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J. Biol. Chem., Vol. 277, Issue 24, 21624-21629, June 14, 2002

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Purification and Characterization of the Escherichia coli Exoribonuclease RNase R

COMPARISON WITH RNase II^*

Zhuan-Fen Cheng and Murray P. Deutscher

From the Department of Biochemistry and Molecular Biology, University of Miami School of Medicine, Miami, Florida 33101

Received for publication, March 27, 2002

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Escherichia coli RNase R, a 3'  5' exoribonuclease homologous to RNase II, was overexpressed and purified to near homogeneity in its native untagged form by a rapid procedure. The purified enzyme was free of nucleic acid. It migrated upon gel filtration chromatography as a monomer with an apparent molecular mass of ~95 kDa, in close agreement with its expected size based on the sequence of the rnr gene. RNase R was most active at pH 7.5-9.5 in the presence of 0.1-0.5 mM Mg²⁺ and 50-500 mM KCl. The enzyme shares many catalytic properties with RNase II. Both enzymes are nonspecific processive ribonucleases that release 5'-nucleotide monophosphates and leave a short undigested oligonucleotide core. However, whereas RNase R shortens RNA processively to di- and trinucleotides, RNase II becomes more distributive when the length of the substrate reaches ~10 nucleotides, and it leaves an undigested core of 3-5 nucleotides. Both enzymes work on substrates with a 3'-phosphate group. RNase R and RNase II are most active on synthetic homopolymers such as poly(A), but their substrate specificities differ. RNase II is more active on poly(A), whereas RNase R is much more active on rRNAs. Neither RNase R nor RNase II can degrade a complete RNA-RNA or DNA-RNA hybrid or one with a 4-nucleotide 3'-RNA overhang. RNase R differs from RNase II in that it cannot digest DNA oligomers and is not inhibited by such molecules, suggesting that it does not bind DNA. Although the in vivo function of RNase R is not known, its ability to digest certain natural RNAs may explain why it is maintained in E. coli together with RNase II.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Escherichia coli contains eight distinct exoribonucleases that play important roles in every aspect of RNA metabolism (1, 2). One of these enzymes, now termed RNase R, was originally identified and partially purified and characterized as a nonspecific residual exoribonuclease present in strains lacking RNase II (3, 4). It was subsequently rediscovered in our laboratory as a nuclease active against rRNA and given the name RNase R (5, 6). The enzyme accounts for ~2% of poly(A)-degrading activity remaining in crude extracts of cells lacking RNase I and RNase II (7). Partially purified RNase R is a 3' 5' exoribonuclease that releases 5'-nucleoside monophosphates, and catalytically, it resembles RNase II (3, 4).

RNase R is encoded by the rnr gene located at 95 min on the E. coli chromosome (7). This gene was originally termed vacB; and in Shigella and in enteroinvasive E. coli, it is necessary for expression of virulence (8). In laboratory strains of E. coli, RNase R is dispensable for cell viability (7). Moreover, multiple mutant strains lacking RNases R and T, RNases R and PH, and RNases R, II, D, and BN grow essentially normally on rich media (7). On the other hand, a double mutant strain devoid of RNase R and polynucleotide phosphorylase is inviable (7). This observation suggests that RNase R and polynucleotide phosphorylase serve some overlapping essential function(s) in E. coli that cannot be satisfied by any of the other cellular exoribonucleases. Preliminary data indicate that at least one of these functions is an RNA quality control process that eliminates defective rRNA.¹

RNase R, together with RNase II, is a member of the RNR superfamily of exoribonucleases (1). As might be expected from their similarity in catalytic properties, RNase R and RNase II also share structural properties, including ~60% sequence homology. Interestingly, RNase R is even more widespread among eubacteria than is RNase II (1). In fact, RNase R is the only one of the eight known bacterial exoribonucleases to be present in Mycoplasma (1).

