Exoribonucleases and Endoribonucleases
Module
4.6.3
ZHONGWEI LI1* AND MURRAY P. DEUTSCHER2*
[SECTION EDITOR: GLENN BJÖRK]
Posted December 29, 2004
1Department of Biomedical Sciences, Florida Atlantic University, 777 Glades Road, BC308, Boca Raton, FL 33431-0991, and 2Department of Biochemistry and Molecular Biology, University of Miami School of Medicine, P.O. Box 016129, Miami, FL 33101-6129
*Corresponding authors. Z. Li, Phone: (561) 297-3178; Fax: (561) 297-2221; E-mail:
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. M. P. Deutscher, Phone: (305) 243-3150, Fax: 305-243-3955, E-mail:
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Ribonucleases (RNases) play a central role in many aspects of RNA metabolism. The reactions catalyzed by this group of enzymes include those that process RNA precursors to their mature, functional forms, some that convert RNA molecules to alternate forms, and those that degrade RNAs that are damaged or no longer needed by the cell. As our knowledge of the intricacies of RNA metabolism has grown, so has the number of RNases needed to carry out these processes, and it is now clear that a single cell can contain a multitude of such enzymes (e.g., approaching 20 in Escherichia coli). While considerable progress in the study of RNases has been made for several organisms in recent years, the depth of our understanding remains highest for E. coli. Thus, physiological roles for many of the E. coli RNases have now been identified, and structural and mechanistic studies of the enzymes are progressing rapidly.
RNases can be placed into two groups based on their mode of action. The exoribonucleases digest RNA molecules from either a 3' or 5' terminus, releasing nucleotide residues. Some exoribonucleases act processively, releasing residues continuously as they traverse a substrate without dissociating. Depending on the enzyme, digestion may go to completion or may stop due to the presence of secondary structure in the RNA. Other exoribonucleases act distributively, dissociating and rebinding after one or a few catalytic events. In eubacteria, the known exoribonucleases all initiate degradation at the 3' terminus of the RNA and release 5' mononucleotides (79, 353). Inasmuch as eubacterial mRNAs lack a 5' cap structure, the absence of 5' to 3' exoribonucleases may serve to avoid unwanted degradation initiating from this end of the molecule.
RNases in the second category, the endoribonucleases, cleave RNA molecules internally, releasing RNA fragments of various sizes. The action of these enzymes may be highly specific, cleaving at only one or a few sites per RNA molecule, or nonspecific, cleaving throughout the RNA chain and leading to extensive degradation. Depending on the reaction mechanism of the RNase, cleavage can be on either side of the phosphodiester linkage.
In this chapter, we provide a description of the known E. coli RNases, focusing on their structures, catalytic properties, genes, physiological roles, and possible regulation. For primary references, emphasis will be placed on more recent literature. The role of RNases in mRNA decay is treated in detail in another EcoSal chapter, "mRNA Stability" (chapter 4.6.4), by Kushner. Other recent reviews dealing with various aspects of E. coli RNases can be found in references 37, 58, 79, 156, and 353.
Currently, eight E. coli exoribonucleases are known. These are RNases II, R, D, T, PH, BN, polynucleotide phosphorylase (PNPase), and oligoribonuclease (ORNase) (Table 1). As noted, all digest RNA in the 3' to 5' direction. Six of the RNases release nucleoside 5' monophosphates, and two, PNPase and RNase PH, release nucleoside diphosphates. Based on sequence analysis and catalytic properties, the eight exoribonucleases have been grouped into four families (353). These are the RNR family, including RNase II and RNase R; the DEDD family, including RNase D, RNase T, and ORNase; the RBN family, consisting of RNase BN; and the PDX family, including PNPase and RNase PH.
Table 1E. coli exoribonucleases |
The two members of the RNR family are large, single-chain proteins. Both contain a C-terminal S1 RNA-binding domain (27) and, in addition, a conserved central region of ~400 amino acids with several highly conserved motifs (353). Nevertheless, the two subfamilies can be distinguished, and although E. coli contains a member of each subfamily, many other eubacteria contain only an RNase R homologue (58, 353). Both RNase II and RNase R are processive, nonspecific exoribonucleases; however, their catalytic properties differ significantly (44).
RNase II.
This enzyme is the most active exoribonuclease in extracts of E. coli, accounting for as much as 98% of hydrolytic activity when poly(A) is the substrate (46). Because of its high level of activity, RNase II was identified and purified early on (293). Over the years, it has been purified to homogeneity in various laboratories from both normal and overexpressing strains (e.g., see reference 54). Based on the sequence of the rnb gene, RNase II is a protein of 644 amino acids with a calculated molecular mass of 72.5 kDa (54). Upon gel filtration, RNase II in extracts migrates with an apparent molecular mass of ~80 kDa (81), confirming that it is a single-chain protein and suggesting that it is not associated with other macromolecular components.
RNase II can act on a variety of RNA substrates in a reaction that requires Mg2+ (10 mM optimum), and it is stimulated by monovalent cations (293). However, its action on synthetic homopolymers exceeds that on natural RNAs, particularly stable RNAs, by as much as several hundredfold (44), most likely due to the extensive secondary structure of the latter substrates. RNase II is sensitive to secondary structure both in vitro and in vivo. Studies with a synthetic RNA substrate in vitro indicate that RNase II slows dramatically as it approaches within ~10 nucleotides (nt) of a double-stranded region (54). However, the strength of the double-stranded region affects the stalling and dissociation of RNase II from structured substrates (298). Nevertheless, RNase II can bind to short, single-stranded regions (36), and, given sufficient enzyme, limit products 3 to 5 nt in length can be generated (44). Moreover, in vivo, RNase II can generate mature 3' termini on tRNAs (176), indicating that it can digest to within 4 nt of the double-stranded aminoacyl stem. Other studies with synthetic RNA substrates have led to the suggestion that RNase II contains an anchoring site that binds residues 15 to 25 nt from the 3' end of the substrate and that during catalysis residues at this site remain fixed while the 3' end threads through the catalytic site (35, 36). However, such an anchoring site cannot be essential for RNase II action given that much shorter molecules can be substrates for the enzyme.
In addition to its action on RNA, RNase II can also digest a single-stranded DNA oligomer, dT17, at a slow rate (44). The extent of its action on other DNA substrates remains to be explored. RNase II does not require a free 3'-hydroxyl group to initiate degradation since the presence of a 3'-phosphoryl does not hinder its action (44).
RNase II appears to be involved primarily in mRNA metabolism in vivo. However, its action in this regard is somewhat paradoxical. Together with PNPase, RNase II participates in mRNA degradation, and, in the absence of both enzymes, fragments of mRNA accumulate (88). On the other hand, 10-fold overexpression of RNase II does not alter mRNA degradation (87), and in some cases RNase II actually stabilizes mRNA against degradation by removing the poly(A) tails that are needed for degradation of mRNAs with stable terminal hairpins (54). A genome-wide analysis of the stabilizing effects of RNase II revealed that steady-state levels of 31% of E. coli mRNAs are decreased in the absence of RNase II, whereas only 7% are increased (227). Based on these data, a critical role of RNase II may be to protect mRNAs against the action of other exoribonucleases (203). It has also been suggested that RNase II may interfere with the action of an endoribonuclease, RNase E (66).
RNase II can also participate in the metabolism of stable RNAs. In the absence of other exoribonucleases, RNase II becomes an important enzyme for the maturation of tRNA (176; see Fig. 1) and other small, stable RNA species (178). However, when other exoribonucleases are present, there is no evidence for a significant role for RNase II in stable RNA metabolism. This conclusion is supported by the fact that RNase II− single mutants display no obvious phenotype. RNase II levels do affect the length of poly(A) tails on 23S rRNA (224); however, the significance of this polyadenylation is not understood.
RNase II is encoded by the rnb gene at 29.0 min of the E. coli genetic map (283, 349). The gene is monocistronic but contains two functional promoters (350). The amount of RNase II in cells can be influenced by other RNases. For example, RNase E acts on the RNase II message, and, as a consequence, rnb mRNA and RNase II activity increases when RNase E is inactivated (351). Likewise, PNPase modulates RNase II expression (350). In PNPase-deficient strains, RNase II levels are elevated, and in PNPase-overproducing strains, they are decreased, leading to an overall variation of ~fivefold. Likewise, PNPase is elevated in an rnb deletion strain. The mechanism by which these two RNases are interrelated is not yet known, but it may indicate significant cross-regulation among RNases with overlapping functions. RNase II is also regulated by a protein, termed Gmr, which influences the stability of RNase II protein (28). In the absence of Gmr, RNase II can be elevated as much as threefold. RNase II levels are also affected by growth media. However, the physiological relevance of all these regulatory phenomena remains to be elucidated since RNase II levels are not known to vary under normal conditions.
RNase R.
In the absence of RNase II, the major hydrolytic exoribonuclease in extracts active on synthetic polynucleotides is RNase R (46). Studies of the rnr (vacB) gene encoding RNase R indicates that it is a protein of 813 amino acids with a calculated molecular mass of 92.1 kDa (46), in close agreement with its size determined by gel filtration (44). Recently, RNase R was purified to apparent homogeneity from an overexpressing strain (44), and, as was found with the partially purified protein (147), it has a strong tendency to self-aggregate, particularly at salt concentrations below 300 mM (44). This property may complicate studies to determine whether RNase R associates with other proteins.
Purified RNase R is most active at pH 7.5 to 9.5, 0.1 to 0.5 mM Mg2+, and 50 to 500 mM monovalent cation (44). As with RNase II, RNase R is most active against synthetic polynucleotides, such as poly (A). However, in contrast to the former enzyme, RNase R displays significant activity against ribosomal RNAs and tRNA (44). In fact, 16S and 23S rRNAs can be digested essentially to completion. Such data indicate that RNase R can work through extensive secondary structure. On the other hand, RNase R cannot act on a perfect double-stranded 17-mer oligoribonucleotide or on a 13-mer with a 4-nt single-strand extension (44). These data imply that RNase R may need a single-stranded region to bind and to initiate degradation. In contrast to RNase II, RNase R is essentially inactive on the DNA substrate, dT17, but like RNase II, it can act on a 3'-phosphoryl-terminated oligonucleotide. Limit digests are 2 to 3 nt in size. On short oligonucleotides, 6 to 17 nt in length, RNase R acts processively, whereas RNase II becomes distributive (44).
RNase R is encoded by the rnr (formerly vacB) gene located at 94.9 min on the E. coli genetic map (46, 283). Although not yet studied in detail, rnr appears to be part of a three-gene operon, with the unknown yjeB gene upstream and rlmB, which encodes a 23S rRNA methyltransferase, downstream (46 and unpublished observations). Single mutant strains lacking RNase R grow normally on rich or minimal media at temperatures between 31 and 42°C. Likewise, multiple mutant strains lacking RNase R together with RNases I, II, D, BN, with RNase T or with RNase PH, are all viable. In contrast, double mutant strains lacking RNase R and PNPase do not grow (45, 46), suggesting that the two exoribonucleases overlap in some important function. Analysis of a temperature-sensitive mutant strain lacking RNase R and containing a PNPts mutation revealed that, in the absence of the two RNases, fragments of 16S and 23S rRNA accumulate to high levels and ribosome assembly is defective (45). These data revealed that RNase R and PNPase participate in a quality control process that normally removes these rRNA fragments as rapidly as they are generated. The source of these fragments is not yet known, but they could arise at some frequency during ribosome assembly or by premature transcription termination. Recent evidence indicates that RNase R also plays a role in mRNA decay (Z.-F. Cheng and M. P. Deutscher, unpublished observation).
RNase R is a cold-shock protein, increasing 7- to 8-fold after transfer of a culture to 10°C (29). Induction is thought to result from stabilization of the rnr message. PNPase appears to play a role in the ultimate decay of the rnr message at the end of the acclimatization period. It has also been proposed that RNase R is important both for growth and for tmRNA maturation under cold-shock conditions (29). RNase R levels also increase under other stress conditions, such as stationary phase and starvation (C. Cheng and M. P. Deutscher, unpublished observation). What the role of elevated RNase R might be under these various stress conditions remains to be determined.
Three RNase members of the DEDD family are present in E. coli: RNase D, RNase T, and ORNase. While their overall structures differ, the three RNases share a common catalytic core that contains four invariant acidic residues as well as other conserved residues distributed in three separate motifs (353). The nucleases of this superfamily, which includes many DNA exonucleases, are thought to act by a common catalytic mechanism that involves two metal ions (306). The DEDD nucleases fall into two subfamilies based on whether they contain a histidine or tyrosine in motif III (353). RNase D is a member of the DEDDy subfamily, and RNase T and ORNase are in the DEDDh subfamily. The latter two enzymes also are more closely related structurally and catalytically (see below).
RNase D.
The discovery of RNase D was the first indication that exoribonucleases could display a high degree of specificity (70, 71). The other exoribonucleases (RNase II and PNPase) and phosphodiesterases known at that time all displayed broad substrate specificities. In contrast, RNase D was specific for tRNA-like molecules and was essentially inactive against homopolymers, such as poly (A). In fact, RNase D was originally identified because of its action on "denatured" tRNAs (105). Subsequently, it was shown to be able to generate mature tRNA from an artificial tRNA precursor, suggesting that its role in vivo might be in the 3' maturation of tRNA (67).
