were 5′-end-labeled using [γ-32P]-ATP and T4 polynucleotide kinase (New England Biolabs) purified [from the non-incorporated nucleotide] by Sephadex G50 gel filtration. Uniformly labeled [32P]-RNA substrate was synthesized by in vitro transcription with T7 RNA polymerase and [α-32P]-CTP using a DNA template obtained by PCR amplification of plasmid pAZ101 with oligonucleotides FG1626 (taatacgactcactatagggCATTAGCCGCGCGAACCTC, where lowercase letters indicate the T7 promoter, and uppercase letters indicate E. coli sequence) and FG1627 (GAATGATCTTCCGTTGCAGAG) (46).
PNPaseEco, PNPaseBsu and SsbA were over-expressed in E. coli BL21(DE3) cells bearing plasmid borne pnp, pnpA or ssbA genes, purified and stored as previously described (46–48). RecA and RecN were over-expressed in B. subtilis BG214 cells bearing plasmid borne recA or recN gene, purified and stored as previously described (49,50). PNPaseBsuR399D R400D and PNPaseBsuD493G variants were purified using as previously described for PNPaseBsu (47).
Assays of PNPase activities
The 5′-end [32P]-labeled oligo-A56 or oligo-A ssDNA (0.25 nM in molecules) was incubated with PNPaseEco or PNPaseBsu (10 nM) in buffer A [10 mM Tris–HCl (pH 7.5), 10 mM KCl, 2 mM MnCl2, 0.75 mM DTT, 2% PEG-6000], buffer B (as buffer A but with 2 mM MgCl2 instead of MnCl2) or buffer C (as buffer A but lacking MnCl2) containing the indicated concentration of dADP, dCDP, dGDP, dTDP, rATP or dATP, or no nucleotide, with the indicated concentrations of sodium phosphate. The 5′-end labeled [32P]-ssDNA was pre-incubated with RecN and then with increasing concentrations of PNPaseBsu in buffer A containing the indicated concentration of dADP and rATP or dATP. The samples were separated on denaturing 6% or 12% (as indicated) polyacrylamide gel electrophoresis (dPAGE) (46).
Thin layer chromatography of the radiolabeled DNA degradation products was performed on polyethyleneimine–cellulose (PEI) plates with buffer D [0.4 M K2HPO4 (pH 8.0), 0.7 M H3BO3] as the mobile phase as described (51).
To exponentially growing cells (OD560 = 0.5) 50 ng ml−1 of MMC was added and the culture was incubated for 30 min. The cells were collected, the DNA was stained with 4′,6-diamidino-2-phenylindole (DAPI; final concentration 0.2 µg ml−1) and fluorescence microscopy was performed as described (22).
Absence of PNPase abrogates formation of a discrete RecN focus per nucleoid
DNA-damage response recruits complex molecular machinery involved in DNA repair to DSBs. In vivo, B. subtilis RecN is among the first responders leading to the formation of a discrete RC per nucleoid (22), and a null pnpA (ΔpnpA) mutation is epistatic to ΔrecN (27). In vitro, RecN-bound (d/r)ATP recognizes ssDNA ends and tethers them onto a nucleoprotein complex, rosette-like structure (26,50). To determine whether PNPaseBsu acts in concert with RecN, the fate of the RecN:YFP protein was studied by fluorescence microscopy in pnpA+ and ΔpnpA strains expressing a recN:yfp fusion upon MMC treatment. RecN:YFP appeared evenly distributed within the cells in both pnpA+ and ΔpnpA exponentially growing cultures (Figure 1). Thirty minutes upon addition of MMC (50 ng ml−1), DNA damage-induced RecN foci were detected in both pnpA+ (72%, n = 808) and ΔpnpA (70%, n = 491) cells (Figure 1). pnpA+ cells generally contained a discrete RecN:YFP focus per nucleoid in the majority of cells (~69%, n = 580), whereas two or more foci were rarely observed (~3% of the cells) (22) (Figure 1A). In contrast, a large fraction of ΔpnpA cells contained two (37%, n = 350) or multiple foci that appeared patchy (11%, n = 350), whereas a discrete focus was observed in ~22% of the cells (Figure 1B). RecN:YFP can thus form foci at DSBs early upon DNA damage in the absence of PNPaseBsu, whereas the efficient formation of a single RC appears to require an active PNPaseBsu enzyme.