As part of our continuing efforts to determine the physiological role of RNase R, we have developed a simple procedure to purify the enzyme to near homogeneity. In this study, we describe the purification of RNase R and provide a detailed characterization of its structure, catalytic properties, and substrate specificity. In view of its high degree of similarity to RNase II, for many experiments, we have directly compared the properties of RNase R with those of a homogeneous preparation of RNase II (9).

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Materials-- Restriction endonucleases, T4 DNA ligase, T4 polynucleotide kinase, and the Klenow fragment of DNA polymerase I were obtained from New England Biolabs Inc. Calf intestine alkaline phosphatase was from Promega. The QIAEX II gel extraction kit was purchased from QIAGEN Inc. [³²P]Orthophosphate and [-³²P]ATP (6,000 Ci/mmol) were obtained from PerkinElmer Life Sciences. Polyvinylidene difluoride membranes were a product of Pall Corp. [³H]Poly(A) and Ultrogel AcA44 were obtained from Amersham Biosciences. Poly(A), DNase I, and bacterial alkaline phosphatase (from E. coli) and its substrate p-nitrophenyl phosphate were from Sigma. Affi-Gel blue-agarose (100-200 mesh) and protein size markers were obtained from Bio-Rad. SequaGel, used to make urea-polyacrylamide gels, was purchased from National Diagnostics, Inc. All other chemicals were reagent-grade. RNA oligomers were synthesized by Dharmacon Research Inc., and DNA oligomers were synthesized by the DNA Core Facility of our department. The RNA oligomers used for substrate specificity studies were 5'-CCCCACCACCAUCACUU-3' (17-mer), its 17-mer complement, and the homo-oligomers A₆, A₈, A₁₀, and A₁₇. The DNA oligomers used were 5'-GATGGTGGTGGGG-3' (13-mer) and 5'-AAGTGATGGTGGTGGGG-3' (17-mer).

Bacterial Strains and Plasmids-- The E. coli K12 strain CMA201 (rnb201::tet) was a kind gift from Dr. C. Arriano (Centro de Tecnologia Química e Biológica, University of Lisbon, Lisbon, Portugal) (10). It was used to prepare the P1vir lysate for conversion of strain BL21(DE3)/pLys (Novagen) to an RNase II strain by P1-mediated transduction. The resulting strain, BL21II(DE3)/pLys, was the host for plasmid pETR-dependent overexpression of RNase R. The RNase II strain was employed to avoid problems in the assay of RNase R during purification due to the greater activity of RNase II on poly(A). Strain CA244I was used to prepare ribosomes and rRNAs.

Plasmid pETR was generated by insertion of a NaeI-BclI fragment containing the rnr gene and a portion of the upstream yjeB gene at the NcoI (blunted)-BamHI site of plasmid pET15b (Novagen) (Fig. 1). During the cloning procedure, the His tag coding region of pET15b was removed, and the recombinant RNase R protein was not tagged.

View larger version (12K): [in this window] [in a new window]   Fig. 1.   Cloning of the E. coli gene rnr for overexpression. A shows the sequence immediately upstream of rnr. The residues shown in boldface are the two possible initiation codons for rnr; underlined is the stop codon of yjeB; and the italicized sequence is a possible Shine-Dalgarno sequence assuming that the downstream ATG is the actual initiation codon. B shows a linear representation of the rnr gene of E. coli and its flanking genes. C shows plasmid pETR used to overexpress RNase R. Plasmid pETR was constructed as described under "Experimental Procedures." It contains the rnr gene and a portion of the upstream gene yjeB cloned immediately after the lac operator and the Shine-Dalgarno sequence of the vector.

Overexpression of RNase R-- Strain BL21II(DE3)/pLys harboring pETR was grown to A₅₅₀  1 with vigorous shaking at 37 °C in 4 liters of yeast-tryptone medium supplemented with 100 µg/ml ampicillin plus 34 µg/ml chloramphenicol to maintain the pLys plasmid. Isopropyl--D-thiogalactopyranoside was then added to a final concentration of 1 mM to induce RNase R expression, and 100 µg/ml additional ampicillin was added to maintain pETR. One h after adding isopropyl--D-thiogalactopyranoside, the culture was quickly cooled on ice, and cells were harvested by centrifugation. Five-ml portions of culture were taken before and after induction for analysis. The cell pellet was frozen at 80 °C until used.