RNase D has been purified to homogeneity from normal (70) and overexpressing (338) strains. Based on its deduced amino acid sequence and gel filtration, RNase D is a single-chain protein of 375 amino acids with a molecular mass of 42.7 kDa (70, 338). Moreover, there is no evidence that RNase D associates with other proteins in cell extracts. Recently, the crystal structure of RNase D has been resolved to 1.6 Å (Y. Zuo and A. Malhotra, personal communication). The protein consists of three domains that form a funneled ring structure. The catalytic center containing the DEDD residues is found on one of these domains in a region rich in ß-sheet. The other two domains, which contain mainly α-helical structure, form a region of positively charged residues on the side of the funnel away from the catalytic center. This region may serve as the RNA substrate-binding site.
In vitro, RNase D is most active on tRNA molecules containing residues following the mature 3' terminus or on molecules lacking all or part of the –CCA sequence. It is much less active on mature tRNA (69). The relative resistance of mature tRNA does not appear to be due to the –CCA sequence per se because a second –CCA sequence added following the original one can be removed rapidly (69). Rather, the data suggest that RNase D is sensitive to tRNA structure. This conclusion is supported by the increased activity of RNase D on denatured tRNA (105). RNase D also is sensitive to the 3'-terminal residue; tRNA-C-Cp, containing a 3'-terminal phosphate residue, is inactive as a substrate, and tRNAs that have been oxidized by periodate or oxidized and reduced by borohydride are much less active than those with an intact 3'-terminal ribose (69). However, the identity of the 3'-terminal base does not appear to play a role.
RNase D is most active at pH 8 to 10. A divalent cation, which may be Mg2+, Mn2+, or Co2+, is required for activity. RNase D acts on tRNA distributively, dissociating from the substrate upon removal of the terminal residue (69).
The physiological role of RNase D is much less clear. Strains devoid of RNase D grow essentially normally and plate bacteriophage T4 normally (20). However, RNase D becomes essential for viability when other exoribonucleases, RNases II, BN, T, and PH, are eliminated (153). Cells lacking the four exoribonucleases, but retaining RNase D, grow relatively poorly, doubling in 64 min compared with 24 min for wild-type cells, indicating that RNase D cannot take over completely the functions of the missing RNases. RNase D can act on tRNA molecules in vivo when it is present at elevated levels (339) or when the aforementioned exoribonucleases are absent (176, 275), but its action on precursors of different tRNA species varies (Fig. 1). In the absence of the other RNases, RNase D also participates in the maturation of several small E. coli RNAs (178). Thus, RNase D can function as a backup enzyme when the primary nucleases are missing, but when they are present, it is not required. However, the primary function of RNase D that has resulted in its maintenance in E. coli probably has not yet been discovered.
RNase D is encoded by the rnd gene located at 40.6 min on the E. coli genetic map (283, 337, 340). Extensive deletion analysis and primer extension studies identified a single promoter upstream of the RNase D coding region (341). The start site of transcription is located 70 nt upstream of a UUG initiation codon. RNase D expression is regulated not only by the UUG codon but also by other sequences in the upstream region, particularly an apparent rho-independent terminator immediately downstream of the promoter. Mutagenesis of this region revealed that, in fact, the apparent transcription terminator affected RNase D expression at the translational level (341, 342). Thus, removal of the stem-loop structure has no effect on levels of rnd mRNA, but it does affect the amount of RNase D. The sequence of eight U residues following the stem-loop were shown to serve as a ribosome-binding site, and their mutation decreased RNase D protein and activity by as much as 95%. Overall RNase D expression is affected by the UUG codon, the Shine-Dalgarno sequence, the stem-loop structure, and the U-rich region (342). However, how these unusual features actually modulate RNase D levels in vivo has not been established.
RNase T.
The end-turnover of tRNA, in which the terminal AMP residue of the –CCA sequence is removed and then restored by tRNA nucleotidyltransferase, requires the action of a nuclease for the removal step (77). RNase T was identified as the enzyme that carries out this reaction both in vitro (81) and in vivo (82). In fact, it is the only nuclease in E. coli that can efficiently remove the terminal residue from intact tRNA, due to its ability to digest close to the double-stranded aminoacyl stem (see below). Subsequent studies, to be discussed below, revealed that RNase T participates in many other RNA metabolic processes and that it is required for normal growth.
RNase T has been purified to homogeneity from both normal and overexpressing cells (80, 183). Based on SDS-PAGE, gel filtration, and the sequence of the rnt gene, RNase T is an α2 dimer with a molecular mass of ~50 kDa and a monomer size of 215 amino acids with a molecular mass of 23.5 kDa (80, 120). The dimer form of RNase T is required for activity both in vitro and in vivo (182, 183). RNase T does not associate with any other components in crude extracts (81).
A detailed structural and functional model of RNase T has been developed based on site-directed mutagenesis and its similarity to the known structure of ORNase (354, 355). RNase T was originally found to be extremely sensitive to sulfhydryl reagents and to air oxidation (182, 183). This led to the identification of a cysteine residue at position 168 as an important determinant for dimer formation and RNase activity (183). Subsequent studies revealed that residues within the C-terminal region of RNase T are essential for dimerization and that the monomeric form is inactive (354). These studies also identified the DEDD residues in the catalytic center and confirmed their importance for catalysis. Additionally, three highly conserved sequence segments that form a surface patch rich in positively charged residues were shown to be involved in substrate binding (355). In the model proposed, this surface patch is on the face of the RNase T monomer opposite to that of the DEDD catalytic center. The substrate-binding site and the catalytic center were proposed to come together by dimerization to form the complete active site (355). This was confirmed experimentally by reconstituting active dimers from monomers derived from two inactive mutant proteins, one defective in catalysis and one in substrate binding. The model explains why RNase T must dimerize to function.
Purified RNase T is most active at pH 8 to 9 in the presence of either Mg2+, Mn2+, or Co2+. Enzyme activity is strongly inhibited by elevated ionic strength (>100 mM) (80). The preferred substrate for RNase T is intact tRNA-CCA, from which it removes primarily only the terminal AMP residue. tRNA-CA and tRNA-CCA-CC are also weak substrates. On the other hand, tRNA-CC and tRNA-CU are essentially inactive as substrates (81). Detailed analysis of the base specificity of RNase T using oligonucleotide substrates revealed that it discriminates against pyrimidines, particularly C residues (356). A single 3'-terminal C residue can reduce RNase T action by >100-fold, and two consecutive terminal C residues essentially stop the enzyme. These unusual properties explain why only the single AMP residue is removed from tRNA-CCA. It also explains why certain precursors are substrates for RNase T in vivo and others are not.
RNase T can act on oligonucleotides as small as dimers, although molecules shorter than six residues are poor substrates. On all substrates, RNase T acts distributively, releasing from the substrate after each catalytic cycle (356). RNase T requires a free 3'-hydroxyl group to initiate degradation. Neither tRNA-CCp, a 3'-phosphate-terminated oligoA16, aminoacyl-tRNA, nor a molecule with a dideoxy terminus can serve as substrates. RNase T works actively on single-stranded substrates, but it stops at double strands. However, in contrast to other E. coli exoribonucleases, RNase T can digest right up to the double-stranded region (356). It also has the unusual property of actively digesting single-stranded DNA (321, 352). In fact, the Km values for DNA are much lower than for RNA substrates. This property, coupled with its ability to digest right up to a double strand, has made RNase T an ideal reagent for generating blunt-end DNAs for cloning protocols (352).
RNase T plays an important role in RNA metabolism in E. coli. Thus, it is surprising that the enzyme is restricted to only a small group of bacteria, the γ division of proteobacteria (353). There is no evidence that it normally participates in DNA metabolism, although in high copy it can suppress UV repair defects of a RecJ, Exo I, Exo VII mutant (322). In E. coli, the absence of RNase T leads to a small but significant decrease in doubling time (from 25 to 30 min) in otherwise wild-type cells, and to an even greater decrease in cells lacking RNases I, II D, and BN (from 30 to 45 min) (257). The absence of RNase T also leads to a delay in the recovery of cells from starvation. These data indicate that the other exoribonucleases present in E. coli cannot completely take over the function of RNase T. On the other hand, from among the five exoribonucleases RNase II, D, BN, PH, and T, RNase T is the most effective in restoring growth to an inviable strain lacking all five RNases (153). The presence of just RNase T from among the five results in a doubling time of 35 min compared with 24 min for wild type. This observation indicates that RNase T can substitute significantly for the other missing enzymes.
The explanation for these findings is that RNase T is uniquely required in E. coli for certain 3'-maturation reactions, whereas for other processes, it overlaps in function with additional exoribonucleases. Thus, in the absence of RNase T, neither 5S rRNA (175) nor 23S rRNA (179) is matured to completion (Fig. 2). Both RNAs are assembled into large ribosomal subunits as slightly 3'-extended precursors. Although cells survive and grow in this situation, the mutant ribosomes may be responsible for the decreased growth rates observed in RNase T− strains. In addition to these reactions, RNase T also is a major contributor to the 3' maturation of tRNA (176) and other small, stable RNAs (178). However, for these processes, other RNases can substitute for RNase T (Fig. 1). The unique requirement for RNase T in some processing reactions, but not others, is because only RNase T can approach close to double-stranded regions, which are known to be present near the 3' ends of many RNAs that require exonucleolytic maturation (Fig. 3). When the precursor sequence to be removed is close to a double-stranded stem, such as in 5S or 23S RNA, only RNase T can catalyze the reaction. For tRNA precursors, in which the residues to be removed are >4 from the aminoacyl stem, other nucleases also can participate. Consistent with this notion, RNase T is the only enzyme responsible for the end-turnover of tRNAs, a process that removes nucleotides closer to the aminoacyl stem.
RNase T is encoded by the rnt gene located at about 37.2 min on the E. coli genetic map (40, 283). A typical σ70 promoter is located upstream of the rnt coding region (120). rnt is cotranscribed with a very large open reading frame, termed lhr, which encodes a putative helicase (276). Immediately downstream of lhr is a likely rho-independent transcription terminator, suggesting that rnt and lhr form a bicistronic operon. However, it is not known whether there is any functional relationship between RNase T and Lhr. Likewise, nothing is known regarding regulation of rnt expression.
ORNase.
The third DEDD exoribonuclease in E. coli, oligoribonuclease, was initially identified on the basis of its ability to digest short oligoribonucleotides (241). Its unusual properties distinguished it from RNase II and PNPase, the two E. coli exoribonucleases known at that time (74). Subsequent work, years later, indicated that ORNase activity was not due to any of the newly discovered E. coli exoribonucleases (336), and that, therefore, it is a distinct enzyme.
ORNase has been purified to homogeneity, and, based on studies of the purified protein and the orn gene, it is an α2 dimer with 180-amino-acid subunits of 20.7 kDa (345). The crystal structure of ORNase recently has been resolved to 1.7 Å (T. Fiedler and A. Malhotra, personal communication).
Based on studies of the partially purified enzyme (241), ORNase works best at pH 8 to 9 in the presence of Mn2+; Mg2+ also supports activity but less effectively. The enzyme appears to work processively, and the smaller the oligoribonucleotide, the more rapid is the rate of hydrolysis. Only single-stranded chains are substrates, and a free 3'-hydroxyl group is preferred. However, more complete characterization of ORNase activity, using the homogenous enzyme, needs to be done.
ORNase is encoded by the orn gene located at 94.6 min on the E. coli genetic map (283, 345). Upstream of orn is a reading frame that is transcribed in the opposite direction, and downstream is the glyV operon, suggesting that orn is monocistronic. However, at present, the orn promoter has not been identified, and nothing is known about orn expression.
Interruption of the orn gene suggested that ORNase is essential for E. coli viability (106). This is likely because, at normal cellular levels, none of the other exoribonucleases can degrade oligoribonucleotides. Thus, in the absence of ORNase, cells accumulate oligonucleotides 2 to 5 residues in length derived from pulse-labeled RNA (106). Since neither RNase II nor PNPase can act on small oligonucleotides, it appears that ORNase is necessary to complete the degradation of mRNA to mononucleotides. However, why accumulation of oligonucleotides might lead to loss of viability remains unanswered. As can be gleaned from this short description, much remains to be learned about ORNase.
The single RNase member of the RBN family present in E. coli is RNase BN. The members of this family do not share with other exoribonucleases highly conserved residues typical of metal ion-binding sites (353). Rather, these proteins are hydrophobic with extensive helical structure.
RNase BN.
Studies of the maturation of bacteriophage T4-encoded tRNAs in E. coli suggested that four of the eight tRNAs, those lacking an encoded –CCA sequence, required a host nuclease for 3' processing (291). This nuclease apparently was missing in the mutant strains BN and CAN (198), which resulted in their inability to support the growth of a mutant T4 phage that depended on the synthesis of a suppressor tRNASer (291). The fact that strains BN and CAN contained normal levels of RNase II and RNase D, and that mutant strains lacking these RNases could still support growth of the mutant T4 phage (282), indicated that another nuclease was involved. This enzyme termed RNase BN was subsequently identified and shown to be absent in strains BN and CAN (12, 13).