We also monitored the distribution of GFP-tagged PNPaseBsu in the absence and presence of MMC (Supplementary Figure S1). In ~80% of the cells (n = 665 cells), GFP:PNPaseBsu was found either to form foci at the cell poles (14–18%) (Supplementary Figure S1A, left panels), or distributed within the cell (Supplementary Figure S1B) in a way that was reminiscent of the filamentous helical structures (62–66%) previously described for E. coli PNPase (52); the remaining fraction showed fluorescence distributed throughout the cells or no fluorescence at all. The frequency of foci and helical structures (14–18% and 62–66%, respectively), however, was the about same both in untreated exponentially growing cells and upon addition of MMC.
PNPaseEco catalyzes Mn2+-dependent phosphorolysis
PNPaseBsu is an Mn2+-dependent 3′ → 5′ ssDNA exonuclease (27). To shed light into the role of PNPase in DNA processing, we compared in vitro exonucleolytic DNA degradation by purified E. coli and B. subtilis enzymes on a 60-nt long ssDNA potentially forming secondary structures (oligo-A) or an unstructured (oligo-A56) ssDNA. Both PNPases degraded oligo-A56 ssDNA with a 3′ → 5′ polarity in the presence of Mn2+ and low Pi (10 µM) (Figure 2A, lanes 3–12 and 3A). Independent of protein concentration, PNPaseEco could run through the entire molecule whereas PNPaseBsu could not proceed more than ~25-nt (Figure 3B, lanes 4–6 and 10–12). When oligo-A ssDNA (Figure 3D) was used, PNPaseBsu only nibbled off 2-nt, whereas PNPaseEco could run through the entire molecule, although pausing at preferential sites (Figure 2B and 3C, lanes 3–5 and 8–10). Such differences could not be imputed to different specific activities of the two PNPase preparations, as no similar effects were observed when RNA was degraded (Supplementary Figure S2). Overall these data suggest that PNPaseBsu is less processive or efficient at DNA degradation and more sensitive to DNA secondary structures than PNPaseEco.
ssDNA degradation occurred in the presence of low or even in the absence of supplemented Pi (27) (Figure 3A). This was likely due to contaminating Pi, since degradation of [α-32P]-dATP uniformly labeled DNA by PNPase generated dADP both with and without added Pi, as shown by thin layer chromatography (Supplementary Figure S3). Thus PNPase promoted genuine ssDNA phosphorolysis. Unlike RNA phosphorolysis, which occurs at Pi concentrations as high as 10–30 mM (53), the Mn2+-dependent DNase activity of both PNPases was completely inhibited by 100 µM or higher Pi concentrations (27) (Figure 2C). No ssDNA degradation could be detected when Mn2+ was replaced by Mg2+ (27) (Supplementary Figure S4A, lanes 3–12).
Bacterial PNPase catalyzes template-independent dNDP polymerization into ssDNA
In the presence of Mn2+ and dADP, both PNPases catalyzed template-independent ssDNA polymerization on both unstructured oligo-A56 and structured oligo-A ssDNA (Figure 2A and B). High dADP concentrations (1 mM) (or contaminating Pi possibly present in dADP commercial stocks), however, inhibited polymerization (Figure 2C, lanes 19 and 25).