Preparation of Cell Extracts-- To determine the amount of RNase R expressed from plasmid pETR, cells from the 5-ml portions of culture taken before and after isopropyl--D-thiogalactopyranoside induction were resuspended in 1 ml of buffer containing 10 mM Tris-Cl (pH 7.6), 1 mM dithiothreitol, 500 mM KCl, and 0.1 mM phenylmethylsulfonyl fluoride. Extracts were prepared by sonication using two 15-s pulses.

For RNase R purification, ~12 g of wet cells overexpressing RNase R were suspended in 60 ml of buffer A (10 mM Tris-Cl (pH 7.6), 1 mM dithiothreitol, 10 mM MgCl₂, 0.1 mM phenylmethylsulfonyl fluoride, and 15 units/ml DNase I). Cells were ruptured by two passes through an Aminco French press at 12,000 p.s.i. The suspension was centrifuged at 30,000 × g for 2 h. The pellet obtained was resuspended in 30 ml of buffer B500 (10 mM Tris-Cl (pH 7.6), 1 mM dithiothreitol, 500 mM KCl, 0.5 mM EDTA (potassium salt; pH 7.4), 10% glycerol, and 0.1 mM phenylmethylsulfonyl fluoride). The sample was centrifuged at 30,000 × g for 45 min and then at 150,000 × g for 2 h to obtain an S150 supernatant fraction. All steps of the purification were carried out at 4 °C or below.

Affi-Gel Blue Chromatography-- The S150 fraction (7.5 ml) was loaded onto an Affi-Gel blue column (0.8 × 10 cm) prepared in buffer B500, washed with 3 bed volumes of the same buffer, and then eluted with 2 bed volumes of the same buffer containing 1 M KCl. Peak fractions were combined and dialyzed against buffer B500 to lower the KCl concentration. The dialyzed sample of purified RNase R was quickly frozen in an ethanol/dry ice bath and stored at 80 °C.

Gel Filtration-- For molecular mass measurement, purified RNase R was applied to an Ultrogel AcA44 gel filtration column (1.0 × 56.5 cm) using either buffer B500 (containing 500 mM KCl) or buffer A (containing no KCl) as the running buffer. Blue dextran 2000 and bacterial alkaline phosphatase (86 kDa) were added to the samples and co-chromatographed as internal molecular mass markers.

Enzyme Assays-- RNase R assays were carried out as described (7) using [³H]poly(A) as substrate, unless indicated. The activity of RNase R is expressed as nmol of nucleotide released in 1 min at the given temperature, usually 37 °C. RNase II assays were carried out in 50-µl reactions containing 10 mM Tris-Cl (pH 8.2), 100 mM KCl, 10 mM MgCl₂, 36 µg of [³H]poly(A), and 0.06 µg of RNase II. The activity of RNase II is expressed as described for RNase R. Acid-soluble radioactivity was measured as described (7). Bacterial alkaline phosphatase assay of the gel filtration column fractions was carried out by mixing 10 µl of each column fraction with 1 ml of 1 mM p-nitrophenyl phosphate dissolved in 1 M Tris-Cl (pH 8.0) in a cuvette and measuring the A₄₁₀/min (11).

Protein Determination-- Protein was determined by the method of Bradford (12) or by absorbance at 280 nm.

SDS-PAGE-- Proteins were resolved on 8% SDS-polyacrylamide gels (13). Gels were stained with Coomassie Brilliant Blue R-250 to visualize the protein bands.

N-terminal Sequencing-- Twenty µg of purified RNase R was resolved on an 8% SDS-polyacrylamide gel and transferred to a polyvinylidene difluoride membrane by electroblotting. The membrane was stained with Ponceau S. Protein bands were subjected to N-terminal sequencing by the Protein Analysis Facility of our department.