RNase BN has been purified to near homogeneity (33). The purified enzyme has a native molecular mass of ~65 kDa; and, based on SDS-PAGE of the monomers and the derived sequence of the putative rbn gene (32, 33), it is an α2-dimer. A dimer form for the native enzyme is also supported by the finding that an interrupted rbn gene carried on plasmids displays a dominant-negative phenotype (32). Purified RNase BN is extremely sensitive to sulfhydryl reagents (33).
The catalytic properties of RNase BN are quite unusual (33). The enzyme is most active at pH 6.5 in the presence of Co2+ and 200 to 400 mM monovalent cation; Mg2+ is considerably less effective. RNase BN also has an extremely narrow substrate specificity. It is active on tRNA substrates in which residues within the –CCA sequence have been altered, but other closely related molecules are essentially inactive as substrates. Thus, while tRNA-CU and tRNA-CA are efficient substrates, intact tRNA-CCA, tRNA-CC, tRNA-CCA-Cn, diesterase-treated tRNA, rRNA, and poly(A) are poor or inactive substrates. RNase BN also cannot act on RNA or DNA oligonucleotides, ranging in length from 11 to 17 residues. Inasmuch as RNase BN’s action in vivo is on phage T4 tRNAs, which do not contain a -CCA terminus, its substrate specificity fits with such a role. RNase BN releases the terminal residue from either tRNA-CU or tRNA-CA as a mononucleotide, and it appears to act in a distributive manner.
The physiological role of RNase BN is unclear. Mutant cells with an interrupted rbn gene grow normally on YT media at 31°C to 42°C (32). However, such mutant cells do not support the growth of the bacteriophage T4 mutant that requires synthesis of the suppressor tRNASer. RNase BN can contribute to the maturation of cellular tRNAs and to growth when RNases II, D, T, and PH are absent; but it does so extremely poorly (Fig. 1). Cells with only RNase BN from among these five exoribonucleases grow with a ~100-min doubling time (153) and synthesize a suppressor tRNATyr at only ~10% the wild-type level (275). Given these data, it is unlikely that maturation of cellular tRNA is a major function of RNase BN, although it remains possible that a specific tRNA might be a substrate. The high degree of specificity of RNase BN for tRNAs with an altered –CCA sequence also makes its role in tRNA metabolism questionable, as such altered tRNAs are not known to arise in cells. So, although RNase BN can act on tRNA molecules under certain conditions (i.e., phage infection or when multiple other RNases are absent), it is likely that its normal physiological role lies elsewhere.
A gene, now termed rbn, localized at 87.8 min on the E. coli genetic map, has been suggested to encode RNase BN (32, 283), but strictly speaking, this has not yet been proven conclusively. Thus, cloning of the putative rbn gene on a multicopy plasmid leads to overexpression of RNase BN, and its interruption results in loss of activity. On the other hand, sequence information has not been obtained from the purified protein to confirm that it is encoded by rbn. Also, the size of the RNase BN monomer on SDS-PAGE (~37 kDa) differs somewhat from that deduced from the rbn coding region (32.8 kDa). Moreover, the mutations present in strains BN and CAN that lead to loss of RNase BN activity are not in the coding region of rbn or in the immediate flanking regions. rbn is the second gene of a four-gene operon. The mutations that affect RNase BN activity in the mutant strains are located within a cloned ~4-kb fragment that includes the entire operon (32), but their exact location has not yet been determined. Clearly, further work will be required to prove that rbn actually encodes RNase BN.
In addition to the six hydrolytic exoribonucleases, E. coli contains two phosphorolytic nucleases, PNPase and RNase PH, members of the PDX family. Enzymes of this family use Pi instead of H2O as the nucleophile and generate nucleoside diphosphates, rather than monophosphates, as the product. PNPase from E. coli, as well as from other organisms, is a large, multidomain protein, whereas RNase PH members generally are small, single-domain proteins (353). PNPase polypeptides generally contain several highly conserved domains; the central PDX domain is common to all members of the PDX family and is the part of the protein homologous to RNase PH (220, 353). The three characteristic motifs of this domain include the Pi-binding region and the catalytic center. Structually, this region of PNPase and RNase PH are almost identical (123).
PNPase.
Although studies of E. coli PNPase date back to the 1950s and essentially homogenous preparations of the enzyme were made as early as the 1960s (early information on PNPase can be found in reference 190), PNPase is still under active investigation. Recent studies of its structure, physiological role, and regulation continue to provide fascinating information.
E. coli PNPase has been purified as an α3-trimer (264) and also as an α3β2-structure (265), now known to contain two subunits of the glycolytic enzyme enolase (269). A portion of PNPase also purifies with the degradosome, a multienzyme complex that contains an endoribonuclease, RNase E, as well as an RNA helicase, RhlB, and enolase (38, 221). As noted, PNPase is a large protein. Based on the sequence of the pnp gene, the PNPase polypeptide is 711 amino acids in length with a calculated molecular mass of 77.1 kDa (274). Although the crystal structure of E. coli PNPase has not yet been determined, that of the closely related enzyme from Streptomyces antibioticus has been solved (309). The enzyme has two core domains closely related to RNase PH that pack closely together in each of the three subunits, resulting in a core hexameric ring structure containing a central channel. Three additional domains from each subunit are located near the upper and lower surfaces of the core. The two upper domains are homologous to KH and S1 RNA-binding domains, whereas the lower domain of unknown function is all α-helical. Based on the location of tungstate, a phosphate analog, in the crystal structure, the second of the two core domains is predicted to contain the PNPase active site. How RNA binds to PNPase and accesses the active site is not yet understood.
PNPase catalyzes three reactions in vitro, the synthesis of RNA from nucleoside diphosphates, an exchange reaction between Pi and nucleoside diphosphates, and the phosphorolytic degradation of RNA to form nucleoside diphosphates (190). Although in vivo the primary role of PNPase is RNA degradation (227), some situations have been found in which PNPase can also act in the synthetic mode (225, 277). All the reactions of PNPase require the presence of a divalent cation; Mg2+ is most effective, but Mn2+ can also function. Early studies suggested that the active site of PNPase contained several subsites, whose roles varied depending on the reaction catalyzed (190). In addition, it was predicted that PNPase binds long RNAs at sites more than 20 nt away from its site of action and that this accounts for its processivity. Deletion of the KH or S1 RNA-binding domains does not abolish catalytic activity, although some activities are partially decreased (132). Whether the enzyme remains processive was not examined in this study.
In the physiologically important degradative mode, PNPase can rapidly and processively digest RNA chains (190, 216). However, the enzyme is markedly inhibited by RNA helical structure, resulting in stalling and dissociation of six to nine residues from the base of stable stem-loop structures (298). Nevertheless, in vivo, PNPase is an important RNA degradative enzyme (45, 56, 88, 227). Thus, the presence of an RNA helicase in the degradosome (221, 269), as well as repeated cycling on polyadenylated tails (55, 331), enables PNPase to eventually proceed through secondary structure in many RNAs. As the length of an RNA substrate shortens, PNPase action becomes more distributive, and ultimately the enzyme can no longer remain bound (190). The small oligonucleotides resistant to PNPase and RNase II action are degraded in vivo by ORNase (106).
E. coli cells devoid of PNPase retain viability, although their growth rate slows somewhat (217). However, such cells become increasingly sensitive to antibiotics. The basis of this phenotype is not clear. PNPase is known to be induced upon cold shock (138), and it is required to repress the production of other cold-shock proteins at the end of the acclimation phase (332). In the absence of PNPase, mRNAs for cold-shock proteins are not degraded, and cells are unable to resume growth after the initial cold-shock response. PNPase also plays a role in quality control of tRNA (181). In the absence of PNPase, cells accumulate elevated levels of a temperature-sensitive tRNATrp that normally is degraded. Similar results were also found with a temperature-sensitive tRNASer (1). These data indicate that PNPase is specifically required for the essential degradation of certain defective RNA molecules and that in its absence the RNAs accumulate. In some of these examples, the PNPase defect is accentuated in pcnB mutants, which eliminate poly(A) polymerase (181, 332). These findings support the conclusion that polyadenylation serves to facilitate the action of PNPase, presumably by providing an optimal binding site for initiating the degradative process (226).
Even more dramatic effects of the lack of PNPase become evident when it is coupled with the absence of a second exoribonuclease. Thus, cells devoid of both PNPase and RNase II are inviable, and they accumulate large fragments of mRNA (88), indicating that both enzymes contribute to mRNA degradation. While this apparent functional overlap occurs, more recent information indicates that it is probably true only for a subset of mRNAs, and that for many messages, PNPase and RNase II serve different functions (227). In a second example, cells lacking both PNPase and RNase R also are inviable (46). These cells accumulate large amounts of rRNA fragments that are thought to arise due to errors in transcription or ribosome assembly (45). Finally, cells lacking PNPase and RNase PH, while viable at 37°C, are extremely cold sensitive at 31°C and are unable to assemble 50S ribosomes at this temperature (346). In this group of examples, PNPase serves as one of a pair of enzymes able to carry out a specific, essential function. The presence of either RNase is sufficient to support almost normal growth, indicating a high degree of functional overlap in the specific process. However, when both exoribonucleases are absent, none of the other RNases present can take over the missing function.
PNPase is encoded by the pnp gene at 71.3 min on the E. coli genetic map (283). pnp lies downstream of the rpsO gene, encoding ribosomal protein S15, as part of a two-gene operon (283). pnp is expressed from either of two promoters, one upstream of rpsO, generating a dimeric transcript, and one just upstream of pnp (94). Expression of PNPase is autoregulated at the posttranscriptional level by several mechanisms. In one case, the endoribonuclease RNase III cleaves the pnp mRNA leader in a stem-loop region that destabilizes the message by creating a 3' end at which PNPase binds and initiates degradation (133). Mutations that eliminate PNPase activity or that remove the KH and S1 RNA-binding domains abolish autocontrol (132). Likewise, mutations that inactivate RNase III elevate PNPase expression (266). The elevation of PNPase expression at low temperatures also is related to autocontrol (205). At the low temperatures, PNPase is less efficient at degrading its own message, resulting in increased pnp expression. In a second type of control, PNPase levels are affected by the amounts of polyadenylated transcripts. Conditions that lead to elevated polyadenylation of E. coli transcripts stabilize the pnp message and result in elevation of PNPase (226). It has also been reported that PNPase expression can be down-regulated by increased levels of RNase II present in cells (349). While the mechanism of this regulation has not been elucidated, RNase II may also degrade the pnp message after RNase III cleavage.
RNase PH.
The second Pi-dependent exoribonuclease in E. coli, RNase PH, was identified when extracts of a cell lacking RNase II, D, BN, and T were found to mature the 3' terminus of a tRNA precursor in a reaction dependent on Pi (68). Further examination revealed that the processing reaction was not due to PNPase, the only phosphorolytic nuclease known at that time (83). Subsequent purification of RNase PH to homogeneity from both a normal (251) and an overexpressing strain (135) and characterization of its properties (154) confirmed that RNase PH is a distinct Pi-dependent exoribonuclease. RNase PH is now known to be the founding member of a large family of nucleases widely distributed in all kingdoms (353).
E. coli RNase PH was found to be structurally somewhat unusual. Sequence analysis of the rph gene indicated that the polypeptide is 238 amino acids in length with a molecular mass of 25.5 kDa (252, 267). Yet, it migrates on SDS-PAGE as a 30- to 33-kDa molecule and often shows multiple bands (135, 154). Moreover, the protein tends to aggregate. The smallest active form on gel filtration appears to be the dimer, but aggregates close to 200 kDa are also observed (154). The crystal structure of the E. coli enzyme has not yet been determined, but recent structures of the Aquifex aeolicus (123) and the Pseudomonas aeruginosa (47) enzymes indicate that RNase PH forms a hexameric ring structure as a trimer of dimers. This structure is very similar to that reported for the core of PNPase (309) and supports the suggestion that PNPase evolved by duplication of an RNase PH-type enzyme (309). Mutational analysis has shown that RNase PH dimers are inactive and that the hexameric ring is essential for activity of the P. aeruginosa enzyme (47). The data suggest that all members of the PDX family may function as oligomeric rings (309). These findings also explain the tendency of E. coli RNase PH to aggregate, and they suggest that the active dimers observed by gel filtration probably oligomerized in the presence of the RNA substrate during assay (154).
RNase PH is active as a degradative enzyme only in the presence of Pi (154). Like PNPase, it can also act as a polymerizing enzyme, adding any nucleoside diphosphate to the 3' terminus of a variety of RNAs (251). As a degradative enzyme, RNase PH is most active at pH 8 to 9 in the presence of Mg2+ at 10 mM Pi; Mn2+ and Co2+ also function to some degree (154). RNase PH acts on homopolymers and on tRNA-type substrates. With the latter, the preferred substrate is one containing a few residues following the –CCA sequence from which it can generate mature functional tRNA-CCA. tRNA-CCA and tRNA-CC are considerably poorer as substrates. The Km value for tRNA-CCA is ~10-fold higher than that for tRNA-CCA-C2-3, suggesting that the mature tRNA would tend to dissociate from the enzyme after processing. RNase PH requires a free 3'-hydroxyl group to act on tRNA precursors (154).
