PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of jbacterPermissionsJournals.ASM.orgJournalJB ArticleJournal InfoAuthorsReviewers
 
J Bacteriol. 2017 February 1; 199(3): e00739-16.
Published online 2017 January 12. Prepublished online 2016 November 28. doi:  10.1128/JB.00739-16
PMCID: PMC5237116

Characterization of Runella slithyformis HD-Pnk, a Bifunctional DNA/RNA End-Healing Enzyme Composed of an N-Terminal 2′,3′-Phosphoesterase HD Domain and a C-Terminal 5′-OH Polynucleotide Kinase Domain

Richard L. Gourse, Editor
Richard L. Gourse, University of Wisconsin—Madison;

ABSTRACT

5′- and 3′-end-healing reactions are key steps in nucleic acid break repair in which 5′-OH ends are phosphorylated by a polynucleotide kinase (Pnk) and 3′-PO4 or 2′,3′-cyclic-PO4 ends are hydrolyzed by a phosphoesterase to generate the 5′-PO4 and 3′-OH termini required for sealing by classic polynucleotide ligases. End-healing and sealing enzymes are present in diverse bacterial taxa, often organized as modular units within a single multifunctional polypeptide or as subunits of a repair complex. Here we identify and characterize Runella slithyformis HD-Pnk as a novel bifunctional end-healing enzyme composed of an N-terminal 2′,3′-phosphoesterase HD domain and a C-terminal 5′-OH polynucleotide kinase P-loop domain. HD-Pnk phosphorylates 5′-OH polynucleotides (9-mers or longer) in the presence of magnesium and any nucleoside triphosphate donor. HD-Pnk dephosphorylates RNA 2′,3′-cyclic phosphate, RNA 3′-phosphate, RNA 2′-phosphate, and DNA 3′-phosphate ends in the presence of a transition metal cofactor, which can be nickel, copper, or cobalt. HD-Pnk homologs are present in genera from 11 bacterial phyla and are often encoded in an operon with a putative ATP-dependent polynucleotide ligase.

IMPORTANCE The present study provides insights regarding the diversity of nucleic acid repair strategies via the characterization of Runella slithyformis HD-Pnk as the exemplar of a novel clade of dual 5′- and 3′-end-healing enzymes that phosphorylate 5′-OH termini and dephosphorylate 2′,3′-cyclic-PO4, 3′-PO4, and 2′-PO4 ends. The distinctive feature of HD-Pnk is its domain composition, i.e., a fusion of an N-terminal HD phosphohydrolase module and a C-terminal P-loop polynucleotide kinase module. Homologs of Runella HD-Pnk with the same domain composition, same domain order, and similar polypeptide sizes are distributed widely among genera from 11 bacterial phyla.

KEYWORDS: 3′ phosphatase, nucleic acid repair, polynucleotide kinase

INTRODUCTION

Polynucleotide kinases (Pnks) are a widely distributed class of cellular and virus-encoded nucleic acid repair enzymes that convert 5′-OH termini into 5′-PO4 ends that can be sealed by RNA or DNA ligases. Pnks are members of the P-loop phosphotransferase superfamily; they catalyze metal-dependent transfer of the γ phosphate of a nucleoside triphosphate (NTP) donor to a 5′-OH polynucleotide acceptor. In many repair systems, a Pnk enzyme is fused in a modular fashion to one or more other repair enzymes within a single multifunctional polypeptide.

For example, bacteriophage T4 encodes a bifunctional 5′-OH polynucleotide kinase-3′-phosphatase (Pnkp) consisting of an N-terminal Pnk domain fused to a C-terminal phosphoesterase domain of the DXDXT acylphosphatase superfamily (1,4). T4 Pnkp is the end-healing component of a tRNA repair system that includes the separately encoded T4 RNA ligase 1 (Rnl1). 3′ end healing by T4 Pnkp entails transformation of a 2′,3′-cyclic phosphate (indicated by >p) into a ligatable 3′-OH or 2′-OH end (5). A homologous bifunctional mammalian DNA repair enzyme that heals 3′-PO4/5′-OH breaks is composed of an N-terminal DXDXT acylphosphatase domain fused to a C-terminal Pnk domain (6, 7). The eukaryal baculovirus Autographa californica nuclear polyhedrosis virus (AcNPV) encodes a trifunctional RNA repair enzyme composed of an N-terminal Rnl1-like RNA ligase domain, a central Pnk domain, and a C-terminal DXDXT acylphosphatase domain, which executes the full suite of RNA end-healing and sealing reactions (8). The bacterium Capnocytophaga gingivalis elaborates an RNA repair complex composed of three polypeptides, one of which is a bifunctional Pnkp that is homologous to T4 Pnkp (i.e., it has a C-terminal DXDXT acylphosphatase domain) and is encoded in a two-gene operon with an RNA ligase (9).

In other repair systems, Pnks are fused to different types of 3′-end-healing enzymes. For example, the RNA repair system of Clostridium thermocellum (recapitulated in scores of diverse bacterial taxa) includes a trifunctional enzyme composed of an N-terminal Pnk domain, a central 2′,3′-phosphoesterase domain of the binuclear metallophosphoesterase superfamily, and a C-terminal RNA ligase domain (Rnl5 type) (10,20). In yet another variation, the tRNA splicing systems of fungi and plants exploit a trifunctional RNA repair enzyme consisting of an N-terminal RNA ligase domain (Rnl1 type), a central Pnk domain, and a C-terminal 2′,3′-cyclic phosphodiesterase (CPDase) domain that belongs to the 2H phosphoesterase superfamily (21,24).

We suspect that the level of diversification in the organization of nucleic acid end-healing and sealing enzymes described above is just the tip of the iceberg. Consequently, our laboratory aims to discover and to characterize novel enzymes that either reveal new principles of end modification and ligation chemistry (e.g., references 25,27) or illuminate distinctive organizations of catalytic modules within multidomain repair systems. Here we characterize HD-Pnk, a new bifunctional 3′- and 5′-end-healing enzyme from the bacterium Runella slithyformis that consists of an N-terminal 2′,3′-phosphoesterase module of the HD domain family fused to a C-terminal Pnk module (Fig. 1). HD domain enzymes are widely distributed binuclear metal-dependent phosphodiesterases and monoesterases that act on nucleic acids, nucleotides, and other phosphate-containing substrates (28,41) in the service of a broad spectrum of biological functions.

FIG 1
Genomic context and primary structure of Runella HD-Pnk. (Top) The ORF encoding Runella slithyformis HD-Pnk is preceded by three cooriented upstream ORFs encoding putative enzymes with likely roles in RNA or DNA repair. (Bottom) The amino acid sequence ...