Preparation of Ribosomes and rRNAs-- Ribosomes were isolated from E. coli cells (usually in log phase) as described (14). To isolate rRNAs, 10 µl of 20% SDS was added to 400 µl of the ribosome suspension. The mixture was extracted once with an equal volume of buffered phenol (buffered with 20 mM sodium acetate (pH 5.3)) and once with an equal volume of buffered phenol/chloroform/isoamyl alcohol (25:24:1), and then 2.5 volumes of 95% ethanol was added to the aqueous phase to precipitate rRNA. The total rRNA pellet was dissolved in 200 µl of diethyl pyrocarbonate-treated H₂O and resolved on a 1% agarose gel (15). 23 S, 16 S, and 5 S RNAs were then isolated using RNaid w/Spin kit.

Preparation of a Substrate with a 3'-Phosphate-- An oligonucleotide substrate containing a 3'-phosphate terminus (A₁₆P) was generated by periodate oxidation of A₁₇ (16).

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Overexpression and Purification of RNase R-- Upon isopropyl--D-thiogalactopyranoside induction of strain BL21II(DE3)/pLys harboring plasmid pETR, RNase R was overexpressed to an extent that it was the most abundant protein in the sonicated extract (Fig. 2, compare lane 2 with lane 1). Quantitatively, with poly(A) as substrate, an extract of cells collected after induction had a specific activity of 1,550 nmol/min/mg of protein, which was ~100-fold higher than that of an extract from uninduced cells.

View larger version (28K): [in this window] [in a new window]   Fig. 2.   SDS-PAGE of RNase R purification fractions. Ten µg of protein from each fraction was denatured and loaded on an 8% SDS-polyacrylamide gel. After electrophoresis, the gel was stained with Coomassie Brilliant Blue R-250. Lane 1, sonicated cell extract before induction; lane 2, sonicated cell extract after induction; lane 3, no-salt soluble fraction; lane 4, S150; lane 5, pooled Affi-Gel blue (AGB) peak fractions. The migration positions of molecular mass standards are shown on the left (in kDa). The position of RNase R is indicated by the arrow.

RNase R was rapidly purified from the induced cell extract by taking advantage of its insolubility in low salt (4). Thus, most of the proteins in the extract were soluble in buffer A (no KCl), whereas RNase R was not detectable in this fraction (Fig. 2, lane 3). However, resuspension of the pellet in buffer B500 (containing 500 mM KCl) and recentrifugation revealed that RNase R was the major protein in the S150 supernatant fraction (Fig. 2, lane 4). These precipitation steps resulted in an ~5-fold purification of RNase R (Table I). A portion of the S150 fraction was further purified on an Affi-Gel blue column, leading to an additional 2-3-fold purification (Table I). SDS-PAGE of the purified RNase R indicated that it was at least 95% pure (Fig. 2, lane 5). Based on this simple procedure, 7.5 mg of highly purified RNase R was obtained from the equivalent of 3 g of wet cells (Table I).

Spectral Analysis-- Spectral analysis of purified RNase R in the range of 200-700 nm revealed no unusual peaks (data not shown). The A₂₈₀/A₂₆₀ ratio was 1.87, indicating that purified RNase R does not contain nucleic acid and therefore that RNase R does not require nucleic acid for activity.

Molecular Mass of RNase R-- When gel filtration of purified RNase R was performed in buffer B500 (containing 500 mM KCl), RNase R eluted as a globular protein of ~95 kDa, in close agreement with its predicted size of 92 kDa based on the rnr gene sequence. This observation supports our previous suggestion that RNase R is a monomer (7). In contrast, when gel filtration was carried out in buffer A (lacking KCl), RNase R eluted in the void volume, indicating that it aggregates at low ionic strength, as originally observed by Kasai et al. (4) with partially purified enzyme. The fact that activity could be detected when the protein was aggregated suggests either that aggregation does not abolish RNase R activity or that the protein dissociates in the assay mixture.