The recent X-ray structures have identified the catalytic site of RNase PH (47, 123) and have identified four residues important for binding Pi, which are conserved among bacterial RNase PHs. This Pi-binding site superimposes well on the second core domain of PNPase, which was predicted to be the catalytic center of that enzyme (309). The Pi-binding site in RNase PH is present at the bottom of a cleft, and this structure would only allow the 3'-single-stranded region of a tRNA precursor to enter easily. This would explain the preferred activity on precursor tRNAs compared with the mature forms. In addition, the structural analysis indicates that the interface between dimers is important for RNA substrate binding and that regions from the adjacent dimer would interact with the double-stranded aminoacyl stem of tRNA (47), explaining the need for the oligomeric structure. The importance of adjacent subunits for RNA binding has also been proposed for RNase T action (355).
Although RNase PH is not an essential enzyme in E. coli (null mutants grow normally and synthesize mature tRNA normally), when rph mutants are coupled with mutations in a variety of other exoribonucleases, cells grow more slowly and synthesize tRNA poorly (155). This phenotype varies dramatically with the additional nuclease missing. In particular, RNase T– RNase PH– double-mutant cells display a major growth defect. These two RNases are the major contributors to the 3' maturation of tRNA molecules (Fig. 1), and in their absence large amounts of tRNA precursors accumulate (176). Even when RNase PH is absent by itself, certain tRNA precursors are incompletely processed, indicating that the presence of all the other RNases is insufficient to carry out the required processing reactions. Quantitatively, only 30% as much suppressor tRNATyr is made in the RNase T- RNase PH– double-mutant strain as in wild type, and this level decreases even further upon removal of additional exoribonucleases (275). After RNase T, RNase PH is most effective in supporting growth of cells lacking multiple exoribonucleases (153). Consequently, RNase PH can take over the functions of the missing RNases quite well. In addition to tRNA, RNase PH also participates in the 3' maturation of a number of other small, stable RNAs in E. coli (178). For most RNA substrates, both RNase T and RNase PH serve as the processing enzymes, indicating that both a hydrolytic and a phospholytic nuclease are effective. On the other hand, as discussed earlier, only RNase T can complete 3' maturation of 5S and 23S rRNAs (Fig. 2). These findings suggest that RNase PH cannot approach as close to a double-stranded stem as RNase T can.
A second pair of mutations that lead to a dramatic phenotype is one that eliminates both phosphorolytic exoribonucleases, RNase PH and PNPase (346). As noted earlier, these cells are extremely cold sensitive, are defective in 50S subunit assembly, and degrade 23S rRNA. Even ribosomes made at 42°C (a temperature at which cells grow almost normally) are defective, because they are unable to function upon a temperature shift to 31°C. These findings demonstrate that some essential process in ribosome metabolism can be carried out only by the phosphorolytic nucleases, and they raise the possibility of a connection between Pi levels and ribosome metabolism mediated through the action of these two nucleases.
RNase PH is encoded by the rph gene located at 82.2 min on the E. coli genetic map, and it is part of an operon together with the downstream pyrE gene, which encodes orotate phosphoribosyltransferase, an enzyme of pyrimidine biosynthesis (252, 268, 283). The two genes are coupled translationally, such that translation of rph is necessary to transcribe past the pyrE attenuator (267). Moreover, the rph message differs depending on pyrimidine metabolism (8). When the cellular UTP pool is low, a dicistronic rph-pyrE message is made; however, when UTP levels are high, a monocistronic rph message is formed due to attenuation of pyrE expression. Normally, the dicistronic message is cleaved to generate two monocistronic messages. The significance of the complex relationship between rph and pyrE is presently not understood.
In many widely used E. coli strains, such as W3110 and MG1655, a GC base pair is missing near the 3' end of the rph gene (134). This results in a frameshift that reduces the size of RNase PH by 10 amino acids and essentially eliminates RNase PH activity in these strains. Also, as a consequence of the altered translation of rph, orotate phosphoribosyltransferase is decreased and the cells become starved for pyrimidines in minimal medium. The consequences of the mutation for RNA metabolism in these strains and their derivatives have not been examined.
RNase I.
RNase I was the first E. coli endoribonuclease to be identified. A description of this work and other early studies of the enzyme are presented in a detailed review by Shen and Schlessinger (293), which will be used as the source for many of the older references. RNase I is unusual among the well-characterized E. coli RNases in that it is the only one active in the absence of a divalent cation and is the only one to generate 3'-phosphoryl-terminated RNA products. RNase I also differs from the other RNases with regard to its subcellular localization. Thus, while it is often found bound to 30S ribosome subunits in extracts, as much as 90% of the enzyme is released upon preparation of spheroplasts, indicating that most of it actually resides in the periplasmic space in vivo (293, 348). However, forms of RNase I are also found in the debris fraction (10) and within the cell (34, 348).
Based on the sequence of the rna gene encoding RNase I (219, 347), it is a member of the T2 family of RNases (122), which are widely distributed among prokaryotes and eukaryotes. An enzyme similar to E. coli RNase I also has been characterized from Salmonella enterica serovar Typhimurium (41). Many endoribonucleases with properties similar to RNase I, termed RNase IV (297), RNase F (111), RNase I* (34), RNase M (33a), and RNase R (302) have been reported to be present in E. coli. However, examination of the E. coli genome reveals no other genes related to rna (307). Thus, these enzymes either are forms of RNase I with slightly altered properties or they are derived from genes unrelated to rna despite the similarity of the proteins. It is already known that RNase I* is derived from the rna gene (34) and that the protein purified as RNase M (33a) is actually a mutant form of RNase I (307). It has been suggested (156) that the original properties attributed to RNase M (33a, 218) are due to a different protein, but this needs to be verified. Clearly, further work on the relationship among all of these activities is warranted.
Based on the sequence of the rna gene (219), RNase I is a protein of 245 amino acids and a calculated molecular mass of 27.2 kDa. The protein fractionates and apparently functions as a monomer (81). As judged by activity assays (293) and physical measurements (258), RNase I is a very stable protein. The protein contains eight cysteine residues (219), and their oxidation state may have an important influence on the properties of RNase I activity. For example, RNase I*, which is found in spheroplasts and is at least partially in the reduced state, differs in several respects from the completely oxidized form of RNase I present in the periplasm (34). The possibility of multiple alternate structures, arising from the formation of different disulfide bonds, might account for the various proteins apparently related to RNase I.
RNase I degrades all types of RNA molecules with no obvious specificity for the phosphodiester bond (293). The initial cleavage generates a cycle 2',3'-nucleotide, which ultimately is converted to a 3'-phosphate. Limit digests consist primarily of 3'-mononucleotides. Divalent cations are not needed, and, in fact, Mg2+ is somewhat inhibitory.
The physiological role of RNase I is unknown. Mutant E. coli (103, 348) or S. enterica (328) strains grow normally and show no major metabolic defects. Under certain stress conditions and conditions that damage the cell membrane, periplasmic RNase I may enter the cell, leading to extensive RNA degradation (reviewed in reference 78). However, it is unlikely that this is the primary role of RNase I, because, in effect, its entry into the cell leads to suicide. Alternatively, it is possible that the smaller, intracellular pool of the enzyme may serve a degradative function under certain conditions.
The rna gene encoding RNase I is located at 13.9 min on the E. coli genetic map (283). It has an unusual promoter with a poor –35 region that is located within a stem-loop structure that likely is a transcription-termination site for an upstream gene (348). The gene encodes a 23-amino-acid leader peptide that probably serves as the signal peptide for transport of the protein to the periplasmic space (219).
RNase III.
Excellent reviews dealing with RNase III have been published (60, 239). Originally identified in 1968, RNase III is the only endoribonuclease in E. coli that specifically hydrolyzes double-stranded RNA (dsRNA) (279). Moreover, it is widely distributed in almost all bacteria (58). RNase III has been purified to homogeneity by affinity chromatography based on its dsRNA-binding activity and shown by gel filtration to be active as a 50-kDa homodimer (91). Based on the sequence of the rnc gene, which encodes RNase III, the monomer is a polypeptide of 226 amino acids (200, 235) with a calculated molecular mass of 25.6 kDa.
The C-terminal one-third of RNase III is a dsRNA-binding domain (dsRBD) that is conserved in a variety of proteins able to bind dsRNA (157, 303, 314). The isolated dsRBD can bind dsRNA in vitro, but it is unable to cleave the substrate (238). On the other hand, a shortened form of RNase III, lacking the dsRBD, has activity similar to intact RNase III under certain in vitro conditions. These data indicate that the N-terminal endonuclease domain of RNase III can function in the absence of the dsRBD (308). Nevertheless, the dsRBD does play a role in determining substrate specificity and cleavage site location, based on an analysis of RNase III hybrid proteins (59). The N-terminal domain of RNase III possesses a "signature box"of 11 conserved amino acids (220, Prosite PS00517) as well as additional, highly conserved Asp and Glu residues (281). The involvement of some of these residues in the catalytic action of RNase III was proven by a combination of modification of the carboxylate groups (239) and mutational and structural analyses. For example, an E117K mutant of RNase III is inactive (63). It can bind, but it cannot cleave the RNA substrate (172).
The structure of the dsRBD of E. coli RNase III has been solved (157). It is a compact ellipsoid with an αβββα tertiary fold. The same fold also has been found in a number of other proteins that bind dsRNA (60, 239). Analysis of such a structure from the Xenopus laevis RNA-binding protein A complexed with dsRNA revealed that the dsRBD makes contact with both RNA strands on the 2'-hydroxyl groups within a 16-bp region. These interactions would explain why RNase III selectively recognizes dsRNA and why it displays an apparent lack of sequence specificity (285).
More recently, the crystal structure of the N-terminal endonuclease domain of the A. aeolicus RNase III (Aa-RNase III) was solved, both in a form free of ligand as well as in a complex with Mn2+ (19). This structure displays a novel protein fold containing seven α-helices and a 310-helix, but it has no β-strands. Dimerization of the monomers creates a valley containing two compound active centers, which accommodate the dsRNA substrate. Mn2+ binding facilitates the formation of two potential RNA-cutting sites within each compound active center. Six negatively charged residues, E40/D44/D107/E110 and E37/E64, are tightly placed in the two sites. Some of these amino acids are known to be important for catalysis based on mutational analysis of the corresponding residues in the E. coli protein (19, 73, 172). Modeling of the intact RNase III structure bound to dsRNA suggests a mechanism of substrate cleavage in which four cuts are made by each dimer enzyme.
Cleavage on both strands of dsRNA by RNase III creates fragments with 5'-phosphate and 3'-OH groups and containing 2-nt 3' overhangs (92, 237). The action of RNase III requires the divalent cation Mg2+, but it can be substituted by Mn2+, Co2+, or Ni2+ (238). RNase III recognizes complicated features of its RNA substrates. A common structural element required by RNase III for cleavage is a dsRNA of 15 to 20 bp (63, 278). The minimum length that confers reactivity is a 12-bp RNA hairpin, slightly greater than one α-helical turn, with a single-stranded 5' extension (239). However, the site of cleavage is selected according to specific RNA sequence and structural elements (reactivity epitopes) and not solely by the length of helix (42, 343). Sequence alignment of RNase III substrates indicates that specific Watson-Crick base pair sequences at defined positions relative to the cleavage site act as antideterminants (343). Thus, as shown by mutation, the presence of certain base pair sequences in a 3-bp "proximal box" and a 2-bp "distal box" in the helix are inhibitory to RNase III cleavage.
Some E. coli RNase III substrates contain an internal loop within the double-stranded region, which directs cleavage to one of the single strands within the loop (239). Mutational analysis of such a substrate with an asymmetric internal loop revealed that the asymmetry per se determines the cleavage of only one strand and that the sequence of the internal loop is not the determinant for reactivity (30). A model supported by most of the results is that the cleavage site is determined by the site of RNase III binding on the helix, which in turn is dictated by the base pair sequence. In addition, an internal loop appropriately positioned within the helix can force a cleavage on only one of the two strands (239).
RNase III participates in cellular RNA metabolism by cleaving dsRNA substrates. Its primary role is in the maturation of rRNA. RNase III cleaves in the double-stranded regions bracketing the 16S and 23S rRNAs in the precursor to convert the primary transcript into shorter intermediates (22, 295, 335; Fig. 2). Other activities, some of which are still unidentified, complete the processing of the intermediates (104, 117, 175, 179, 180, 327; Fig. 2). The processing reactions are carried out preferentially on preribosomal particles (4, 300, 301), and RNase III cannot be replaced by any other processing activities. Thus, inactivation of RNase III results in aberrant processing and accumulation of incompletely processed, yet functional, 23S RNA species (158). In S. enterica, RNase III has been shown to be responsible for removal of intervening sequences in the 23S rRNA (26). However, this activity is dispensable for 23S rRNA function as well as for cell growth (209).
RNase III also participates in the maturation or decay of a number of cellular and phage mRNAs, of some tRNAs, and of transcripts from plasmids (60, 63, 239, 312). Although global mRNA decay is not affected by a mutation in the rnc gene, degradation of a few mRNAs is initiated by RNase III cleavages, indicating that RNase III does play a limited role in mRNA degradation (109, 273, 330). Also, RNase III action on some tRNA-mRNA and mRNA-mRNA cotranscripts is required for their maturation (239, 272). For example, translation of E. coli adhE mRNA and at least two mRNAs derived from phage T7 are activated by RNase III cleavages in their 5'-UTR, which remove sequences blocking the ribosome binding site (11, 63, 237).