The suggestion of a nucleic acid repair function for HD-Pnk is fortified by the location of the enzyme within a cooriented gene cluster of the Runella slithyformis chromosome that encodes other candidate repair enzymes, including (i) a polynucleotide ligase gene immediately upstream of the HD-Pnk gene, in a putative repair operon in which the 3′ end of the ligase open reading frame (ORF) overlaps the 5′ end of the HD-Pnk ORF, and (ii) an upstream two-gene cassette, separated from the ligase ORF by a 152-nucleotide spacer, that encodes RNA 2′-phosphotransferase (Tpt1) and a predicted bifunctional S-adenosylmethionine (AdoMet)-dependent methyltransferase (MTase)-histidine triad (HIT) phosphotransferase enzyme (Fig. 1). Tpt1 is the enzyme that, in yeast, removes a 2′-phosphate at the splice junction of yeast tRNAs (42). The HIT family includes repair enzymes that resolve abortive nucleic acid reaction intermediates generated by polynucleotide ligases (43).

Our interest in HD-Pnk was enhanced by the fact that a ligase-HD-Pnk operon is present in diverse bacterial taxa, including pathogenic strains of Escherichia coli. In order to place HD-Pnk within a nucleic acid repair framework, we set out (i) to produce and to purify recombinant Runella HD-Pnk, (ii) to investigate whether it has 3′- and 5′-end-healing activities in vitro (it does), (iii) to analyze whether the 3′- and 5′-end-healing activities are catalyzed by the HD and Pnk domains, respectively, and are functionally separable (they are), and (iv) to determine the cofactor requirements and substrate preferences of the 5′- and 3′-end-healing activities.

RESULTS

HD-Pnk as a candidate end-healing enzyme.

The 368-amino-acid Runella HD-Pnk polypeptide and its 370-amino-acid homolog from E. coli strain UTI89 share 238 positions of side chain identity or similarity (Fig. 1). The C-terminal half of HD-Pnk contains two hallmark motifs found in polynucleotide kinases, i.e., (i) the GXXGXGK “P-loop,” which engages the phosphates of the NTP donor substrate via hydrogen bonds from the P-loop lysine ζ nitrogen and several of the P-loop main-chain amide nitrogens, and (ii) a DXXR motif that engages the 5′ HONp nucleotide of the phosphate acceptor, such that the aspartate (Asp254 in HD-Pnk) acts as a general base catalyst to activate the O5′ nucleophile for its attack on the NTP γ phosphorus and the arginine coordinates the 3′ phosphate (14, 15). The N-terminal half of HD-Pnk contains the signature HDXXK motif of HD phosphoesterases, which coordinates the metal cofactors via the histidine and aspartate side chains and the scissile phosphoester via the lysine side chain (35, 39).

Polynucleotide kinase activity of recombinant HD-Pnk.

To assess what biochemical activities, if any, are associated with Runella HD-Pnk, we produced in E. coli and purified recombinant wild-type (WT) HD-Pnk and three mutants, in which alanines replaced predicted Pnk active site residues Asp254 and Arg257 and the HD motif His73 residue (Fig. 2A). Recombinant Runella HD-Pnk proteins were produced as His10-Smt3 fusions and were isolated from soluble bacterial extracts by adsorption to Ni-agarose resin and elution with imidazole. The His10-Smt3 tag was removed with the Smt3-specific protease Ulp1, and the tagless HD-Pnk proteins were separated from the tag by a second round of Ni-agarose chromatography. A final purification step of Superdex-200 gel filtration showed that the 43-kDa HD-Pnk polypeptide eluted as a single component at a position (relative to size standards) consistent with its being a monomer in solution (Fig. 2C). SDS-PAGE analysis highlighted the equivalent purity of the recombinant wild-type and mutant HD-Pnk preparations (Fig. 2A).

FIG 2
Polynucleotide kinase activity of recombinant HD-Pnk. (A) WT and mutant HD-Pnk proteins. Aliquots (5 μg) of the recombinant WT HD-Pnk and the H73A, D254A, and R257A mutants were analyzed by SDS-PAGE. The Coomassie blue-stained gel is shown. The ...

A polynucleotide kinase activity of wild-type HD-Pnk was demonstrated by incubating the protein with magnesium, [γ-32P]ATP, and a 21-mer 5′-OH DNA oligonucleotide; this resulted in label transfer from ATP to the 21-mer DNA, as assessed by denaturing PAGE and autoradiography (Fig. 2B). The kinase activity was effaced by the D254A and R257A mutations in the Pnk DXXR motif but was not affected by the H73A mutation in the HD motif (Fig. 2B). The extent of DNA phosphorylation was proportional to the HD-Pnk concentration (Fig. 2D). From the slope of the titration curve, we estimated a specific activity of 50 pmol of 5′ ends phosphorylated per pmol of wild-type enzyme. Whereas the specific activity of the H73A mutant was similar to that of the wild-type enzyme, the D254A and R257A mutants were effectively inert (Fig. 2D). We conclude that the Pnk activity observed inheres to the recombinant HD-Pnk protein and relies on enzymatic components that are conserved and essential in other Pnk enzymes.

We used Tris-acetate (pH 6.0) for all subsequent HD-Pnk kinase assays in this study, after determining that the kinase was active over a broad pH range of 4.5 to 9.5 in Tris-acetate or Tris-HCl buffers. DNA phosphorylation required a divalent cation (Fig. 2E), with optimal activity at 0.5 to 5 mM magnesium. A comparison of magnesium, manganese, calcium, nickel, cobalt, and zinc at 5 mM showed that magnesium was the preferred metal cofactor (Fig. 2E).

Runella slithyformis is a psychrotolerant mesophile of the phylum Bacteroidetes that grows at temperatures as low as 4°C; its optimal growth temperature is 20°C to 30°C (44). The temporal profile of the HD-Pnk DNA kinase reaction was assessed at three different enzyme concentrations (25 nM, 100 nM, and 200 nM HD-Pnk), in reactions performed in parallel at 22°C and 37°C. The initial rates were proportional to the enzyme concentration and did not differ at 22°C versus 37°C (data not shown). The HD-Pnk DNA kinase activity displayed a hyperbolic dependence on the ATP concentration, with an apparent Km of 104 μM ATP and a kcat of 11 min−1 (Fig. 2F).

Effects of oligonucleotide chain length on DNA kinase activity.