N-terminal Sequencing of RNase R-- To confirm that RNase R is encoded by the rnr gene and also to conclusively determine which of the two potential translation start sites is used, purified RNase R was subjected to N-terminal sequencing. The enzyme was resolved by 8% SDS-PAGE, followed by electroblotting onto a polyvinylidene difluoride membrane. Subsequent N-terminal analysis revealed the sequence SQDPFQE, in agreement with that predicted in the Swiss Protein Database, which is MSQDPFQE. These data indicate that 1) the rnr gene encodes RNase R; 2) the open reading frame predicted in the Swiss Protein Database is correct, but that the N-terminal formyl-Met residue is removed; and 3) the downstream AUG codon is the initiation site for RNase R synthesis.

Heat Stability of RNase R-- To determine the response of RNase R to heat, its activity was measured at various temperatures. RNase II activity was measured at the same temperatures for comparison. Both RNase R and RNase II displayed similar temperature activity profiles, being most active at ~50 °C in a 5-min assay. RNase R stability at 50 °C was further examined by preincubating at that temperature for varying lengths of time and then assaying at 37 °C. The enzyme was relatively stable, losing ~25% of its activity in 15 min and ~80% over a 30-min period (data not shown).

Optimal Reaction Conditions for RNase R Activity on Poly(A)-- Purified RNase R was assayed on poly(A) under a variety of conditions to assess its requirements for optimal activity; RNase II was assayed in parallel for comparison. Both RNase R and RNase II displayed a broad optimal pH range between 7.5 and 9.5. Both enzymes required a divalent cation, preferably Mg²⁺, for activity. In the presence of 1 mM EDTA, the activity of each enzyme was abolished. However, the optimal Mg²⁺ concentrations for the enzymes were quite different. RNase R was most active at 0.1-0.5 mM Mg²⁺, whereas RNase II was most active at 10 mM Mg²⁺. Both RNase R and RNase II were stimulated by the presence of a monovalent cation. RNase R was stimulated ~2-fold by 50-500 mM K⁺ and 40% more by Rb⁺. For RNase II, K⁺ was the most stimulatory of the monovalent cations tested, increasing activity ~5-fold over a similarly wide range of concentrations (data not shown).

Mechanism of Action of RNase R-- Earlier work with partially purified enzyme indicated that RNase R is an exoribonuclease releasing 5'-nucleoside monophosphates (4). To ensure that the product released was not due to a secondary reaction by a contaminating activity, we re-examined the mechanism of action of RNase R using the purified enzyme. Thus, when 5'-³²P-labeled RNA substrates (of varying length from oligomer to polymer) were digested by purified RNase R, intermediates with sizes between the starting material and final product were not observed to accumulate (Fig. 3 and data not shown). This observation is consistent with the conclusion that RNase R is a processive exoribonuclease. To confirm this point, the products of the reaction catalyzed by highly purified RNase R were examined.

View larger version (78K): [in this window] [in a new window]   Fig. 3.   Digestion of oligonucleotide substrates by RNase R and RNase II. Oligonucleotide substrates were labeled at their 5'-ends with 32P. Reactions were carried out at 37 °C in 50-µl reaction mixtures containing 20 mM Tris-Cl (pH 8.2), 100 mM KCl, and 10 µM labeled oligomer. For RNase R reactions, MgCl2 was present at 0.25 mM, and 0.15 µg of purified RNase R was added; for RNase II reactions, MgCl2 was present at 10 mM, and 0.06 µg of purified RNase II was added. Nine-µl aliquots were taken at 0, 5, 10, 15, and 20 min as indicated, and the reactions were stopped with 2 volumes of RNA loading buffer. The upper panel (marked R) presents the results of RNase R digestion, and the lower panel (marked II) presents the results of RNase II digestion.