RNase III is encoded by the rnc gene located at 58.2 min on the E. coli genetic map (283). It is the first gene in an operon that contains era and recO downstream (310). Transcription initiates from a promoter 170 nt upstream of rnc (15). Although RNase III serves many important functions in E. coli, mutants lacking the enzyme remain viable; however, their growth is impeded (310). Likewise, overexpression of RNase III does not have a deleterious effect on cells (43, 171, 201).
The activity of RNase III is controlled at multiple levels. RNase III can cleave its own mRNA in its 5'-UTR at a portable, double-stranded region, termed rncO, to autoregulate expression of the genes in the operon (15, 206, 207, 208, 273). A very similar, if not identical, autoregulation mechanism is present in S. enterica (9). At the protein level, RNase III undergoes phosphorylation following T7 infection, a process that stimulates its activity fourfold (210, 239). RNase III may associate with ribosomes in vivo. It copurifies with ribosomes (4, 279) and associates with high-molecular-weight components in cell extracts (131). Moreover, a ribosomal protein gene mutation can be suppressed by an rnc mutation (234). Interaction of RNase III with the ribosome has been shown to affect the specificity of RNase III (4). It has been suggested that association with ribosomes may serve to locally enrich the amount of this less abundant enzyme (239).
RNase P.
E. coli RNase P has been the subject of several recent reviews (7, 52, 99, 159, 165). The enzyme was originally discovered during a search for tRNA maturation enzymes in E. coli extracts (280). Subsequent studies of RNase P revealed the surprising finding that the enzyme contains an essential RNA component, termed M1 RNA (305), and that the RNA is the catalytic subunit (110). A catalytic RNA subunit is now known to be a common feature of bacterial RNase Ps. The E. coli RNA is 377 nucleotides in length with a mass of ~130 kDa. The protein component of RNase P, termed C5, is a basic polypeptide of 119 amino acids with a molecular mass of 13.8 kDa. Both M1 RNA and C5 protein have been purified to homogeneity (162), and the two components can be combined to reconstitute the holoenzyme in vitro. RNase P action generates products with 5'-phosphate and 3'-hydroxyl termini in a reaction that utilizes Mg2+, but other cations also can function. Remarkably, under certain conditions M1 RNA can digest tRNA precursors in the absence of C5 protein (110). However, the presence of C5 protein greatly enhances cleavage under physiological conditions, consistent with the finding that C5 is required for RNase P action in vivo (5, 256).
Extensive analysis of RNase P has led to the generation of low-resolution structures of the RNA and protein moieties; however, high-resolution structures are not yet available (52). Our current information about the structure of RNase P RNA and its interaction with tRNA is based on phylogenetic analysis, chemical crosslinking, nucleotide analog interference mapping (NAIM), and mutational studies (165). The current model suggests that M1 RNA folds into a number of helices that form two functional regions: a specificity domain (S domain, containing helices 7 to 14) and a catalytic domain (C domain, the rest of the RNA) (52, 99, 165, 204a, 255a).
Considerable conservation of RNA sequence is found among bacterial RNase Ps, primarily in the C domain (112, 165). Modeling studies have resulted in a putative three-dimensional structure for the RNA subunit (329), and for the holoenzyme (316). Recently, the crystal structure of the specificity domain of Bacillus subtilis RNase P was solved, and it agrees well with the other analyses (164). Mutational and crosslinking studies have shown that the small C5 protein interacts with only part of the M1 RNA, near the catalytic site (18, 108, 292, 311, 320). The interaction involves several regions of both M1 RNA and C5 protein. High-resolution structures of the protein subunit of RNase P from B. subtilis by X ray (304) and from Staphylococcus aureus by NMR (299) have been solved. These structures revealed three potential RNA-binding motifs, a metal-binding loop, and a conserved cleft, which appear to be involved in the interactions with the pre-tRNA substrate and with the RNase P RNA.
RNase P cleaves 5'-leader sequences from precursors of all tRNAs, generating their mature 5' termini. Recognition of the tRNA substrate by RNase P has been studied extensively (reviewed in references 7, 52, and 159). Since structure is so highly conserved among tRNAs, it was thought likely that specific structural domains within tRNA precursors would be recognized by RNase P. In fact, it is now known that RNase P recognizes the coaxially stacked acceptor stem and the T-stem-loop of tRNA, as well as the conserved 3'-CCA (165). It is thought that interaction of the S domain of RNase P RNA with the T-stem-loop of tRNA is needed for efficient cleavage. This conclusion is supported by the observation that the S domain of the B. subtilis enzyme possesses a substrate-interacting clamp in its three-dimensional structure (164), which by itself can bind the pre-tRNA substrate with micromolar affinity (270). The C domain of the catalytic RNA interacts with substrate residues at and near the RNase P cleavage site (52). In the presence of its protein component, the isolated C domain of B. subtilis RNase P RNA retains significant pre-tRNA processing activity, indicating that the C domain contains the active site (191). In the C domain of E. coli M1 RNA, a GGU sequence in the P15 loop interacts with the 3'-RCCA sequences of the pre-tRNA (159), and it was shown that this interaction between the R and U residues is important for cleavage-site recognition, cleavage efficiency, and Mg2+ coordination in this region (23, 159). Other residues and secondary structural regions in both the S and C domains that participate in substrate recognition and catalysis are discussed in detail in recent reviews (52, 165).
Addition of the C5 protein decreases the catalytic requirement of RNase P for metal ions (110), possibly through promotion of specific interactions with the substrate, as suggested by studies using the B. subtilis enzyme (64, 166, 240). As a result, RNase P recognizes only part of the pre-tRNA substrate. Such a minimal substrate can be constructed by combining the T-stem-loop and an acceptor stem with a 3'-CCA (211). Bipartite molecules with nicked T loops, which preserve the minimal structure, also can be cleaved by RNase P (98a). These studies led to the identification of an "external guiding sequence (EGS)" that directs cleavage of a target RNA by RNase P (6).
Divalent metal ions, such as Mg2+, and the metal-binding sites on M1 RNA play important roles in the structure of the RNA and in substrate cleavage. However, because of the numerous metal ions that potentially could interact with RNase P RNA, the positions of the functionally important metal ions are poorly understood (52, 159). Based on NAIM and kinetic studies, residues A62 to C70, A130 and A132, and the P15 loop all have been suggested to interact with functionally important metal ions (159). Other studies indicate that a cluster of metal ion interactions in the P1–P4 multihelix junction define the catalytic core of RNase P (51). Additional work suggests that two to three Mg2+ are required for substrate binding and catalysis, possibly through an SN2 in-line attack by a solvent nucleophile on the scissile phosphodiester bond, resulting in a trigonal bipyrimidal transition state (165).
The major function of RNase P is maturation of the 5' ends of tRNA (Fig. 1). Inasmuch as tRNAs are made as precursors containing 5'-leader sequences, RNase P cleavage is essential to generate the complement of functional tRNA molecules. No other enzyme can substitute for RNase P in tRNA maturation. Therefore, RNase P is essential for cell growth, and temperature-sensitive mutants of either M1 RNA or C5 protein accumulate high levels of tRNA precursors (286, 287, 288). In addition to tRNA, RNase P is responsible for maturation of a number of other RNAs. These include the 5' ends of 4.5S RNA (260), tmRNA (163), and C4 antisense RNA of bacteriophages P1 and P7 (114); maturation of the 3' end of tRNA-leuX (242); and processing of a polycistronic mRNA (3). A global survey of RNase P function demonstrated a number of RNase P cleavage sites on polycistronic mRNAs (174) that affect expression of downstream regions of the mRNAs. Analysis of the operons affected suggests that RNase P participates in transport and amino acid biosynthesis, as well as other processes.
M1 RNA is encoded by the rnpB gene located at 70.4 min of the E. coli genetic map (283). The RNA is transcribed from multiple promoters, producing precursors with extra nucleotides at their 3' ends (192). Maturation of M1 RNA precursors is initiated by RNase E cleavage 1 or 2 nucleotides downstream of the mature 3' end, followed by exonucleolytic trimming (178, 192). Strongest transcription is from the +1 position of M1 RNA under the control of the closest promoter (228a, 285a). Transcription from distal start sites results in transcripts having various lengths of 5' extra sequences, the longest of which has four additional openreading frames upstream of the transcript of rnpB (192, 228a). Maturation of the 5' endrequires processing by an as yet unknown activity. The rnpA gene encoding the C5 protein maps at 83.7 min (283). It is cotranscribed with the upstream rpmH gene encoding the ribosomal protein L34 (112a).
RNase P is under control of the relA locus and ppGpp (7, 86, 139, 259). A consensus discriminator sequence responsive to stringent conditions is present in the promoter of the rnpB gene. Mutation of this sequence (139), or of the flanking region (259), alters the response of M1 RNA production to stringent conditions. This result contrasts with the finding that M1 RNA levels increase as a function of growth rate but are independent of ppGpp or FIS (Factor for Inversion Stimulation [86]).
Seven well-characterized endoribonucleases are known in E. coli. These are RNases I, III, P, E, G, HI, and HII (Table 2). In addition, two toxins, RelE and MazF, were recently shown to cleave mRNAs in a highly specific manner (262, 344). Homologues to most of these enzymes are also present in Salmonella (58; Z. Li, unpublished result). Most of the endoribonucleases cleave RNA in the presence of divalent cations, producing fragments with 3'-hydroxyl and 5'-phosphate termini. RNase I, on the other hand, acts by a different mechanism, generating products with 3'-phosphates and does not require a divalent cation for activity. Some of the RNases are highly specific with regard to their substrates and to their cleavage sites, whereas others cleave RNA relatively indiscriminately, resulting in extensive degradation.
Table 2E. coli endoribonucleases |
Members of the RNase E/G endoribonuclease family contain an S1 RNA-binding domain. RNase G is a smaller protein, highly homologous to the catalytic N-terminal half of RNase E (58, 141, 213), whereas RNase E contains additional domains located in its C-terminal region. RNase E and G were both first identified in E. coli (102, 180, 327), but RNase E, RNase G, and RNase E/G-like proteins are now known to be present in almost all subdivisions of bacteria. Some, such as the α-, β-, and γ-divisions of proteobacteria contain both RNase E and G (58, 141; National Center for Biotechnology Information [NCBI] COG1530), whereas others have only one family member.
RNase E.
RNase E is an interesting multifunctional enzyme (57). Full-length RNase E polypeptide consists of 1,061 amino acids (39). With a calculated size of 118 kDa; however, the protein migrates on SDS-PAGE with an apparent molecular mass of 180 kDa. RNase E contains multiple domains. The N-terminal half of the protein (N-Rne), which is required for cell viability, confers the RNase activity and contains the S1 RNA-binding domain (27, 212, 313). The aromatic residues Phe-57, Phe-67, and Lys-112 in the S1 RNA-binding domain have been shown to be important for RNase E activity (85). Recent studies showed that the N-terminal 62 kDa organizes into a catalytically active tetramer in solution (31). The C-terminal half contains an arginine-rich region with RNA-binding activity (ARRBS; 39, 53, 140, 212). However, this region is not required for endonucleolytic cleavage (212). Also present in the C-terminal half of the protein are three proline-rich regions, which may be responsible for its slow migration on SDS-PAGE (39, 53, 212).
RNase E is the central component of the degradosome, a membrane-associated complex containing multiple proteins involved in RNA degradation (38, 189, 319; recently reviewed in reference 37). The "minimal" active RNA degradosome, which can be reconstituted in vitro, includes RNase E, PNPase, and the DEAD-box RNA helicase, RhlB (38, 55, 56, 269). Enolase, usually a glycolytic enzyme, also is an integral component (269). The C-terminal domain of RNase E serves as the scaffold for assembly of the degradosome complex (319).
RNase E binds RNA (61) and cleaves at single-stranded, AU-rich sequences (93, 188, 193, 194, 215), generating products with 5'-monophosphate and 3'-hydroxyl groups (233). It requires monovalent and divalent cations (Mg2+ or Mn2+) for activity (233). No simple consensus sequence on RNA substrates for recognition and cleavage by RNase E has yet been defined. Instead, it appears that cleavage is affected by the continuity of the AU-rich regions (188, 215). Recently, it has been shown that RNase E binds to the 5' end of RNA substrates, preferentially on monophosphate termini (195), and its activity depends on the 5' end (195, 196). Further study showed that N-Rne binds to 5'-monophosphate RNA termini, scans over the substrate in a 3' to 5' direction, and cuts the substrate when accessible cleavage sites are found (97). RNA secondary structures close to the cleavage sites, such as stem-loops, affect RNase E cleavage, but their role has been controversial. Secondary structures either may facilitate binding of RNase E or they may enhance the single-stranded structure of the cleavage sites (62, 93, 197, 236). On the other hand, stem-loops also were found to inhibit RNase E cleavage (21, 214), presumably due to blocking the interaction of RNase E with the 5' end or to inhibition of the 3' to 5' scanning of the enzyme on its substrate.