We assayed the HD-Pnk-catalyzed transfer of 32P from 100 μM [γ-32P]ATP to a series of unlabeled 5′-OH DNA oligonucleotides (10 μM) of different lengths but identical 5′-terminal nucleobase sequences (Fig. 3). The reaction products were resolved by PAGE to reveal a descending ladder of 32P-labeled DNAs according to the size of the DNA strand (Fig. 3). DNAs of 21 to 9 nucleotides were effective phosphoacceptor substrates for HD-Pnk, but a 6-mer oligonucleotide was not (Fig. 3). Parallel controls showed that the 6-mer DNA was readily phosphorylated by recombinant bacteriophage T4 Pnkp (Fig. 3), an enzyme for which the minimal phosphoacceptor is a 5′-OH nucleoside-3′-monophosphate (45).

FIG 3
Effect of oligonucleotide chain length on DNA kinase activity. HD-Pnk kinase reaction mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 5 mM MgCl2, 2.5 mM DTT, 100 μM [γ-32P]ATP, 100 pmol (10 μM) 5′-OH 21-mer, ...

HD-Pnk kinase use of any NTP as the phosphate donor.

Pnks can vary greatly in their NTP donor specificities. Whereas T4 Pnkp, Clostridium thermocellum Pnkp, and the kinase of the plant tRNA ligase Arabidopsis thaliana RNA ligase (AtRNL) use any NTP as a phosphate donor (13, 23), other Pnks are more fastidious. The Pnks of fungal tRNA ligases have a strong preference for GTP (24, 46, 47). In contrast, the human RNA kinase Clp1 favors ATP over GTP (48). To assess the NTP substrate preference of HD-Pnk, we employed an alternative kinase assay in which phosphate is transferred from an unlabeled NTP donor to the 5′-OH end of a 3′-32P-labeled 21-mer DNA substrate to form a 5′-phosphorylated product that migrates characteristically faster than the 5′-OH DNA substrate during denaturing PAGE (Fig. 4A). In the experiments shown in Fig. 4B, we reacted 0.25 μM HD-Pnk with 20 nM 3′-32P-labeled 21-mer DNA substrate for 5 min at 22°C, in the absence of added NTP or in the presence of 1, 10, 100, or 1,000 μM unlabeled ATP, GTP, CTP, UTP, or dATP. The reactions were quenched with EDTA, the products were analyzed by urea-PAGE, and the extents of 5′ phosphorylation were quantified. No phosphorylation was detected in the absence of added NTP. Varying the NTP concentrations from 1 to 10 to 100 μM elicited progressive and similar increases in the extents of 5′ phosphorylation for each NTP tested, indicating that HD-Pnk neither discriminates the nucleobase of the NTP donor nor displays a strong bias for a ribose versus deoxyribose sugar.

FIG 4
NTP donor requirements for the HD-Pnk kinase. (A) Kinase assay. A reaction mixture (60 μl) containing 100 mM Tris-acetate (pH 6.0), 5 mM MgCl2, 2.5 mM DTT, 100 μM ATP, 20 nM 3′-32P-labeled 21-mer HODNA (depicted at the bottom, ...

RNA CPDase activity of recombinant HD-Pnk.

Endonucleases that incise RNA via transesterification generate 2′,3′-cyclic phosphate and 5′-OH termini at the broken RNA ends. The fusion in Runella HD-Pnk of an HD phosphoesterase domain and a catalytically proficient 5′-OH polynucleotide kinase domain raised the prospect that the HD component might heal RNA>p ends. To test this idea, we prepared a 10-mer RNA substrate with 5′-OH and 2′,3′-cyclic phosphate ends and a single radiolabel between the 3′-terminal and penultimate nucleosides (Fig. 5A). A control experiment verified that reaction of the RNA>p substrate with T4 Pnkp, in the presence of magnesium and the absence of ATP (to limit activity to the acylphosphatase domain), resulted in quantitative conversion of RNA>p to a more slowly migrating RNAOH product (Fig. 5A, T4). The salient finding was that reaction with wild-type HD-Pnk in the presence of nickel also elicited the conversion of RNA>p to RNAOH (Fig. 5A, WT). Whereas the CPDase activity of HD-Pnk was apparently unaffected by the D254A and R257A mutations that eliminated the Pnk function, the H73A mutation in the HD motif strongly suppressed the formation of the fully dephosphorylated RNAOH product while allowing the conversion of a minority of the input RNA>p strands to a more rapidly migrating product that corresponded to a 10-mer RNAp with a terminal phosphomonoester (Fig. 5A).

FIG 5
RNA 2′,3′-cyclic phosphodiesterase activity of HD-Pnk. (A) CPDase assay. Reaction mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 5 mM NiCl2, 20 nM (0.2 pmol) 32P-labeled 10-mer HORNA>p (depicted at the bottom, ...

Because HD enzymes typically require transition metals as cofactors, we tested the metal dependence and metal specificity of HD-Pnk. Omission of exogenous metals from the reaction mixture plus inclusion of 10 mM EDTA was needed to eliminate CPDase activity. A low level of residual CPDase, leading to formation of RNAp and RNAOH species, was evident in the absence of added metals without EDTA supplementation, suggesting that some of the enzyme preparation retained metal cofactors in the HD CPDase active site. Among the divalent metals tested as CPDase cofactors at 5 mM, nickel, cobalt, and copper supported the greatest extent of conversion of RNA>p to RNAOH (Fig. 5B). In contrast, activity in the presence of calcium or zinc was hardly different than that in the absence of added metals (Fig. 5B). Manganese supported efficient substrate consumption but yielded a mixture of RNAp and RNAOH species. Magnesium elicited more modest stimulation of substrate consumption (versus the no-metal control), and the RNAp species predominated over RNAOH (Fig. 5B). Based on these findings, we assayed CPDase activity in the presence of nickel.

The extent of conversion of RNA>p to RNAOH during a 30-min reaction was proportional to the input wild-type HD-Pnk; there was scant RNAp present at any enzyme concentration (Fig. 5D). From the kinetic profile of the CPDase reaction of 20 nM RNA>p with 500 nM HD-Pnk, we saw that RNAp did accumulate transiently at early times, representing 21% and 23% of total labeled RNA at 1 and 3 min, respectively, when 35% and 54% of the input RNA>p had been converted to RNAOH (Fig. 5C). RNAp declined steadily thereafter (to 4% at 15 to 30 min), as RNAOH attained an endpoint value of 96% (Fig. 5C). Titration of the H73A mutant indicated that His73 subtraction had a greater impact on the phosphomonoesterase step than on the CPDase step, as inferred from the higher levels of RNAp versus RNAOH with 500 nM enzyme (Fig. 5D, 5 pmol).