If RNase R were an exoribonuclease, the major acid-soluble product with [³H]poly(A) as substrate would be expected to be AMP. Upon treatment with alkaline phosphatase, this would be converted to the uncharged molecule, [³H]adenosine, and would elute with the nucleoside fraction from an anion-exchange column (Dowex AG 1-X2). If, on the other hand, RNase R were an endoribonuclease, the acid-soluble oligonucleotides produced would remain charged after phosphatase treatment and would elute with the "nucleotide" fraction. As shown in Table II, 90% of the acid-soluble radioactivity was eluted from the anion-exchange column with water, consistent with its being the nucleoside, adenosine.

A second test of the mechanism of RNase R was its action on tRNA. Because tRNA is a relatively poor substrate of RNase R, we reasoned that RNase R might act on it in a distributive fashion such that tRNA-CC might be generated due to removal of 1 AMP residue from tRNA-CCA. This product would be a substrate for the re-incorporation of AMP by tRNA nucleotidyltransferase. As shown in Table II, this prediction was fulfilled. Thus, there was a dramatic increase in AMP incorporation into tRNA by tRNA nucleotidyltransferase that was dependent on the amount of RNase R added (40 pmol with 0.15 µg of RNase R and 578 pmol with 3 µg of the enzyme). The incorporation of [³H]AMP in the absence of RNase R was due to the presence of some tRNA-CC in the tRNA preparation. Nevertheless, these data show that RNase R can slowly remove a single AMP residue from tRNA to generate tRNA-CC, supporting the conclusion that RNase R is an exoribonuclease.

Substrate Specificity of RNase R-- Important clues to the physiological role of RNase R may come from an analysis of its substrate specificity. Earlier work suggested that RNase R is a nonspecific enzyme able to act on homopolymers, rRNA, and mRNA (4, 7). Inasmuch as the previous studies of RNase R were carried out with only partially purified preparations, we felt that it was important to examine the specificity of the purified enzyme. We have used a greatly expanded catalogue of substrates and also compared its specificity with that of its homolog, RNase II.

The relative activities of RNase R and RNase II on poly(A) and the major classes of RNA are presented in Table III. RNase II was ~4-fold more active on poly(A) compared with RNase R, whereas RNase R was much more active on the natural RNAs. Both enzymes worked best on poly(A). The K_m value for poly(A) in the RNase R reaction was ~30 nM. Among the natural RNAs, RNase R worked well on 23 S and 16 S RNAs, but relatively poorly on 5 S RNA and tRNA. Nevertheless, 5 S RNA and tRNA were potent inhibitors of RNase R action on poly(A) (data not shown). Fig. 4 shows that the acid-soluble material released from the 16 S and 23 S rRNA preparations actually led to the disappearance of the rRNA bands and thus could not be due to degradation of a contaminating RNA. In separate experiments using gel analysis rather than acid solubility, degradation of homopolymers by RNase R was in the order poly(A) > poly(U) > poly(C) poly(G) (data not shown). The latter polymer was essentially inactive as a substrate. The limit products of digestion of the poly(A), poly(U), and poly(C) homopolymers were di- and trinucleotides in each instance.

View larger version (60K): [in this window] [in a new window]   Fig. 4.   Digestion of 16 S and 23 S rRNAs by RNase R and RNase II. Fifty µg of total RNA (purified from isolated ribosomes) was treated with 0.15 µg of RNase R or RNase II in 50-µl reactions under the conditions described under "Experimental Procedures," except that KCl was present at 100 mM. Nine-µl aliquots were taken at the times indicated, and reaction products were resolved on a 1.1% agarose gel, followed by Northern hybridization using oligomers complementary to the 5'-ends of 16 S and 23 S RNAs. The oligomer used to detect 16 S RNA was 5'-CCTGTTACCGTTCGACTTGC-3', and the oligomer used to detect 23 S RNA was 5'-CCTTCATCGCCTCTGACTGCCA-3'. Quantitation was done on a PhosphorImager (Molecular Dynamics, Inc.).