RNase E is responsible for the cleavage of a diverse set of RNA substrates in E. coli. It was first discovered as an activity responsible for converting the 9S RNA to a precursor of 5S rRNA (102, 104, 223; Fig. 2). Subsequently, it was shown to participate in mRNA degradation (250). RNase E cleavages of specific mRNAs have been well studied (see the EcoSal chapter "mRNA Stability" [chapter 4.6.4]). In a genome-wide analysis, steady-state levels of more than 40% of cellular mRNAs are found to increase at least 1.5-fold upon depletion of RNase E (169). RNase E also participates in maturation of 16S rRNA (180; Fig. 2), tRNA (177, 253; Fig. 1), tmRNA (187), and M1 RNA (192, 294). It also cleaves the antisense RNAI of pBR322 and several other small RNAs with antisense or regulatory functions (24, 72, 95, 136, 186, 204, 228, 290, 296). In almost all cases, RNase E cleavage is the initial step of the processing or degradation pathways. Upon inactivation of RNase E mutants at a nonpermissive temperature, or upon overexpression of RraA, an inhibitor of RNase E (170), genome-wide accumulation of RNase E-targeted transcripts occurs (169, 170, 177, 253, 263). It is therefore not surprising that RNase E is an essential enzyme in E. coli.
A recent study demonstrated that overexpression of an extended form of RNase G restores growth of rne mutant cells without affecting tRNA processing (76). These data suggest that the effect of RNase E on tRNA maturation is not responsible for the inability of rne mutant cells to grow, in contrast to an earlier suggestion (253). Several lines of evidence suggest that, in addition to RNA degradation and processing, RNase E may also be involved in other cellular activities (39, 233), but these need to be examined in more detail.
The gene encoding E. coli RNase E has been named rne, ams, and hmp (14, 39, 104, 230, 250). It is located at 24.6 min on the genetic map of E. coli (104, 283). The rne gene is transcribed from three promoters, although the promoter closest to the start codon is most active (53, 129, 254). The 3.6-kb rne mRNA is a substrate of RNase E itself, providing a mechanism for RNase E to autoregulate its own expression (129, 229). Autoregulation appears to require transcription from the less active promoters (254). An evolutionarily conserved RNA stem-loop in the 5'-UTR was shown to affect the degradation rate of rne mRNA (84, 137). At least 10 to 20% of the normal cellular level of RNase E is required for cell growth, and autoregulation is able to increase RNase E synthesis when its concentration drops below normal levels (130, 296a). In addition, synthesis of RNase E is promoted when cellular poly(A) levels increase (226). Moreover, cellular RNase E concentration is affected by medium composition (168). Recently, a specific protein inhibitor of RNase E, RraA, was identified (170). This evolutionarily conserved 17.4-kDa protein interacts with RNase E without affecting its cleavage specificity. The C-terminal domain of RNase E is not required for the inhibition, but it does modulate the action of RraA.
RNase G.
RNase G was first identified as the CafA protein since its overexpression causes the formation of cytoplasmic axial filaments (249), and subsequently it was found to be an endoribonuclease and named RNase G (180, 327). The protein shares 34% identity and 50% similarity with the N-terminal 470 amino acids of RNase E (212, 213, 313) and it also shares some functional overlap with RNase E (326). Interruption of the rng gene shows no growth defect, but an rne-ts mutant plates less well in the rng background at semipermissive temperatures. RNase G can partially suppress the temperature-sensitive growth phenotype of rne mutant bacteria when expressed from a multicopy plasmid. The RNase activity of RNase G was proven with the demonstration that, together with RNase E, it is needed to generate the mature 5' end of 16S rRNA (180, 327).
RNase G polypeptide is 488 amino acids in length with a predicted molecular mass of 55 kDa. It is translated from the second possible start codon in the mRNA followed by removal of the N-terminal methionine residue (25). Purified RNase G exists largely as a dimer, but it is in equilibrium with monomers and higher multimers (25). Dimerization is required for activity. Substitution of two of its cysteine residues, C405 and C408, with serine results in a partial loss of activity, and a shift in the distribution of RNase G multimers toward monomers.
RNase G functions in vitro at pH 7.5 in the presence of 10 mM Mg2+ and ~100 mM monovalent cations (137, 180, 315). Purified RNase G cleaves single-stranded AU-rich sequences, similar to RNase E (137, 315).Some RNase E substrates, such as RNAI and its derivatives, and a region from ompA mRNA, can be cleaved by purified RNase G. However, the 5'-RNase E cleavage site of pre-5S RNA is not cleaved by RNase G (137, 315). Poly(A), a proven substrate of RNase E (121), is poorly cut by RNase G (315). Like RNase E, RNase G also depends on the 5' end of the RNA substrate for its activity (25, 137, 315). However, RNase G has a nondirectional and distributive mode of action, different from the scanning mode of RNase E (97).
RNase G is responsible for maturation of the 5' terminus of 16S rRNA in E. coli (180, 327; Fig. 2). When both RNase E and G are inactive, the processing of 17S precursor stops. Efficient production of the mature 5' end by RNase G depends on an initial cleavage at the +66 position by RNase E. In the absence of RNase G, the +66-nt 16.3S intermediate accumulates. It can be converted slowly to 16S RNA by additional RNase E cleavages close to the mature 5' end. The processing reactions occur when the precursors are assembled into ribosome particles, rather than as free RNA (71). A mutant, termed BUMMER, previously shown to accumulate the 16.3S product (71) was found to have a deletion in the coding region of the rng gene (327).
RNase G also participates in mRNA decay. It is involved, for example, in degradation of adhE and eno mRNAs (142, 318, 325). A genome-wide survey identified 18 mRNAs whose abundance is increased more than 1.5-fold upon deletion of rng, identifying these mRNAs as possible targets of RNase G action (169). Eleven of the mRNAs were decreased in abundance when RNase G was overexpressed. The transcripts affected are mostly involved in utilization of energy sources and stress responses, suggesting a role for RNase G in these processes. In addition, RNase G overexpression reduces the amount of about 100 mRNAs whose levels are increased in an rne deletion mutant, suggesting that RNase G can complement RNase E deficiency for some mRNAs (169). However, neither RNase G nor N-Rne can restore the decay of these RNAs as completely as does RNase E. These data indicate a limited role for RNase G in mRNA decay.
The limited role of RNase G is supported by observations that its cellular concentration is only 3% that of RNase E (169). Although it plays some role in mRNA degradation and in the processing of 9S rRNA, it does not participate in either tRNA or M1 RNA maturation, processes normally carried out by RNase E (255). RNase G’s limited role may be due to its inability to cleave important RNase E substrates. In fact, overexpression of RNase G to 20 times the normal level of RNase E does not rescue an rne mutant phenotype. In contrast, some unnatural forms of RNase G that contain extensions at their termini are better able to complement RNase E deficiency (76).
RNase G is encoded by the rng gene located at 73.2 min on the E. coli genetic map (283, 324). Little is known about expression of RNase G. Overexpression of the enzyme causes abnormal cellular structure and morphology, a phenomenon not yet understood (249). RNase E may downregulate the expression of RNase G by stimulating decay of the rng transcript (169). In RNase E-deficient cells, the steady-state level of rng mRNA increases by threefold, whereas the amount of RNase G protein produced from a multicopy plasmid is increased fourfold.
RNase H selectively hydrolyzes the RNA strand of RNA·DNA hybrids (65, 144). Members of the RNase H family are widely distributed among prokaryotic and eukaryotic organisms (58, 65) in three distinct lineages, RNases HI, HII, and HIII (247). Two of them, RNases HI and HII, are present in E. coli as well as in S. enterica. RNases HII and HIII are homologous and are believed to have arisen from a common ancestor, whereas RNase HI has limited sequence homology with the other members and is believed to be a product of convergent evolution (124, 167).
RNase HI.
E. coli RNase HI has been the subject of extensive reviews (118, 144). The enzyme, originally designated as RNase H prior to the discovery of RNase HII (124), is encoded by the rnhA gene. The enzyme was identified based on its cleavage of the RNA strand in a RNA·DNA duplex and is now believed to participate in DNA replication and repair (64). It is a protein of 155 amino acids with a calculated mass of 17.6 kDa (143) and functions as a monomer. RNase HI accounts for more than 90% of the RNase H activity in E. coli extracts (65) and has been purified from both normal and overexpressing cells (144). Cleavage by RNase HI produces 5'-phosphate and 3'-hydroxyl termini. The enzyme requires Mg2+ for activity, but this can be partially substituted by Mn2+ (17).
Crystal structures of E. coli RNase HI have been determined at high resolution (149, 150, 333). In addition, the solution structure has been resolved by NMR (100). Based on these studies, RNase HI contains five α-helices, five β-strands, and five reverse turns. A conserved triad of carboxylate side chains, Asp10, Glu48, and Asp70 (89), clustered on a concave surface, are essential for catalysis (148, 151). Site-directed mutagenesis of these residues abolishes the catalytic activity of RNase HI (146). A single divalent metal ion binds in the vicinity of the triad (119, 149, 244), and four other highly conserved residues, Ser71, His124, Asn130, and Asp134 (89), also are located nearby. Mutational analysis of His124 and Asp134 indicates that these residues also are involved in the catalytic function of the enzyme (115, 245).
Two different mechanisms have been proposed for catalysis by RNase HI: a two-metal-ion mechanism (75, 333) and a general acid-base mechanism that involves only a single metal ion (232, 243). The latter mechanism seems more likely based on a variety of X-ray (151), NMR (244), biochemical (119), and enzymatic (152) studies. In the currently proposed general acid-base model (152, 317), Asp10 and Asp70 provide ligands for metal binding, His124 activates a H2O molecule that acts as the general base and Glu48 anchors a second H2O molecule that acts as the general acid. The metal ion coordinates with the scissile phosphodiester bond through a H2O molecule. It is thought that Glu48 is required for Mg2+- but not for Mn2+-mediated catalysis (317) and that a second Mn2+ can bind through interactions with Asp10 and Asp134 and thereby inhibit catalysis (107, 152). Consistent with these notions, Glu48 and Asp134 are dispensable for the Mn2+-dependent activity (317).
Substrate recognition has been studied by X-ray, NMR, and site-directed mutagenesis (144, 145, 150, 232, 333), and many amino acid residues have been shown to be important for substrate binding. For example, upon substitution of these residues, the Kmvalue for substrate binding is greatly increased (144). Some of these residues, such as Thr43, Asn44, Asn45, and Asn130, are highly conserved among RNase HIs. In addition, a "basic protrusion" consisting of seven basic amino acids starting with Lys87 is important for substrate binding through charge interactions (145). This information, combined with docking simulations of the structures of the free enzyme and RNA·DNA hybrids, has led to a model of how RNase HI interacts with its substrate (118, 144). RNase HI is proposed to interact with the minor groove of the duplex, in which the aforementioned amino acid residues and the active site make contact with the RNA or DNA. It seems that RNA·DNA hybrids assume neither the A-type nor the B-type geometry (96), suggesting a means by which the enzyme might discriminate between RNA·DNA hybrids and pure DNA or RNA duplexes (118, 144).
RNase HI shows little nucleotide specificity for the position of cleavage, although there may be a slight preference for phosphodiester bonds adjacent to pyrimidines (144). A DNA-RNA sequence as small as a tetramer embedded within a larger DNA or RNA duplex can be recognized and cleaved by RNase HI. The enzyme also can cleave a homopolymeric poly(rA)·poly(dT) duplex, yielding mainly tri- and tetra-adenylates as limit products of digestion. The low yield of AMP indicates that the enzyme does not contain exonuclease activity.
RNase HI does not cleave phosphodiester bonds at the junction of RNA and DNA. Likewise, it can bind, but it does not cleave an RNA·RNA duplex (184). RNase H, in general, is quite sensitive to modifications of the DNA (222) or RNA (334) strands in the substrate. Alterations of the structure of the RNA or the DNA also can influence RNase HI activity (173, 185). However, single-stranded RNA adjacent to an RNA·DNA duplex can be cleaved (184), indicating that RNase HI can tolerate some limited substrate variation.
The substrate specificity of RNase HI suggests that it is involved in DNA replication and repair. In fact, it was found that RNase HI participates in the replication of ColE1 plasmids and in the replication of chromosomal DNA from oriC (118, 144, 160). RNase HI is required for the processing of the RNA transcripts that are used as primers for replication of ColE1 plasmids in vitro, and it may also be responsible for eliminating the primers after initiation of replication (127). Nevertheless, replication of ColE1 plasmids can take place in rnhA mutants by initiating at sites other than Ori, such as by using RNA that results from aberrant transcription termination (113). Furthermore, RNase HI can suppress replication initiated outside of Ori (144).
The role of RNase HI in chromosomal DNA replication includes removal of RNA primers from Okazaki fragments (246) and degradation of RNA in R-loops (90, 323). The latter action would help ensure that replication initiates from OriC. However, these activities are nonessential since other enzymes can substitute for RNase HI. For example, removal of R loops can be carried out by two alternative pathways, both of which depend on recBCD (161). However, inactivation of components of recBCD in the rnhA background is lethal (125).
The rnhA gene encoding RNase HI is located at 5.1 min of the E. coli genetic map (283). The promoter of rnhA overlaps with that of dnaQ, which runs in the opposite direction (199). In E. coli mutants constitutively expressing the pleiotropic SOS response, a strong reduction in rnhA and an increase in dnaQ transcription are observed, suggesting that the two genes are inversely regulated (271). It is estimated that about 100 RNase HI molecules are present per cell (128), but the enzyme can be overexpressed 15-fold without affecting cell growth (143). On the other hand, a 1,000-fold overproduction is detrimental to cell growth (144).