In the presence of Mg2+, template-independent polymerization occurred with a much lower efficiency than with Mn2+ (compare Figure 2, lanes 13–22 with Supplementary Figure S4A, lane 13–22). PNPaseEco was able to catalyze template-independent polymerization with oligo-A as a primer in the presence of Fe3+, confirming a previous report (41), although less efficiently than with Mn2+ (compared Figure 2B, lanes 13–17 with Supplementary Figure S4A, lanes 2–7). When Fe3+ was replaced by Fe2+ polymerization was almost negligible (Supplementary Figure S4B). PNPase-mediated polymerization reaction was inhibited when Fe ions exceeded the 0.5 mM concentration (Supplementary Figure S4B).
All four dNDPs could be used as substrates for polymerization, although with different reaction rates, for both PNPases (dCDP ≈ dADP >> dTDP ≈ dGDP) (Supplementary Figure S5).
Polymerization of dADP by PNPase was stimulated by sub-stoichiometric concentrations of rADP but at equimolar or higher concentrations rADP exerted a negative effect (Supplementary Figure S6). In such conditions rADP was incorporated into co-polymers, as shown by alkali degradation of the polymerization products (Supplementary Figure S6).
To unambiguously determine the function of PNPase and to examine whether the amino acids important for RNA phosphorolysis and NDP polymerization activities were also important for ssDNA phosphorolysis and dNDP polymerization we created two site-specific PNPaseBsu mutants. The PNPaseBsuR399D R400D and PNPaseBsuD493G variants, which correspond to the catalytically inactive PNPaseEcoR398D R399D and PNPaseEcoD492G proteins, respectively (45), were purified in parallel with wt PNPase, using the same purification protocol. Both PNPaseBsu mutant proteins were unable to catalyze degradation or template-independent ssDNA polymerization even when the mutant variants were present at a 1000-fold higher concentration than wt PNPaseBsu (Supplementary Figure S7). These data indicate that both RNA and ssDNA share the same catalytic site.
Bacterial PNPase can process a 3′-tailed duplex
To test whether PNPase was able to process duplex DNA with a 3′-protruding or recessed end, oligo-A ssDNA was annealed with a 2-fold excess of oligos complementary to different portions of the labeled probe (oligo-B, -C or -D) or the mock non-complementary oligo-E (Figure 4). PNPaseEco degraded 40-nt (A:B) or 20-nt (A:C) long 3′-protruding ends, but failed to degrade 3′-recessed ends (A:D) (Figure 4A, lane 16–20).
Both PNPases extended dsDNA digested with a restriction enzyme leaving a 4-nt long 3′-protruding, a blunt end or a 5′- protruding end (Figure 4B). Template-independent ssDNA polymerization by PNPase was significantly enhanced if the length of the tailed ssDNA was increased. Both PNPaseEco (Figure 4C) and PNPaseBsu (Figure 4D) efficiently extended the 3′-protruding ends of partially dsDNA substrates. Elongation, however, was less efficient with a 3′-recessed end (A:D) substrate. PNPaseBsu seemed to be more proficient than PNPaseEco with this substrate (Figure 4C and D). The lower efficiency of polymerization with a dsDNA primer with short or no tail as compared with the partially dsDNA oligos (Figure 4C and D) might depend on the lower affinity of PNPase for dsDNA (27,54) and the preference for a ssDNA tail as primer.
rATP and dATP modulate the PNPase activities
Biochemical analyses revealed that (d/r)ATP is an allosteric inhibitor of PNPaseEco RNA degradation and polymerization activities (46) and of PNPaseBsu ssDNA degradation activity (27). To address whether (d/r)ATP affected both PNPase activities on ssDNA, assays were performed in the presence of (d/r)ATP·Mn2+. PNPase ssDNA degradation was inhibited by the presence of 1 mM rATP·Mn2+, and addition of even lower rATP concentrations (100 µM) also inhibited dADP polymerization (Supplementary Figure S8A). Likewise, dATP inhibited both phosphorolysis and polymerization by PNPase (Supplementary Figure S8B). Interpretation of such results is complicated by the fact that r/dNTPs are unavoidably contaminated by Pi and r/dNDPs (see Supplementary Figure S3, lanes 5 and 6), which by themselves modulate PNPase activities. This, however, could not explain the inhibitory effect in both directions by the same nucleotide preparation; moreover, the inhibitory effects of dATP was observed at very low dATP concentrations (10 and 1 µM for degradation and polymerization, respectively) versus concentrations of 10 µM Pi and 100 µM dADP as substrates, where concentration of degradation byproducts may be negligible. In addition, although contaminating Pi might directly inhibit degradation of ssDNA (see above), this is considered unlikely because contaminating Pi concentrations of 100 µM or higher are needed for inhibition of the degradative activity (see above). Thus inhibition of PNPase by r/dATP seems the more reasonable explanation of the bulk of these data.