The wild-type HD-Pnk CPDase was active over a broad pH range (from 4.5 to 9.5) in 100 mM Tris buffers. In reaction mixtures containing Tris-acetate (pH 4.5 to 6.5), the product distribution was similar to that seen in Fig. 5D, WT; the relative abundance of the RNAp species increased in reaction mixtures containing Tris-HCl (pH 8.0 to 9.5), at the expense of RNAOH (data not shown).

RNA 2′- and 3′-phosphomonoesterase activities of HD-Pnk.

We prepared 10-mer HORNA3′p and HORNA2′p substrates, with 3′-phosphate and 2′-phosphate termini, respectively, and a single 32P radiolabel between the 3′-terminal and penultimate nucleosides (Fig. 6A). HD-Pnk converted both substrates to RNAOH products, in an enzyme concentration-dependent fashion, in the presence of nickel (Fig. 6A). The RNA 3′-phosphatase specific activity was ~5-fold greater than the RNA 2′-phosphatase activity. The RNA 3′-phosphatase activity depended on a divalent cation (Fig. 6B), and values were similar with 0.5, 1, and 5 mM nickel (data not shown). Comparison of various metals at 1 mM showed that RNA 3′-phosphate hydrolysis was activated by nickel, cobalt, and copper but not by magnesium, manganese, or calcium (Fig. 6B). Zinc was a feeble cofactor for the RNA 3′ phosphatase (Fig. 6B). Single-turnover RNA 3′-phosphate hydrolysis under conditions of enzyme excess (kobs of 2.1 ± 0.1 min−1) was 5-fold faster than RNA 2′-phosphate hydrolysis (kobs of 0.42 ± 0.03 min−1) (Fig. 6C).

FIG 6
RNA 2′- and 3′-phosphomonoesterase activities of HD-Pnk. (A) Enzyme titration. Reaction mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 5 mM NiCl2, 20 nM (0.2 pmol) 32P-labeled 10-mer RNAs with 2′-phosphate or ...

DNA 3′-phosphatase activity of HD-Pnk.

DNA 3′-end-healing activity was assessed by reaction of HD-Pnk with a 32P-labeled 10-mer pDNAp substrate of the same nucleobase sequence as the RNAp substrates used in the preceding sections. HD-Pnk converted pDNAp to a more slowly migrating pDNAOH product in the presence of a suitable metal cofactor (Fig. 7A). DNA 3′-phosphate hydrolysis was activated best by 1 mM nickel or copper, less well by cobalt, feebly by zinc, and not at all by magnesium, manganese, or calcium (Fig. 7A). The DNA 3′-phosphatase activity was unaffected by the kinase-inactivating D254A and R257A mutations but was strongly suppressed by the H73A mutation (Fig. 7B). Copper was preferred over nickel as the cofactor for the DNA 3′-phosphatase reaction, as determined by enzyme titration (Fig. 7C) and reaction kinetics (Fig. 7D).

FIG 7
DNA 3′-phosphatase activity of HD-Pnk. (A) Divalent cation requirements. Reaction mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 0.1 μM (1 pmol) 5′-32P-labeled 10-mer pDNAp (depicted at the bottom, with the labeled ...

DISCUSSION

The present study provides insights regarding the diversity of nucleic acid repair strategies via the characterization of HD-Pnk as the exemplar of a novel clade of dual 5′- and 3′-end-healing enzymes. The distinctive feature of HD-Pnk is its domain composition, i.e., a fusion of a canonical P-loop polynucleotide kinase module and an HD hydrolase module. The biochemical properties of the 5′ kinase are similar to those of other well-studied bacterial or viral Pnks. The reliance of the kinase activity of HD-Pnk on the DXXR motif suggests that the kinase adheres to the mechanism of general base catalysis by aspartate and 5′-nucleotide recognition by arginine invoked for other Pnks.

The biochemical analysis of the HD-catalyzed 3′-end-healing reactions of Runella HD-Pnk extends the catalytic repertoire of the HD family. We show that HD-Pnk can dephosphorylate RNA 2′,3′-cyclic phosphate, RNA 3′-phosphate, RNA 2′-phosphate, and DNA 3′-phosphate ends in the presence of a transition metal cofactor, which can be nickel, copper, or cobalt. Whereas HD-Pnk has CPDase and phosphomonoesterase activities at nucleic acid 3′ ends, it is neither a nuclease (3′-5′ phosphodiesterase) nor a polynucleotide 5′ phosphatase, as indicated by the stability of the 3′-32P-labeled and 5′-32P-labeled products of the HD-Pnk 3′-end-healing reactions.

Ours is not the first study in which a Pnk domain was noted to be fused to an HD-like module within a putative nucleic acid repair enzyme. Blondal et al. (49) identified a Pnk-like protein encoded by thermophilic bacteriophage RM378, and they characterized its 5′-kinase activity with RNA and DNA substrates. An N-terminal extension with an HD motif flanking the kinase module prompted them to test for phosphoesterase activity. They demonstrated a capacity of the RM378 enzyme to release phosphate from a cyclic mononucleotide substrate (2′,3′-cAMP) in the presence of manganese, but they were unable to detect DNA 3′-phosphatase activity (49). In the same vein, Blasius et al. (50) purified the Deinococcus radiodurans DRB0098 protein, which is composed of a C-terminal Pnk-like domain and an N-terminal HD-like domain. Whereas those authors demonstrated that recombinant DRB0098 had 5′-OH DNA kinase activity in the presence of manganese, they were unable to assign DNA 3′-phosphatase activity to the recombinant protein. Thus, our study of Runella HD-Pnk is the first study to show that the HD module associated with a Pnk has an actual nucleic acid end-healing activity. It is conceivable that the failure of previous investigators to assign a DNA/RNA 3′-end-healing function to phage and bacterial homologs of Runella HD-Pnk stems from the use of an ineffective metal cofactor for the HD hydrolase domain.

Homologs of Runella HD-Pnk with the same domain composition, same domain order, and similar polypeptide size are distributed widely in the bacterial domain of life, spanning genera from at least 11 different phyla (see Table S1 in the supplemental material). Runella belongs to the phylum Bacteroidetes, of which at least 21 other genera have HD-Pnk homologs (Table S1). The phyla Proteobacteria, Actinobacteria, and Cyanobacteria are also rich in genera that encode HD-Pnk (Table S1). As noted here for Runella and E. coli HD-Pnks, other bacterial HD-Pnk proteins are frequently genetically clustered with a ligase-like adenylyltransferase enzyme, suggesting that they do indeed play an end-healing role in bacterial nucleic acid repair. On the basis of our inspection, HD-Pnk homologs are not prevalent in archaea or eukarya.