The activities of RNase R and RNase II on A oligonucleotides 6-17 nucleotides in length were also examined. Both RNase R and RNase II could hydrolyze RNAs as short as a 6-mer (A₆); however, although RNase R acted processively for RNA oligomers as short as 8-mers, RNase II tended to be more distributive, especially for the shorter substrates (Fig. 3). The limit end products of the RNase R reaction were di- and trinucleotides, whereas the major end products of the RNase II reaction were oligonucleotides ranging from 3 to 5 residues in length (Fig. 3). Neither RNase R nor RNase II displayed much activity against DNA oligomers at the enzyme levels usually used. However, when 30-40 times this amount of enzyme was tested, RNase II could hydrolyze the DNA oligomer dT₁₇, in a distributive manner; RNase R worked much more poorly, removing only 1-2 residues from a portion of the substrate molecules (Fig. 5). Moreover, the DNA oligomer dC₁₇ was a potent inhibitor of RNase II activity with poly(A) as substrate, whereas it had little effect on RNase R (data not shown).

View larger version (67K): [in this window] [in a new window]   Fig. 5.   Digestion of the DNA oligomer dT17 by RNase R and RNase II. dT17 was labeled at its 5'-end with 32P. Reactions were carried out at 37 °C for 20 min under the conditions of the T4 polynucleotide kinase reaction: 70 mM Tris-Cl (pH 7.6), 10 mM MgCl2, and 5 mM dithiothreitol with 5 µM dT17 and the indicated amounts of RNase R (R) or RNase II (II).

To examine whether RNase R can hydrolyze double-stranded molecules, a 5'-³²P-labeled RNA oligomer (17-mer) was subjected to RNase R action in the presence of a complementary DNA either 13 or 17 nucleotides in length or a fully complementary RNA (Fig. 6). In the presence of a 2-fold molar excess of the complementary 13-mer (to generate a substrate with 13 base pairs and a 4-nucleotide extension at the 3'-end) or in the presence of a complementary DNA or RNA 17-mer (to generate a completely base-paired substrate), no shortening was evident. In fact, for the assay of the RNA-RNA duplex, 20 times the usual amount of RNase R was used. On the other hand, when even a 20-fold excess of a non-complementary DNA (oligomer dT₁₇) was present, RNase R degraded the RNA substrate at a rate comparable to that when no complementary oligomer was present (Fig. 6). These data show that the complementary strands inhibit RNase R action and that RNase R cannot act on double-stranded molecules even when a 4-nucleotide 3'-RNA overhang is present.

View larger version (39K): [in this window] [in a new window]   Fig. 6.   Digestion of a 17-nucleotide RNA oligomer by RNase R in the presence or absence of a complementary oligomer. An RNA 17-mer was labeled at its 5'-end with 32P. Reactions were carried out as described in the legend to Fig. 3. for the times indicated. Complementary oligomers were present at 20 µM, and dT17 (the non-complementary oligomer) was present at 200 µM. Twenty-fold as much RNase R was present when the substrate was the RNA-RNA duplex.

To ascertain whether RNase R can act on an RNA molecule with a 3'-phosphoryl terminus, its ability to degrade A₁₆P was determined. As shown in Fig. 7, both RNase R and RNase II could function on a substrate with a 3'-phosphate group. The rate at which A₁₆P was degraded by each enzyme was comparable to that previously observed with A₁₇ (see Fig. 3).