RecN and SsbA modulate PNPase polymerization into ssDNA
In vivo analyses in B. subtilis revealed that DNA damage-induced RecN to form a discrete focus per nucleoid (22). In vitro, RecN is a ssDNA-dependent (r/d)ATPase (50). In the (r/d)ATP or (r/d)ADP bound form, RecN, assembles at the 3′-OH ends to form large networks with ssDNA molecules (rosette-like structures) that are insensitive to ExoIII degradation (26,50). Addition of SsbA (counterpart of E. coli SSB or eukaryotic RPA) does not disrupt the rosette-like structures (26). As shown in Figure 1, PNPaseBsu, which co-purifies with RecN (27), altered the formation of a discrete RecN focus per nucleoid. As we failed to detect stable physical PNPaseBsu and RecN interactions in solution using co-immunoprecipitation or protein cross-linking methodologies, we tested whether RecN could modulate any of the PNPase activities.
RecN alone neither polymerized ssDNA (Figure 5A, lane 3) nor degraded ssDNA (27). RecN had two different effects on PNPase-mediated template-independent polymerization. First, RecN·dATP stimulated ~6-fold ssDNA extension when incubated with limiting PNPaseBsu concentrations (e.g. as low as 0.2 nM) (Figure 5A and B). Second, the presence of RecN abolished the inhibitory effect exerted by 1 mM dATP on dADP polymerization (Figure 5A and B, lane 13). It is possible that RecN-mediated rosette-like structures facilitate PNPaseBsu access to a high local concentration of ssDNA ends, and that RecN catalyzed dATP hydrolysis, rather than any other ssDNA-dependent dATPase, contributed to the observed stimulation. As expected RecN failed to activate the catalytic site mutants PNPaseBsuR399D R400D or PNPaseBsuD493G (Supplementary Figure S7A).
RecN binds to the ssDNA ends in a dATP-dependent manner and tethers and bridges them, and such rosette-like structures are insensitive to SsbA addition (26,50). SsbA, at stoichiometric concentrations (1 SsbA tetramer/80-, 40- or 20-nt), showed a significant inhibitory effect on PNPase-catalyzed ssDNA polymerization when compared with the absence of SsbA (Figure 5B, lane 5 versus 6–8). In the presence of RecN·dATP, SsbA inhibited PNPaseBsu polymerizing activity less severely (Figure 5B, lanes 6–8 versus 11–13).