MATERIALS AND METHODS

HD-Pnk purification and mutagenesis.

The HD-Pnk ORF was amplified by PCR from Runella slithyformis (ATCC 49304) genomic DNA (44) with primers that introduced a BamHI site at the translation start codon and an XhoI site flanking the stop codon. The PCR product was digested with BamHI and XhoI and then inserted into the T7 RNA polymerase-based expression vector pET28b-His10Smt3 to fuse HD-Pnk to an N-terminal His10-Smt3 tag. The alanine mutations H73A, D254A, and R257A were introduced into the expression vector by quick-change PCR. The inserts of all plasmids were sequenced to confirm that no unwanted coding changes were introduced during PCR. The resulting pET28b-His10Smt3-HD-Pnk and pET28b-His10Smt3-HD-Pnk-Ala plasmids were transformed into Escherichia coli BL21+(DE3). Cultures (2 liters) derived from single kanamycin- and chloramphenicol-resistant transformants were grown at 37°C in Luria-Bertani medium containing 50 μg/ml kanamycin and 35 μg/ml chloramphenicol until the A600 reached 0.5 to 0.6, at which time the cultures were adjusted to contain 0.4 mM isopropyl-β-d-thiogalactoside and 2% (vol/vol) ethanol and then were incubated for 16 h at 17°C, with continuous shaking. Cells were harvested by centrifugation, and the pellets were stored at −80°C. All subsequent procedures were performed at 4°C. Thawed bacteria were suspended in 50 ml buffer A (50 mM Tris-HCl [pH 7.5], 1 M NaCl, 20 mM imidazole, 0.05% Triton X-100, 10% glycerol), and lysozyme was added to a final concentration of 1 mg/ml, along with one protease inhibitor tablet (Roche). After being mixed for 50 min, the resulting lysates were sonicated to reduce viscosity, and insoluble material was removed by centrifugation at 14,000 rpm for 45 min. The soluble extracts were mixed for 1 h with 5 ml of His60 Ni Superflow resin (Qiagen) that had been equilibrated in buffer A. The resin was washed twice with 50 ml of buffer A and then serially with 30 ml of buffer B (50 mM Tris-HCl [pH 7.5], 3 M KCl) and 50 ml of 40 mM imidazole in buffer C (50 mM Tris-HCl [pH 7.5], 500 mM NaCl, 10% glycerol). The resin was then poured into a column, and the bound material was serially step eluted with 100 mM, 300 mM, and 500 mM imidazole in buffer C. The elution profiles were monitored by SDS-PAGE. The peak His10-Smt3-HD-Pnk-containing fractions were pooled and supplemented with Smt3-specific protease Ulp1 (100 μg) to cleave the tag during overnight dialysis against 4 liters of buffer D (50 mM Tris-HCl [pH 7.5], 500 mM NaCl, 10% glycerol). SDS-PAGE analysis of the dialysates indicated that the His10-Smt3 tag had been cleaved from the protein. The dialysates were applied to 2.5 ml of Ni Superflow resin, and the tag-free HD-Pnk proteins were recovered in the flowthrough and wash fractions in buffer C, which were adjusted to contain 25 mM EDTA. The cleaved His10-Smt3 tag was bound to the resin and recovered in the 500 mM imidazole eluate. The HD-Pnk protein preparations were concentrated by centrifugal ultrafiltration (to ~5 to 7 mg/ml in 5 ml) and then gel filtered, at a flow rate of 1 ml/min, through a 120-ml Superdex 200 column equilibrated with buffer E (50 mM Tris-HCl [pH 7.5], 500 mM NaCl, 1 mM dithiothreitol [DTT], 1 mM EDTA, 10% glycerol), with collection of 2-ml fractions. Peak HD-Pnk fractions were concentrated by centrifugal ultrafiltration and stored at −80°C. Protein concentrations were determined by using the Bio-Rad dye reagent, with bovine serum albumin as the standard. The yields of HD-Pnk proteins were as follows: WT, 18 mg; H73A mutant, 8 mg; D254A mutant, 20 mg; and R257A mutant, 25 mg.

Polynucleotide kinase assay.

Reaction mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 5 mM MgCl2, 2.5 mM dithiothreitol, 100 μM [γ-32P]ATP, 100 pmol (10 μM) 21-mer 5′-OH DNA oligonucleotide d(CTAGAGCTACAATTGCGACCG), and HD-Pnk as specified were incubated at 37°C. The reactions were initiated by adding HD-Pnk and were quenched by adding an equal volume of 90% formamide, 50 mM EDTA, 0.01% bromophenol blue-xylene cyanol. The mixtures were analyzed by electrophoresis (at a constant power of 15 W) through a 15-cm 20% polyacrylamide gel containing 7 M urea in 45 mM Tris-borate, 1 mM EDTA. The radiolabeled 21-mer oligonucleotide products were quantified by scanning the gel with a Fujifilm FLA-7000 imaging device.

Alternatively, kinase activity was assayed using a 3′-32P-labeled 21-mer HODNA oligonucleotide substrate, i.e., HOCTAGAGCTACAATTGCGACCpA (prepared by template-based addition of [32P]dAMP to a 20-mer DNA primer oligonucleotide using Klenow DNA polymerase I, followed by gel purification of the radiolabeled 21-mer strand). Kinase reaction mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 5 mM MgCl2, 2.5 mM DTT, 0.2 pmol (20 nM) 3′-32P-labeled 21-mer HODNA, NTP as specified, and 2.5 pmol (250 nM) HD-Pnk were incubated at 22°C. The reactions were initiated by adding HD-Pnk and were quenched by adding an equal volume of 90% formamide, 50 mM EDTA, 0.01% bromophenol blue-xylene cyanol. The mixtures were analyzed by electrophoresis (at a constant power of 55 W) through a 40-cm 20% polyacrylamide gel containing 7 M urea in 45 mM Tris-borate, 1 mM EDTA. The 32P-labeled DNAs were visualized by autoradiography and quantified by scanning the gel with a Fujifilm FLA-7000 imaging device.

Preparation of 3′-32P-labeled RNA substrates.