View larger version (76K): [in this window] [in a new window]   Fig. 7.   Action of RNase R and RNase II on an oligomer substrate with a 3'-phosphate group. A16P was labeled at its 5'-end with 32P. Reactions were carried out as described in the legend to Fig. 3 for the times indicated. R, RNase R; II, RNase II.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Using the rapid purification procedure described here, which takes advantage of the differential solubility of RNase R in high and low salt, we were able to purify mg quantities of essentially homogeneous RNase R in its native untagged form from 1 liter of culture in just 2 days. Based on activity against rRNA, the preparation obtained was ~20-fold purer than that described by Kasai et al. (4). Moreover, the RNase R described here displayed a native molecular mass of ~95 kDa, as expected based on the sequence of the rnr gene (7). This contrasts with a mass of ~40-50 kDa reported by Kasai et al. (4) based on a comparison of the size of RNase R with that of RNase II. In some preparations of RNase R, we have also found a smaller active form of RNase R that consisted of the central region and the C-terminal S1 RNA-binding domain of the protein (1). Inasmuch as this form of the enzyme could be eliminated by inclusion of EDTA in the buffers during purification and based on the fact that the eliminated N-terminal fragment could be detected by SDS-PAGE (data not shown), we conclude that the smaller protein arises by proteolytic cleavage due to a metal-dependent protease. It is possible that this may explain the form of RNase R purified by Kasai et al. (4). These workers also reported that RNase R was associated with ribosomes. However, this has not been observed with the overexpressed protein.

Both RNase R and RNase II are members of the RNR superfamily of exoribonucleases (1). Structurally, both are large monomeric proteins. Likewise, RNase R and RNase II share many catalytic properties. Thus, both enzymes are relatively nonspecific processive 3'  5' exoribonucleases that release 5'-nucleoside monophosphates and leave a residual oligoribonucleotide core. We have found that this core is somewhat smaller with RNase R (di- and trinucleotides) than is observed with RNase II (tri- to pentanucleotides) (this work and Refs. 17 and 18). The inability of these enzymes as well as polynucleotide phosphorylase to act on small oligoribonucleotides is the apparent reason for the existence of an additional RNase in most cells (termed oligoribonuclease) that digests such residual molecules (19). Another similarity in the properties of RNase R and RNase II is that both enzymes can act on 3'-phosphate-terminated RNA molecules, indicating that neither of them requires a free 3'-hydroxyl group to initiate degradation.

On the other hand, there are a number of significant differences in the actions of RNase R and RNase II. For example, the latter enzyme works relatively efficiently with DNA, and its action on RNA is strongly inhibited by DNA oligomers. In contrast, RNase R acts very poorly on DNA, and its activity is scarcely affected by the presence of a DNA oligomer. Based on this information, it is likely that RNase R binds relatively weakly to DNA. A second major difference between the two RNases is their substrate specificity. Although both enzymes work most effectively on synthetic homopolymers such as poly(A), RNase R also can degrade rRNAs quite well. However, RNase II is essentially inactive against such natural RNA substrates. Considering that rRNA molecules contain extensive secondary structure, these observations suggest that RNase R may be much more effective than RNase II in digesting through such structured regions. It is already known that RNase II slows down as it approaches within 10 nucleotides of a double-stranded RNA structure (20). The data presented here show that RNase R cannot degrade a completely double-stranded short RNA molecule or one with a 4-nucleotide 3'-overhang. Nevertheless, the fact that RNase R can digest both 16 S and 23 S rRNA molecules indicates that it is able to work through secondary structure in the context of the rRNA. How this is accomplished remains to be determined.

The in vivo role of RNase R is not yet known, but the data presented here indicate that the enzyme would be capable of acting on a variety of natural RNA molecules. Perhaps, its ability to act on structured RNAs is the reason why E. coli maintains this enzyme in addition to RNase II.

	ACKNOWLEDGEMENT

We thank Dr. Cecília Arraiano for providing a strain.

	FOOTNOTES

^* This work was supported by Grant GM16317 from the National Institutes of Health.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence and reprint requests should be addressed: Dept. of Biochemistry and Molecular Biology, University of Miami School of Medicine, P. O. Box 016129, Miami, FL 33101. Tel.: 305-243-3150; Fax: 305-243-3955; E-mail: mdeutsch@med.miami.edu.

Published, JBC Papers in Press, April 10, 2002, DOI 10.1074/jbc.M202942200

¹ Z.-F. Cheng and M. P. Deutscher, unpublished data.

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TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

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