RecA and SsbA modulate PNPase degradation of ssDNA
In the presence (d/r)ATP·Mg2+, RecA assembles onto ssDNA to form an extended RecA·ssDNA filament and promotes strand exchange with homologous dsDNA, whereas in the apo form or in the presence of ADP, RecA can bind ssDNA and form a compact filament inactive for RecA-mediated DNA strand exchange (55,56). RecA catalyzes the hydrolysis of dATP and, to a lesser extent, rATP in a ssDNA-dependent manner (48,57). PNPaseEco co-purifies with RecAEco (58) and an interaction between both proteins was predicted (www.compsysbio.org/bacteriome). We failed, however, to detect a stable physical interaction between B. subtilis PNPase and RecA proteins, although an in vivo interaction between PNPase and the RecA-like domain of an RNA helicase was documented by a two-hybrid system (59). To determine whether RecA or RecA and SsbA affect the PNPaseBsu activities, we tested the ability of PNPaseBsu to degrade ssDNA or to polymerize into ssDNA. In the apo or in the nucleotide-bound form RecA, at stoichiometric concentrations (1 RecA/3-nt), neither degrades (see also Figure 6B) nor stimulates template-independent polymerization into ssDNA (21,60). As shown above (Figures 2B and 3C) PNPaseBsu could nibble only a few nucleotides of the structured ssDNA substrate. At very low Pi concentration (no exogenously added Pi), RecA markedly stimulated PNPase-catalyzed ssDNA phosphorolysis >10-fold, leading to extensive degradation of the probe (Figure 6A, compare lanes 2–4 with 8–10). ssDNA degradation by PNPaseBsu accumulates dNDPs, hence we could not discriminate whether RecA exerted the stimulatory effect on PNPase-mediated ssDNA degradation in the apo or in the dADP-bound form.
dATP, which is critical for the formation of an active RecA·ssDNA filament (48,57), limited the extent of PNPase-mediated 3′-end resection (Supplementary Figure S8, lanes 6 and 12). dATP also limited the stimulatory effect exerted by RecA on DNA resection at low Pi concentration and reduced ssDNA degradation in 10 µM Pi (Figure 6A, lanes 11–13 and 23–25). These data suggest that RecA stimulates ssDNA degradation by PNPaseBsu perhaps by melting out secondary structures in the DNA, but RecA fails to overcome the inhibitory effect exerted by dATP on PNPase-mediated 3′-end resection. It is also possible that the products of RecA-mediated dATP hydrolysis and accumulation of Pi concentration higher than 10 µM might limit the extent of degradation. The catalytic site mutants PNPaseBsuR399D R400D or PNPaseBsuD493G could not be activated by the addition of RecA (Supplementary Figure S7B).
Addition of RecA in the apo bound form facilitated full degradation of the ssDNA substrate (Figure 6B, lanes 5 and 10). Addition of SsbA (1 SsbA tetramer/80-, 40- or 20-nt) inhibited PNPaseBsu degradation of ssDNA (Figure 6B, lanes 6–9). Similar inhibition was observed in the presence of RecA. It is likely that SsbA (1 SsbA tetramer/20-nt) inhibits PNPase-mediated degradation of ssDNA and RecA, in the apo or dADP-bound form, cannot overcome such inhibitory effect (Figure 6B, lanes 11 and 13). It is likely that SsbA poses a barrier to PNPase-mediated degradation of ssDNA.
RecA stimulates PNPase degradation of ssDNA
To examine whether RecA affected PNPaseBsu degradation of a ssDNA lacking a 3′OH end we extended the 60-nt ssDNA with ddTTP, rendering a 61-nt long ssDNA with a ddTMP incorporated at the 3′-end. When the 60-nt long ssDNA with a 3′-OH end was used, PNPaseBsu only nibbled off 2-nt (Figure 7, lane 2–4), confirming that PNPaseBsu is sensitive to DNA secondary structures (Figure 3D). On the contrary, PNPaseBsu failed to degrade the 3′-deoxy 61-nt long ssDNA (Figure 7, lanes 6–8); however, the presence of RecA stimulated PNPase-catalyzed phosphorolysis of 3′-deoxy-terminated ssDNA (Figure 7, lanes 9–11). It thus appeared that PNPaseBsu can degrade a 3′-end lacking a 3′-OH group on its sugar moiety in the presence of RecA.