A 10-mer HORNA3′p oligonucleotide labeled with 32P at the penultimate phosphate was prepared by T4 Rnl1-mediated addition of [5′-32P]pCp to a 9-mer synthetic oligoribonucleotide, r(AUCACGCUU), as described previously (25). The 32P-labeled strand AUCACGCUUpCp was then reacted with E. coli RNA 3′-terminal phosphate cyclase (RtcA) and ATP to generate a 2′,3′-cyclic phosphate derivative, HORNA>p (25). The HORNA3′p and HORNA>p strands were gel purified, eluted from an excised gel slice, and recovered by ethanol precipitation. A 2′-PO4-terminated derivative, HORNA2′p, was prepared by reacting HORNA>p with the cyclic phosphodiesterase of AtRNL, as described previously (25). The HORNA2′p strand was gel purified.

Assay of RNA 3′ end healing.

Reaction mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 1 mM or 5 mM NiCl2, 20 nM (0.2 pmol) 10-mer HORNA>p, HORNA3′p, or HORNA2′p as a substrate, and HD-Pnk as specified were incubated at 37°C. The reactions were initiated by adding HD-Pnk and were quenched by adding an equal volume of 90% formamide, 50 mM EDTA, 0.01% bromophenol blue-xylene cyanol. The mixtures were analyzed by electrophoresis (at a constant power of 55 W) through a 40-cm 20% polyacrylamide gel containing 7 M urea in 45 mM Tris-borate, 1 mM EDTA. The products were visualized by autoradiography and quantified by scanning the gel with a Fujifilm FLA-7000 imager.

Assay of DNA 3′ phosphatase activity.

Reactions mixtures (10 μl) containing 100 mM Tris-acetate (pH 6.0), 1 mM CuCl2 or NiCl2, 100 nM (1 pmol) 10-mer 5′-32P-labeled pDNAp substrate (5′-pATCACGCTTCp; prepared by enzymatic phosphorylation of a synthetic 10-mer HODNAp oligonucleotide by phosphatase-inactive T4 Pnkp-D167N and then gel purification), and HD-Pnk as specified were incubated at 37°C. The reactions were initiated by adding HD-Pnk and were quenched by adding an equal volume of formamide-EDTA. The products were analyzed by electrophoresis (at a constant power of 55 W) through a 40-cm 20% polyacrylamide gel containing 7 M urea in 45 mM Tris-borate, 1 mM EDTA, visualized by autoradiography, and quantified by scanning the gel with a Fujifilm FLA-7000 imager.

Supplementary Material

Supplemental material:

ACKNOWLEDGMENT

This research was supported by NIH grant GM46330 (to S.S.).

Footnotes

Supplemental material for this article may be found at https://doi.org/10.1128/JB.00739-16.