RNase HII.
Upon screening an E. coli genomic library for clones that could rescue the lethal phenotype of a mutant strain lacking rnhA and recC, the rnhB gene encoding RNase HII was identified (124). The rnhB gene is located at 4.4 min on the E. coli genetic map and is part of an operon that contains the lpxA and lpxB genes upstream and the dnaE gene downstream (124, 283). RNase HII is a polypeptide of 198 amino acid residues with a calculated mass of 21.5 kDa. Its sequence is only 17% identical to E. coli RNase HI, and the identical residues do not overlap with the residues deemed to be important for RNase HI. Purified RNase HII displays a molecular mass of 23 kDa on SDS-PAGE and of 22 kDa on gel filtration, suggesting that it acts as a monomer (124, 248). Based on analysis of its CD spectrum, the protein has a helical content of 23% (248). Although currently nothing is known about the structure of the E. coli enzyme, comparison of the crystal structures of E. coli RNase HI and Thermococcus kodakaraensis RNase HII (231) indicates that these two enzymes share main-chain folds and suggests that RNase HI and RNase HII proteins are structurally homologous.
Although overexpressed RNase HII complements the lethality of rnhA, recC mutations, its primary function may be related to repair of misincorporated ribonucleotides in DNA (284). Using double-stranded oligonucleotide substrates containing a single ribose residue, an activity was detected in E. coli extracts that nicks at the 5' side of the ribose. This activity is missing in extracts of an rnhB mutant. The activity depends on Mn2+. A similar activity can be detected in human and yeast extracts and is also shown by a highly purified archaeal RNase HII protein. The yeast RNase HII was found to preferentially cleave a substrate that contains a single ribose residue compared with a substrate that contains a stretch of ribose residues. This observation strongly suggests that these enzymes are responsible for removal of misincorporated ribonucleotides in DNA (284). An E. coli mutant strain that lacks both RNase HI and RNaseHII is viable at low temperature, suggesting that these enzymes, although carrying out important functions, are nevertheless dispensable (126).
RelE and MazF.
Very recently, the catalogue of E. coli endoribonucleases has been expanded with the discovery that the known toxins, RelE (49) and MazF (2), participate in RNA cleavage. Both toxins are part of toxin-antitoxin pairs and normally are in an inhibited state during balanced growth. However, physiological conditions that lead to depletion of the antitoxin result in release of free toxin and ultimately cessation of growth. For both RelE and MazF, their mechanism of action appears to be specific cleavage of mRNAs and consequent inhibition of protein synthesis (262, 344).
relE is cotranscribed with the antitoxin gene relB (16), and their protein products form an inactive RelE-RelB complex during normal growth (101). Upon conditions that release free RelE, protein synthesis is strongly inhibited (261). A cell-free E. coli translation system was used to examine the mechanism of this inhibition (262). It was found that purified RelE leads to cleavage of mRNA in the A site of the ribosome and that the cleavage is highly codon specific. For example, among termination codons, cleavage occurs in the order UAG > UAA > UGA. Likewise, certain sense codons, such as UCG and CAG, are cleaved more rapidly than others. Cleavage generally occurs between the second and third nucleotides of the codon in the A site.
As a consequence of the cleavage reaction, ribosomes become stalled on the now incomplete mRNAs. Such structures are substrates for the action of tmRNA, and recent work has shown that tmRNA can relieve the toxicity of RelE by rescuing the stalled ribosomes (48). Free mRNA is not a substrate for RelE. This observation raises the question of whether RelE is an endoribonuclease itself or whether it might activate a nucleolytic activity associated with the ribosome. Nevertheless, further studies of this interesting mRNA degradation system are awaited.
The second toxin, MazF, is part of a toxin-antitoxin system together with MazE (2). As with the relBE system, mazEF consists of adjacent genes. MazE and MazF form a linear heterohexamer structure composed of alternating toxin and antitoxin homodimers (MazF2-MazE2-MazF2) (142a). The labile MazE portion is depleted under certain conditions, such as treatment with certain antibiotics (286a), which results in activation of MazF. Examination of the action of MazF in permeabilized E. coli cells revealed that it inhibits protein synthesis (344). In a cell-free system, purified MazF was found to cleave mRNA, specifically between the A and C residues in an ACA sequence (344). In contrast to RelE, purified MazF cleaves free RNA and thus it clearly is an endoribonuclease. It does not cleave DNA, RNA-DNA hybrids, or RNA-RNA duplexes. It can also act on rRNA, when freed of ribosomal proteins. These data contrast with the suggestion (50) that MazF acts in a manner similar to RelE. Another toxin, ChpBK, is a homologue of MazF and can cleave mRNA in the same manner as MazF (50).
There are two promoters with "alternating palindromic" sequences upstream of mazEF. The MazE-MazF complex binds these sequences, resulting in strong negative autoregulation (202). This allows mazEF expression to respond sensitively to various growth conditions.
The discovery of these endoribonuclease toxins with such unusual specificities raises many questions about their cellular roles and whether other similar RNases might also be present in cells. Cleavage of mRNA A-site codons on paused ribosomes can occur even in cells lacking RelE, MazF, and ChpBK, indicating that other cleavage mechanisms must exist (116).
It is likely that E. coli contains additional endoribonucleases that have not yet been characterized. First of all, endonucleolytic activities are needed for certain known processes that cannot be attributed to any of the known enzymes. These include those responsible for maturation of the 5' termini of 23S, 5S rRNA and M1 RNA, and the 3' terminus of 16S rRNA (79, 98, 117, 179, 180, 192). Second, homologues of known endoribonucleases are present in E. coli. A clear example is the ElaC protein (58), a homologue of RNase Z, that is known to process the 3' termini of tRNA precursors in other organisms (289). Third, endonucleolytic activities have been observed in cell extracts that have different properties from known enzymes. For instance, the activity attributed to RNase M (36, 218) may be conferred by an as yet unidentified protein. Clearly, further work on this interesting class of enzymes is warranted.
It is already clear that E. coli, and also Salmonella species, make a large investment in RNases to carry out the reactions of RNA metabolism. Our knowledge of this class of enzymes has grown dramatically in recent years, but much remains to be learned. In particular, it is likely that we have not yet identified all the RNases present in these cells, as there are several RNA-processing events that have not been associated with any known enzyme. In addition, the discovery that several toxins are RNases raises the possibility that additional enzymes of this type will be found. Second, there are several RNases whose functions are not yet understood, and these need to be determined. Interesting work has begun on the structures and regulation of several of the RNases, and it is to be expected that studies along these lines will expand. The culmination of all these efforts will provide a detailed picture of all the reactions of RNA metabolism and of the enzymes responsible for them.
References
1. Aiso, T., and R. Ohki. 1998. A rne-1 pnp-7 double mutation suppresses the temperature sensitive defect of lacZ expression in a divE mutant. J. Bacteriol. 180:1389–1395.[PubMed]
2. Aizenman, E., H. Engelberg-Kulka, and G. Glaser. 1996. An Escherichia coli chromosomal "addiction module" regulated by guanosine [corrected] 3’, 5’-bispyrophosphate: a model for programmed bacterial cell death. Proc. Natl. Acad. Sci. USA 93:6059–6063. [CrossRef]
3. Alifano, P., F. Rivellini, C. Piscitelli, C. M. Arraiano, C. B. Bruni, and M. S. Carlomagno. 1994. Ribonuclease E provides substrates for ribonuclease P-dependent processing of a polycistronic mRNA. Genes Dev. 8:3021–2031. [CrossRef]
4. Allas, U., A. Liiv, and J. Remme. 2003. Functional interaction between RNase III and the Escherichia coli ribosome. BMC Mol. Biol. 4:8. [CrossRef]
5. Altman, S. 1990. Ribonuclease P. Postscript. J. Biol. Chem. 265:20053–20056.
6. Altman, S. 1995. RNase P in research and therapy. Bio/Technology 13:327–329. [CrossRef]
7. Altman, S., and L. A. Kirsebom. 1999. Ribonuclease P, p. 351–380. In R. F. Gesteland, T. R. Cech, and J. F. Atkins (ed.), The RNA World.Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
8. Andersen, J. T., P. Poulsen, and K.F. Jensen. 1992. Attenuation in the rph-pyrE operon of Escherichia coli and processing of the dicistronic mRNA. Eur. J. Biochem. 206:381–390. [CrossRef]
9. Anderson, P. E., J. Matsunaga, E. L. Simons, and R. W. Simons. 1996. Structure and regulation of the Salmonella typhimurium rnc-era-recO operon. Biochimie 78:1025–1034. [CrossRef]
10. Anraku, Y., and D. Mizuno. 1967. Comparative study on the ribonucleases isolated from the debris and ribosome fraction of Escherichia coli. J. Biochem. (Tokyo) 61:70–80.
11. Aristarkhov, A., A. Mikulskis, J. G. Belasco, and E. C. Lin. 1996. Translation of the adhE transcript to produce ethanol dehydrogenase requires RNase III cleavage in Escherichia coli. J. Bacteriol. 178:4327–4332.
12. Asha, P. K., R. T. Blouin, R. Zaniewski, and M. P. Deutscher. 1983. Ribonuclease BN: identification and partial characterization of a new tRNA processing enzyme. Proc. Natl. Acad. Sci. USA 80:3301–3304. [CrossRef]
13. Asha, P. K., and M. P. Deutscher. 1983. Escherichia coli strain CAN lacks a tRNA-processing nuclease. J. Bacteriol. 156:419–420.
14. Babitzke, P., and S. R. Kushner. 1991. The Ams (altered mRNA stability) protein and ribonuclease E are encoded by the same structural gene of Escherichia coli. Proc. Natl. Acad. Sci. USA 88:1–5. [CrossRef]
15. Bardwell, J. C., P. Regnier, S. M. Chen, Y. Nakamura, M. Grunberg-Manago, and D. L. Court. 1989. Autoregulation of RNase III operon by mRNA processing. EMBO J. 8:3401–3407.
16. Bech, F. W., S. T. Jorgensen, B. Diderichsen, and O. H. Karlstrom. 1985. Sequence of the relB transcription unit from Escherichia coli and identification of the relB gene. EMBO J. 4:1059–1066.
17. Berkower, I., J. Leis, and J. Hurwitz. 1973. Isolation and characterization of an endonuclease from Escherichia coli specific for ribonucleic acid in ribonucleic acid-deoxyribonucleic acid hybrid structures. J. Biol. Chem. 248:5914–5921.
18. Biswas, R., D. W. Ledman, R. O. Fox, S. Altman, and V. Gopalan. 2000. Mapping RNA-protein interactions in ribonuclease P from Escherichia coli using disulfide-linked EDTA-Fe. J. Mol. Biol. 296:19–31. [CrossRef]
19. Blaszczyk, J., J. E. Tropea, M. Bubunenko, K. M. Routzahn, D. S. Waugh, D. L. Court, and X. Ji. 2001. Crystallographic and modeling studies of RNase III suggest a mechanism for double-stranded RNA cleavage. Structure (Cambridge) 9:1225–1236. [CrossRef]
20. Blouin, R. T., R. Zaniewski, and M. P. Deutscher. 1983. Ribonuclease D is not essential for the normal growth of Escherichia coli or bacteriophage T4 or for the biosynthesis of a T4 suppressor tRNA. J. Biol. Chem. 258:1423–1426.
21. Bouvet, P., and J. G. Belasco. 1992. Control of RNase E-mediated RNA degradation by 5'-terminal base pairing in E. coli. Nature 360:488–491. [CrossRef]
22. Bram, R. J., R. A. Young, and J. A. Steitz. 1980. The ribonuclease III site flanking 23S sequences in the 30S ribosomal precursor RNA of E. coli. Cell 19:393–401. [CrossRef]
23. Brännvall, M., B. M. F. Pettersson, and L. A. Kirsebom. 2003. Importance of the +73/294 interaction in Escherichia coli RNase P RNA substrate complexes for cleavage and metal ion coordination. J. Mol. Biol. 325:697–709. [CrossRef]
24. Briani, F., E. Del Vecchio, D. Migliorini, E. Hajnsdorf, P. Regnier, D. Ghisotti, and G. Deho. 2002. RNase E and polyadenyl polymerase I are involved in maturation of CI RNA, the P4 phage immunity factor. J. Mol. Biol. 318:321–331. [CrossRef]
25. Briant, D. J., J. S. Hankins, M. A. Cook, and G. A. Mackie. 2003. The quaternary structure of RNase G from Escherichia coli. Mol. Microbiol. 50:1381–1390. [CrossRef]
26. Burgin, A. B., K. Parodos, D. J. Lane, and N. R. Pace. 1990. The excision of intervening sequences from Salmonella 23S ribosomal RNA. Cell 60:405–414. [CrossRef]
27. Bycroft, M., T. J. Hubbard, M. Proctor, S. M. Freund, and A. G. Murzin. 1997. The solution structure of the S1 RNA binding domain: a member of an ancient nucleic acid-binding fold. Cell 88:235–242. [CrossRef]
28. Cairrao, F., A. Chora, R. Zilhao, A. J. Carpoussis, and C. M. Arraiano. 2001. RNase II levels change according to growth conditions: characterization of gmr, a new Escherichia coli gene involved in the modulation of RNase II. Mol. Microbiol. 39:1550–1561. [CrossRef]
29. Cairrao, F., A. Cruz, H. Mori, and C. M. Arraiano. 2003. Cold shock induction of RNase R and its role in the maturation of the quality control mediator SsrA/tmRNA. Mol. Microbiol. 50:1349–1360. [CrossRef]
30. Calin-Jageman, I., and A. W. Nicholson. Escherichia coli ribonuclease III substrates. Biochemistry 42:5025–5034. [CrossRef]
31. Callaghan, A. J., J. G. Grossmann, Y. U. Redko, L. L. Ilag, M. C. Moncrieffe, M. F. Symmons, C. V. Robinson, K. J. McDowall, and B. F. Luisi. 2003. Quaternary structure and catalytic activity of the Escherichia coli ribonuclease E amino-terminal catalytic domain. Biochemistry 42:13848–13855. [CrossRef]
32. Callahan, C., and M. P. Deutscher. 1996. Identification and characterization of the Escherichia coli rbn gene encoding the tRNA processing enzyme RNase BN. J. Bacteriol. 178:7329–7332.