In this study, we biochemically characterized the Mn2+-dependent 3′ → 5′ phosphorolytic degradation of ssDNA and template-independent polymerization of dNDPs into ssDNA activities of E. coli or B. subtilis PNPase (27, this work). PNPase, which is a multifunctional enzyme, is also responsible for Mg2+-dependent 3′ → 5′ processive phosphorolytic degradation of RNA and template-independent polymerization of rNDPs into RNA (31–36). It is likely that PNPase plays a dual role in response to genotoxic agents, participating both in RNA turn-over (31–36) and in facilitating DNA DSB repair (27). It emerges that PNPase is a versatile enzyme that can be directed toward either RNA or DNA metabolism by metal ions. Its reversible activity not only is determined by substrate (Pi and r/dNDP's) concentration, but also is modulated by nucleotide triphosphates cofactors and can associate with different proteins to build up machines dedicated to cellular functions apparently as diverse as RNA turnover and DNA repair and recombination (9,27).
Genetic data suggest that PNPaseBsu is required for repair of H2O2-induced DSB repair via RecN-dependent HR or Ku-dependent NHEJ, whereas it is dispensable for MMC-induced DSB repair (27). H2O2-induced DSB can chemically alter DNA ends, whereas MMC-induced crosslinks and subsequently the generated DNA DSBs have clean ends. It is likely that PNPase plays an active role when the ends are chemically modified although the resection reaction may proceed more slowly. A similar role has been attributed to Mre11 [reviewed by (5)]. Our cytological results suggest that PNPaseBsu is not involved in RecN foci formation. RecN forms a discrete focus per nucleoid in pnpA+ cells and few foci and patchy structures in pnpA− cells (Figure 1). A similar effect was observed in the absence of long-range resection in the addA and ΔrecJ background (24), suggesting that PNPaseBsu, which co-purifies with RecN (27), contributes to RecN-mediated RCs before and/or concomitantly with long-range end-processing by the AddAB or the RecJ-RecQ/RecS enzymes. This is consistent with the observation that ΔpnpA is epistatic to the ΔrecN or Δku mutations, which by themselves are non-epistatic, whereas ΔpnpA is non-epistatic to addA or ΔrecJ (27).
Our biochemical results demonstrate that RecN stimulates Mn2+-dependent PNPaseBsu polymerization on 3′-OH ends. It is likely that RecN, by binding to 3′-OH ends in an ATP-dependent manner and tethering and bridging them (26), facilitates PNPase-mediated template-independent polymerization (Figure 5A and Supplementary Figure S7A) at the local high concentration of DNA ends (rosette-like structures). RecA, even in the dADP bound form, promotes disassembly of the RecN-induced rosette-like structures and stimulates Mn2+-dependent 3′ → 5′ degradation of ssDNA by PNPaseBsu (Figure 6A and Supplementary Figure S7B). Many bacteria accumulate Mn2+, and high intracellular levels of Mn2+ directly or indirectly allow fast repair of damaged DNA after DSBs (61,62).
A molecular model for PNPase role in DSB repair
What could be the mechanistic role of PNPase? The responses of eukaryotic and bacterial cells to DNA DSBs show some similarities and clear differences. The eukaryotic MRN(X) complex and bacterial RecN are among the first responders to DNA DSBs (10,22,63); both Rad50, as part of the MRN(X) complex, and RecN bind to the DNA ends in an ATP-dependent manner and tether and bridge them (25,26). PNPaseBsu, which might work at early stages of DNA DSB repair process, may be implicated in the repair of two-ended DSBs that are created by H2O2, which leaves non-ligatable termini. This is consistent with the observation that a pnpA mutation is epistatic to recN and ku mutations, which by themselves are non-epistatic (27). Biochemical analyses suggested that in the presence of Mn2+ and dNDPs, PNPase elongates short 3′-protruding, blunt or, with lower efficiency, 3′-recessive ends. In the dATP bound form, RecN enhances PNPaseBsu polymerization even on SsbA-coated ssDNA. It is likely that the RecN interaction with the PNPase-extended 3′-OH ends and SsbA bound to the recombinogenic 3′-ssDNA regions limit the access of the Ku protein to the DNA ends, thus directing the reaction towards error-free DSB repair via HR. This is consistent with the observation that: (i) RecN bound to the 3′-OH ends protects them from ExoIII degradation (50), but facilitates PNPase-mediated extension of the 3′-OHs, and (ii) PNPaseBsu, by adding template independent tails to a DSB end might block NHEJ and indirectly constitutes a control point in DNA repair via HR. In contrast, in the absence of RecN and long-range end processing, PNPaseBsu, or any other ssDNA exonuclease, might degrade the 3′-OHs and indirectly facilitate Ku-mediated NHEJ, suggesting that PNPase might contribute in repair pathway choice. This is consistent with the observation that ΔpnpA and Δku are epistatic (27).