REFERENCES

1. Wang LK, Shuman S 2001. Domain structure and mutational analysis of T4 polynucleotide kinase. J Biol Chem 276:26868–26874. doi:.10.1074/jbc.M103663200 [PubMed] [Cross Ref]
2. Wang LK, Shuman S 2002. Mutational analysis defines the 5′ kinase and 3′ phosphatase active sites of T4 polynucleotide kinase. Nucleic Acids Res 30:1073–1080. doi:.10.1093/nar/30.4.1073 [PMC free article] [PubMed] [Cross Ref]
3. Wang LK, Lima CD, Shuman S 2002. Structure and mechanism of T4 polynucleotide kinase: an RNA repair enzyme. EMBO J 21:3873–3880. doi:.10.1093/emboj/cdf397 [PubMed] [Cross Ref]
4. Galburt EA, Pelletier J, Wilson G, Stoddard BL 2002. Structure of a tRNA repair enzyme and molecular biology workhorse: T4 polynucleotide kinase. Structure 10:1249–1260. doi:.10.1016/S0969-2126(02)00835-3 [PubMed] [Cross Ref]
5. Das U, Shuman S 2013. Mechanism of RNA 2′,3′-cyclic phosphate end-healing by T4 polynucleotide kinase-phosphatase. Nucleic Acids Res 41:355–365. doi:.10.1093/nar/gks977 [PMC free article] [PubMed] [Cross Ref]
6. Garces F, Pearl LH, Oliver AW 2011. The structural basis for substrate recognition by mammalian polynucleotide kinase 3′ phosphatase. Mol Cell 44:385–396. doi:.10.1016/j.molcel.2011.08.036 [PMC free article] [PubMed] [Cross Ref]
7. Coquelle N, Havali-Shahriari Z, Bernstein N, Green R, Glover JNM 2011. Structural basis for the phosphatase activity of PNKP on single- and double-stranded DNA substrates. Proc Natl Acad Sci U S A 108:21022–21027. doi:.10.1073/pnas.1112036108 [PubMed] [Cross Ref]
8. Martins A, Shuman S 2004. Characterization of a baculovirus enzyme with RNA ligase, polynucleotide 5′ kinase and polynucleotide 3′ phosphatase activities. J Biol Chem 279:18220–18231. doi:.10.1074/jbc.M313386200 [PubMed] [Cross Ref]
9. Wang P, Selvadurai K, Huang RH 2015. Reconstitution and structure of a bacterial Pnkp1-Rnl-Hen1 RNA repair complex. Nat Commun 6:6876. doi:.10.1038/ncomms7876 [PMC free article] [PubMed] [Cross Ref]
10. Martins A, Shuman S 2005. An end-healing enzyme from Clostridium thermocellum with 5′ kinase, 2′,3′ phosphatase, and adenylyltransferase activities. RNA 11:1271–1280. doi:.10.1261/rna.2690505 [PubMed] [Cross Ref]
11. Chan CM, Zhou C, Huang R 2009. Reconstituting bacterial RNA repair and modification in vitro. Science 326:247. doi:.10.1126/science.1179480 [PubMed] [Cross Ref]
12. Wang LK, Das U, Smith P, Shuman S 2012. Structure and mechanism of the polynucleotide kinase component of the bacterial Pnkp-Hen1 RNA repair system. RNA 18:2277–2286. doi:.10.1261/rna.036061.112 [PubMed] [Cross Ref]
13. Das U, Wang LK, Smith P, Shuman S 2013. Structural and biochemical analysis of the phosphate donor specificity of the polynucleotide kinase component of the bacterial Pnkp·Hen1 RNA repair system. Biochemistry 52:4734–4743. doi:.10.1021/bi400412x [PMC free article] [PubMed] [Cross Ref]
14. Das U, Wang LK, Smith P, Jacewicz A, Shuman S 2014. Structures of bacterial polynucleotide kinase in a Michaelis complex with GTP·Mg2+ and 5′-OH oligonucleotide and a product complex with GDP·Mg2+ and 5′-PO4 oligonucleotide reveal a mechanism of general acid-base catalysis and the determinants of phosphoacceptor recognition. Nucleic Acids Res 42:1152–1161. doi:.10.1093/nar/gkt936 [PMC free article] [PubMed] [Cross Ref]
15. Das U, Wang LK, Smith P, Munir A, Shuman S 2014. Structures of bacterial polynucleotide kinase in a Michaelis complex with nucleoside triphosphate (NTP)-Mg2+ and 5′-OH RNA and a mixed substrate-product complex with NTP-Mg2+ and a 5′-phosphorylated oligonucleotide. J Bacteriol 196:4285–4292. doi:.10.1128/JB.02197-14 [PMC free article] [PubMed] [Cross Ref]
16. Keppetipola N, Shuman S 2006. Mechanism of the phosphatase component of Clostridium thermocellum polynucleotide kinase-phosphatase. RNA 12:73–82. doi:.10.1261/rna.2196406 [PubMed] [Cross Ref]
17. Keppetipola N, Shuman S 2006. Distinct enzymic functional groups are required for the phosphomonoesterase and phosphodiesterase activities of Clostridium thermocellum polynucleotide kinase/phosphatase. J Biol Chem 281:19251–19259. doi:.10.1074/jbc.M602549200 [PubMed] [Cross Ref]
18. Wang LK, Smith P, Shuman S 2013. Structure and mechanism of the 2′,3′ phosphatase component of the bacterial Pnkp-Hen1 RNA repair system. Nucleic Acids Res 41:5864–5873. doi:.10.1093/nar/gkt221 [PMC free article] [PubMed] [Cross Ref]
19. Smith P, Wang LK, Nair PA, Shuman S 2012. The adenylyltransferase domain of bacterial Pnkp defines a unique RNA ligase family. Proc Natl Acad Sci U S A 109:2296–2301. doi:.10.1073/pnas.1116827109 [PubMed] [Cross Ref]
20. Wang P, Chan CM, Christensen D, Zhang C, Selvadurai K, Huang RH 2012. Molecular basis of bacterial protein Hen1 activating the ligase activity of bacterial protein Pnkp for RNA repair. Proc Natl Acad Sci U S A 109:13248–13253. doi:.10.1073/pnas.1209805109 [PubMed] [Cross Ref]
21. Sawaya R, Schwer B, Shuman S 2003. Genetic and biochemical analysis of the functional domains of yeast tRNA ligase. J Biol Chem 278:43928–43938. doi:.10.1074/jbc.M307839200 [PubMed] [Cross Ref]
22. Wang LK, Schwer B, Englert M, Beier H, Shuman S 2006. Structure-function analysis of the kinase-CPD domain of yeast tRNA ligase (Trl1) and requirements for complementation of tRNA splicing by a plant Trl1 homolog. Nucleic Acids Res 34:517–527. doi:.10.1093/nar/gkj441 [PMC free article] [PubMed] [Cross Ref]
23. Remus BS, Shuman S 2013. A kinetic framework for tRNA ligase and enforcement of a 2′-phosphate requirement for ligation highlights the design logic of an RNA repair machine. RNA 19:659–669. doi:.10.1261/rna.038406.113 [PubMed] [Cross Ref]
24. Remus BS, Shuman S 2014. Distinctive kinetics and substrate specificities of plant and fungal tRNA ligases. RNA 20:462–473. doi:.10.1261/rna.043752.113 [PubMed] [Cross Ref]
25. Tanaka N, Chakravarty AK, Maughan B, Shuman S 2011. A novel mechanism of RNA repair by RtcB via sequential 2′,3′-cyclic phosphodiesterase and 3′-phosphate/5′-hydroxyl ligation reactions. J Biol Chem 286:43134–43143. doi:.10.1074/jbc.M111.302133 [PMC free article] [PubMed] [Cross Ref]
26. Chakravarty AK, Subbotin R, Chait BT, Shuman S 2012. RNA ligase RtcB splices 3′-phosphate and 5′-OH ends via covalent RtcB-(histidinyl)-GMP and polynucleotide-(3′)pp(5′)G intermediates. Proc Natl Acad Sci U S A 109:6072–6077. doi:.10.1073/pnas.1201207109 [PubMed] [Cross Ref]
27. Das U, Chakravarty AK, Remus BS, Shuman S 2013. Rewriting the rules for end joining via enzymatic splicing of DNA 3′-PO4 and 5′-OH ends. Proc Natl Acad Sci U S A 110:20437–20442. doi:.10.1073/pnas.1314289110 [PubMed] [Cross Ref]
28. Aravind L, Koonin EV 1998. The HD domain defines a new superfamily of metal-dependent phosphohydrolases. Trends Biochem Sci 23:469–472. doi:.10.1016/S0968-0004(98)01293-6 [PubMed] [Cross Ref]
29. Yakunin AF, Proudfoot M, Kuznetsova E, Savchenko A, Brown G, Arrowsmith CH, Edwards AM 2004. The HD domain of the Escherichia coli tRNA nucleotidyltransferase has 2′,3′-cyclic phosphodiesterase, 2′-nucleotidase, and phosphatase activities. J Biol Chem 279:36819–36827. doi:.10.1074/jbc.M405120200 [PubMed] [Cross Ref]
30. Nagata M, Kaito C, Sekimizu K 2008. Phosphodiesterase activity of CvfA is required for virulence in Staphylococcus aureus. J Biol Chem 283:2176–2184. doi:.10.1074/jbc.M705309200 [PubMed] [Cross Ref]
31. Zimmerman MD, Proudfoot M, Yakunin A, Minor W 2008. Structural insight into the mechanism of substrate specificity and catalytic activity of an HD-domain phosphohydrolase: the 5′-deoxyribonucleotidase YfbR from Escherichia coli. J Mol Biol 378:215–226. doi:.10.1016/j.jmb.2008.02.036 [PMC free article] [PubMed] [Cross Ref]
32. Jeon YJ, Park SC, Song WS, Kim OH, Oh BC, Yoon SI 2016. Structural and biochemical characterization of bacterial YpgQ protein reveals a metal-dependent nucleotide pyrophosphohydrolase. J Struct Biol 195:113–122. doi:.10.1016/j.jsb.2016.04.002 [PubMed] [Cross Ref]
33. Beloglazova N, Petit P, Flick R, Brown G, Savchenko A, Yakunin AF 2011. Structure and activity of the Cas3 HD nuclease MJ0384, an effector enzyme of the CRISPR interference. EMBO J 30:4616–4627. doi:.10.1038/emboj.2011.377 [PubMed] [Cross Ref]
34. Mulepati S, Bailey S 2011. Structural and biochemical analysis of nuclease domain of clustered regularly interspaced short palindromic repeat (CRISPR)-associated protein 3 (Cas3). J Biol Chem 286:31896–31903. doi:.10.1074/jbc.M111.270017 [PMC free article] [PubMed] [Cross Ref]
35. Huo Y, Nam KH, Ding F, Lee H, Wu L, Xiao Y, Farchione MD, Zhou S, Rajashankar K, Kurinov I, Zhang R, Ke A 2014. Structures of CRISPR Cas3 offer mechanistic insights into Cascade-activated DNA unwinding and degradation. Nat Struct Mol Biol 21:771–777. doi:.10.1038/nsmb.2875 [PMC free article] [PubMed] [Cross Ref]
36. Benda C, Ebert J, Scheltema RA, Schiller HB, Baumgärtner M, Bonneau F, Mann M, Conti E 2014. Structural model of a CRISPR RNA-silencing complex reveals the RNA-target cleavage activity in Cmr4. Mol Cell 56:43–54. doi:.10.1016/j.molcel.2014.09.002 [PubMed] [Cross Ref]
37. Jung TY, An Y, Park KH, Lee MH, Oh BH, Woo E 2015. Crystal structure of the Csm1 subunit of the Csm complex and its single-stranded DNA-specific nuclease activity. Structure 23:782–790. doi:.10.1016/j.str.2015.01.021 [PubMed] [Cross Ref]
38. Bellini D, Caly DL, McCarthy Y, Bumann M, An SQ, Dow JM, Ryan RP, Walsh MA 2014. Crystal structure of an HD-GYP domain cyclic-di-GMP phosphodiesterase reveals an enzyme with a novel trinuclear catalytic iron centre. Mol Microbiol 91:26–38. doi:.10.1111/mmi.12447 [PMC free article] [PubMed] [Cross Ref]
39. Huynh TN, Luo S, Pensinger D, Sauer JD, Tong L, Woodward JJ 2015. An HD-domain phosphodiesterase mediates cooperative hydrolysis of c-di-AMP to affect bacterial growth and virulence. Proc Natl Acad Sci U S A 112:E747–E756. doi:.10.1073/pnas.1416485112 [PubMed] [Cross Ref]
40. Mashhadi Z, Xu H, White RH 2009. An Fe2+-dependent cyclic phosphodiesterase catalyzes the hydrolysis of 7,8-dihydro-d-neopterin 2′,3′-cyclic phosphate in methanopterin biosynthesis. Biochemistry 48:9384–9392. doi:.10.1021/bi9010336 [PubMed] [Cross Ref]
41. Wörsdörfer B, Lingaraju M, Yennawar NH, Boal AK, Krebs C, Bollinger JM, Pandelia ME 2013. Organophosphonate-degrading PhnZ reveals an emerging family of HD domain mixed-valent diiron oxygenases. Proc Natl Acad Sci U S A 110:18874–18879. doi:.10.1073/pnas.1315927110 [PubMed] [Cross Ref]
42. Culver GM, McCraith SM, Consaul SA, Stanford DR, Phizicky EM 1997. A 2′-phosphotransferase implicated in tRNA splicing is essential in Saccharomyces cerevisiae. J Biol Chem 272:13203–13210. doi:.10.1074/jbc.272.20.13203 [PubMed] [Cross Ref]
43. Tumbale P, Appel CD, Kraehenbuehl R, Robertson PD, Williams JS, Krahn J, Ahel I, Williams RS 2011. Structure of an aprataxin-DNA complex with insights into AOA1 neurodegenerative disease. Nat Struct Mol Biol 18:1189–1195. doi:.10.1038/nsmb.2146 [PMC free article] [PubMed] [Cross Ref]
44. Copeland A, Zhang X, Misra M, Lapidus A, Nolan M, Lucas S, Deshpande S, Cheng JF, Tapia R, Goodwin LA, Pitluck S, Liolios K, Pagani I, Ivanova N, Mikhailova N, Pati A, Chen A, Palaniappan K, Land M, Hauser L, Pan C, Jeffries CD, Detter JC, Brambilla EM, Rohde M, Djao OD, Göker M, Sikorski J, Tindall BJ, Woyke T, Bristow J, Eisen JA, Markowitz V, Hugenholtz P, Kyrpides NC, Klenk HP, Mavromatis K 2012. Complete genome sequence of the aquatic bacterium Runella slithyformis type strain (LSU 4T). Stand Genomic Sci 6:145–154. doi:.10.4056/sigs.2475579 [PMC free article] [PubMed] [Cross Ref]
45. Wang LK, Shuman S 2010. Mutational analysis of the 5′-OH oligonucleotide phosphate acceptor site of T4 polynucleotide kinase. Nucleic Acids Res 38:1304–1311. doi:.10.1093/nar/gkp1096 [PMC free article] [PubMed] [Cross Ref]
46. Remus BS, Schwer B, Shuman S 2016. Characterization of the tRNA ligases of pathogenic fungi Aspergillus fumigatus and Coccidioides immitis. RNA 22:1500–1509. doi:.10.1261/rna.057455.116 [PubMed] [Cross Ref]
47. Westaway SK, Belford HG, Apostol BL, Abelson J, Greer CL 1993. Novel activity of a yeast ligase deletion polypeptide: evidence for GTP-dependent tRNA splicing. J Biol Chem 268:2435–2443. [PubMed]
48. Weitzer S, Martinez J 2007. The human RNA kinase hClp1 is active on 3′ transfer RNA exons and short interfering RNAs. Nature 447:222–226. doi:.10.1038/nature05777 [PubMed] [Cross Ref]
49. Blondal T, Hjorleifsdottir S, Aevarsson A, Fridjonsson OH, Skirnisdottir S, Wheat JO, Hermannsdottir AG, Hreggvidsson GO, Smith AV, Kristjansson JK 2005. Characterization of a 5′-polynucleotide kinase/3′-phosphatase from bacteriophage RM378. J Biol Chem 280:5188–5194. doi:.10.1074/jbc.M409211200 [PubMed] [Cross Ref]
50. Blasius M, Buob R, Shevelev IV, Hubscher U 2007. Enzymes involved in DNA ligation and end-healing in the radioresistant bacterium Deinococcus radiodurans. BMC Mol Biol 8:69. doi:.10.1186/1471-2199-8-69 [PMC free article] [PubMed] [Cross Ref]

Articles from Journal of Bacteriology are provided here courtesy of American Society for Microbiology (ASM)