Is end processing conserved among bacteria? B. subtilis RecN is essential for the repair of DNA breaks (64). Like PNPaseBsu (see above), E. coli RecN is dispensable for the repair of ‘clean’ breaks created by EcoRI endonuclease (65), but essential for the repair of breaks created by ionizing radiation or bleomycin (66,67), which can chemically alter DNA ends (68). It is likely that bacterial RecN in concert with PNPase plays an active role when the ends are chemically modified. Similarly to the MRX(N)-Sae2(CtIP) complex (15,17,69), PNPaseBsu might contribute to the formation of an early intermediate for long-range end resection. Both Mre11 (11–14) and PNPaseBsu (27, this work) show an Mn2+-dependent 3′ → 5′ exonuclease activity. PNPase degradation was observed with 3′-tailed duplex DNA (Figure 2), whereas degradation was inefficient or it did not occur on a 3′-recessed end (Figure 4A). In E. coli, PNPase co-purifies with RecA (58). RecA stimulates PNPase 3′-end degradation by a mechanism that neither requires ATP hydrolysis nor formation of an active filament. This stimulatory effect on PNPase-mediated degradation ssDNA takes place even in the absence of a 3′-OH end. This basal end processing might facilitate the removal of ‘non-ligatable’ 3′-ends prior to long-range resection of the 5′ strand of DNA by RecJ-RecQ/RecS or the AddAB complex. PNPaseBsu degradation of 3′-ends might contribute to process the 3′-tailed ends derived from collapsed replication forks with nicks on the lagging strand, because end-processing by the AddAB(RecBCD) complex requires blunt or nearly blunt ends (23). This is consistent with the observation that B. subtilis lacks many 3′ → 5′ ssDNA exonucleases (such as ExoI, ExoIX or ExoX) that are present in E. coli (27).
Based on our results we propose that in B. subtilis RecA plays an early role in DSB repair by dislodging RecN from the overhangs of duplex DNA molecules (26,50) and promoting PNPase degradation prior to long-range resection. However, the potential implication of PNPase alone or in concert with RecA in the generation of the proper substrate for AddAB or RecJ-RecQ/RecS activity remains to be addressed (26,27,50).
PNPase activity is modulated by (d/rATP)
A nucleotide triphosphate allosterically inhibits all PNPase activities on RNA and ssDNA (46, this work), but its hydrolysis products are substrates for phosphorolytic degradation (Pi) and polymerization (r/dADP) on both RNA and ssDNA substrates (27,31,33, this work). The RecN and RecA modulators of PNPaseBsu activities on ssDNA are genuine ssDNA-dependent ATPases (21,50) that lower the local concentration of the nucleotide triphosphate and therefore release the allosteric inhibition. This fits with the previously proposed idea that RNA helicases (which share a RecA-like motif) associated with PNPase at the degradosome may modulate PNPase activities by decreasing the local concentration of ATP (46).
Supplementary Data are available at NAR Online
Funding for open access charge: Ministero dell'Università e della Ricerca and Università degli Studi di Milano (PRIN 2007 - 20074CNBJ2); and Universitàdegli Studi di Milano (FIRST 2007 and PUR 2008) (to G.D.).
Conflict of interest statement. None declared.
The authors thank P.L. Graumann for introducing us in the field of fluorescence microscopy, D. Bechhofer for plasmid-borne pnpA gene, and L. Blanco for critical reading of the manuscript.