33. Callahan, C., D. Neri-Cortes, and M. P. Deutscher. 2000. Purification and characterization of the tRNA processing enzyme RNase BN. J. Biol. Chem. 275:1030–1034. [CrossRef]
33a. Cannistraro, V. J., and D. Kennell. 1989. Purification and characterization of ribonuclease M and mRNA degradation in Escherichia coli. Eur. J. Biochem. 181:363–370. [CrossRef]
34. Cannistraro, V. J., and D. Kennell. 1991. RNase I*, a form of RNase I, and mRNA degradation in Escherichia coli. J. Bacteriol. 173:4653–4659.[PubMed]
35. Cannistraro, V. J., and D. Kennell. 1994. The processive reaction mechanism of ribonuclease II. J. Mol. Biol. 243:930–943. [CrossRef]
36. Cannistraro, V. J., and D. Kennell. 1999. The reaction mechanism of ribonuclease II and its interaction with nucleic acid secondary structures. Biochim. Biophys. Acta 1433:170–187.
37. Carpousis, A. J. 2002. The Escherichia coli degradosome: structure, function and relationship to other ribonucleolytic multienzyme complexes. Biochem. Soc. Trans. 30(Part 2):150–156.
38. Carpousis, A. J., G. Van Houwe, C. Ehretsmann, and H. M. Krisch. 1994. Copurification of E. coli RNAase E and PNPase: evidence for a specific association between two enzymes important in RNA processing and degradation. Cell 76:889–900. [CrossRef]
39. Casaregola, S., A. Jacq, D. Laoudj, G. McGurk, S. Margarson, M. Tempete, V. Norris, and I. B. Holland. 1992. Cloning and analysis of the entire Escherichia coli ams gene. ams is identical to hmp1 and encodes a 114 kDa protein that migrates as a 180 kDa protein. J. Mol. Biol. 228:30–40. (Erratum, 238:867.)[PubMed] [CrossRef]
40. Case, L. M., X. N. Chen, and M. P. Deutscher. 1989. Localization of the Escherichia coli rnt gene encoding RNase T by using a combination of physical and genetic mapping. J. Bacteriol. 171:5736–5737.
41. Chakraburtty, K., and D. P. Burma. 1968. The purification and properties of a ribonuclease from Salmonella typhimurium extract. J. Biol. Chem. 243:1133–1139.
42. Chelladurai, B., H. Li, K. Zhang, and A. W. Nicholson. 1993. Mutational analysis of a ribonuclease III processing signal. Biochemistry 32:7549–7558. [CrossRef]
43. Chen, S. M., H. E. Takiff, A. M. Barber, G. C. Dubois, J. C. Bardwell, and D. L. Court. 1990. Expression and characterization of RNase III and Era proteins. Products of the rnc operon of Escherichia coli. J. Biol. Chem. 265:2888–2895.
44. Cheng, Z-F., and M. P. Deutscher. 2002. Purification and characterization of the Escherichia coli exoribonuclease RNase R. J. Biol. Chem. 277:21624–21629. [CrossRef]
45. Cheng, Z-F., and M. P. Deutscher. 2003. Quality control of ribosomal RNA mediated by polynucleotide phosphorylase and RNase R. Proc. Natl. Acad. Sci. USA 100:6388–6393. [CrossRef]
46. Cheng, Z-F., Y. Zuo, Z. Li, K. E. Rudd, and M. P. Deutscher. 1998. The vacB gene required for virulence in Shigella flexneri and Escherichia coli encodes the exoribonuclease RNase R. J. Biol. Chem. 273:14077–14080. [CrossRef]
47. Choi, J. M., E. Y. Park, J. H. Kim, S. K. Chang, and Y. Cho. 2004. Probing the functional importance of the hexameric ring structure of RNase PH. J. Biol. Chem. 279:755–764. [CrossRef]
48. Christensen, S. K. and K. Gerdes. 2003. RelE toxins from bacteria and archaea cleave mRNAs on translating ribosomes, which are rescued by tmRNA. Mol. Microbiol. 48:1389–1400. [CrossRef]
49. Christensen, S. K., M. Mikkelsen, M. K. Pedersen, and K. Gerdes. 2001. RelE, a global inhibitor of translation, is activated during nutritional stress. Proc. Natl. Acad. Sci. USA 98:14328–14333.[PubMed] [CrossRef]
50. Christensen, S. K., K. Pedersen, F. G. Hansen, and K. Gerdes. 2003. Toxin-antitoxin loci as stress-response-elements: ChpAK/MazF and ChpBK cleave translated RNAs and are counteracted by tmRNA. J. Mol. Biol. 332:809–819. [CrossRef]
51. Christian, E. L., N. M. Kaye, and M. E. Harris. 2002. Evidence for a polynuclear metal ion binding site in the catalytic domain of ribonuclease P RNA. EMBO J. 21:2253–2262. [CrossRef]
52. Christian, E. L., N. H. Zahler, N. M. Kaye, and M. E. Harris. 2002. Analysis of substrate recognition by the ribonucleoprotein endonuclease RNase P. Methods 28:307–322. [CrossRef]
53. Claverie-Martin, F., M. R. Diaz-Torres, S. D. Yancey, and S. R. Kushner. 1991. Analysis of the altered mRNA stability (ams) gene from Escherichia coli. Nucleotide sequence, transcriptional analysis, and homology of its product to MRP3, a mitochondrial ribosomal protein from Neurospora crassa. J. Biol. Chem. 266:2843–2851.
54. Coburn, G. A., and G. A. Mackie. 1996. Overexpression, purification and properties of Escherichia coli ribonuclease II. J. Biol. Chem. 271:1048–1053. [CrossRef]
55. Coburn, G. A., and G. A. Mackie. 1998. Reconstitution of the degradation of the mRNA for ribosomal protein S20 with purified enzymes. J. Mol. Biol. 279:1061–1074. [CrossRef]
56. Coburn, G. A., X. Miao, D. J. Briant, and G. A. Mackie. 1999. Reconstitution of a minimal RNA degradosome demonstrates functional coordination between a 3' exonuclease and a DEAD-box RNA helicase. Genes Dev. 13:2594–2603. (Erratum, 14:1167.) [CrossRef]
57. Cohen, S. N., and K. J. McDowall. 1997. RNase E: still a wonderfully mysterious enzyme. Mol. Microbiol. 23:1099–1106. [CrossRef]
58. Condon, C., and H. Putzer. 2002. The phylogenetic distribution of bacterial ribonucleases. Nucleic Acids Res. 30:5339–5346. [CrossRef]
59. Conrad, C., E. Evguenieva-Hackenberg, and G. Klug. 2001. Both N-terminal catalytic and C-terminal RNA binding domain contribute to substrate specificity and cleavage site selection of RNase III. FEBS Lett. 509:53–58. [CrossRef]
60. Conrad, C., and R. Rauhut. 2002. Ribonuclease III: new sense from nuisance. Int. J. Biochem. Cell Biol. 34:116–129. [CrossRef]
61. Cormack, R. S., J. L. Genereaux, and G. A. Mackie. 1993.RNase E activity is conferred by a single polypeptide: overexpression, purification, and properties of the ams/rne/hmp1 gene product. Proc. Natl. Acad. Sci. USA 90:9006–9010. [CrossRef]
62. Cormack, R. S., and G. A. Mackie. 1992. Structural requirements for the processing of Escherichia coli 5 S ribosomal RNA by RNase E in vitro. J. Mol. Biol. 228:1078–1090. [CrossRef]
63. Court, D. 1993. RNA processing and degradation by RNase III, p. 71–116. In J. G. Elasco and G. Brawerman (ed.), Control of Messenger RNA Stability. Academic Press, New York, N.Y.
64. Crary, S. M., S. Niranjanakumari, and C. A. Fierke. 1998. The protein component of Bacillus subtilis ribonuclease P increases catalytic efficiency by enhancing interactions with the 5' leader sequence of pre-tRNAAsp. Biochemistry 37:9409–9416. [CrossRef]
65. Crouch, R. J. 1990. Ribonuclease H: from discovery to 3D structure. New Biol. 2:771–777.
66. Cruz, A. A., P .E. Marujo, S. F. Newbury, and C. M. Arraiano. 1996. A new role for RNase II in mRNA decay. FEMS Microbiol. Lett. 145:315–324. [CrossRef]
67. Cudny, H., and M. P. Deutscher. 1980. Apparent involvement of ribonuclease D in the 3’ processing of tRNA precursors. Proc. Natl. Acad. Sci. USA 77:837–841. [CrossRef]
68. Cudny, H., and M. P. Deutscher. 1988. 3’ processing of tRNA precursors in ribonuclease-deficient Escherichia coli. Development and characterization of an in vitro processing system and evidence for a phosphate requirement. J. Biol. Chem. 263:1518–1523.
69. Cudny, H., R. Zaniewski, and M. P. Deutscher. 1981. Escherichia coli RNase D. catalytic properties and substrate specificity. J. Biol. Chem. 256:5633–5637.
70. Cudny, H., R. Zaniewski, and M. P. Deutscher. 1981. Escherichia coli RNase D. Purification and structural characterization of a putative processing nuclease. J. Biol. Chem. 256:5627–5632.
71. Dahlberg, A .E., J. E. Dahlberg, E. Lund, H. Tokimatsu, A. B. Rabson, P. C. Calvert, F. Reynolds, and M. Zahalak. 1978. Processing of the 5' end of Escherichia coli 16S ribosomal RNA. Proc. Natl. Acad. Sci. USA 75:3598–3602. [CrossRef]
72. Dam Mikkelsen, N., and K. Gerdes. 1997. Sok antisense RNA from plasmid R1 is functionally inactivated by RNase E and polyadenylated by poly(A) polymerase I. Mol. Microbiol. 26:311–320. [CrossRef]
73. Dasgupta, S., L. Fernandez, L. Kameyama, T. Inada, Y. Nakamura, A. Pappas, and D. L. Court. 1998. Genetic uncoupling of the dsRNA-binding and RNA cleavage activities of the Escherichia coli endoribonuclease RNase III—the effect of dsRNA binding on gene expression. Mol. Microbiol. 28:629–640. (Erratum, 30:679.) [CrossRef]
74. Datta, A. K., and S. K. Niyogi. 1975. A novel oligoribonuclease of Escherichia coli. II. Mechanism of action. J. Biol. Chem. 250:7313–7319.
75. Davies, J. F., II, Z. Hostomska, Z. Hostomsky, S. R. Jordan, and D. A. Matthews. 1991. Crystal structure of the ribonuclease H domain of HIV-1 reverse transcriptase. Science 252:88–95. [CrossRef]
76. Deana, A., and J. G. Belasco. 2004. The function of RNase G in Escherichia coli is constrained by its amino and carboxyl termini. Mol. Microbiol. 51:1205–1217.[PubMed] [CrossRef]
77. Deutscher, M. P. 1990. Ribonucleases, tRNA nucleotidyltransferase, and the 3’ processing of tRNA. Prog. Nucleic Acids Res. Mol. Biol. 39:209–240. [CrossRef]
78. Deutscher, M. P. 2003. Degradation of stable RNA in bacteria. J. Biol. Chem. 278:45041–45044. [CrossRef]
79. Deutscher, M. P., and Z. Li. 2000. Exoribonucleases and their multiple roles in RNA metabolism. Prog. Nucleic Acids Res. Mol. Biol. 66:67–105. [CrossRef]
80. Deutscher, M. P., and C. W. Marlor. 1985. Purification and characterization of Escherichia coli RNase T. J. Biol. Chem. 260:7067–7071.
81. Deutscher, M. P., C. W. Marlor, and R. Zaniewski. 1984. Ribonuclease T: new exoribonuclease possibly involved in end-turnover of tRNA. Proc. Natl. Acad. Sci. USA 81:4290–4293. [CrossRef]
82. Deutscher, M. P., C. W. Marlor, and R. Zaniewski. 1985. RNase T is responsible for the end-turnover of tRNA in Escherichia coli. Proc. Natl. Acad. Sci. USA 82:6427–6430. [CrossRef]
83. Deutscher, M. P., G. T. Marshall, and H. Cudny. 1988. RNase PH: an Escherichia coli phosphate-dependent nuclease distinct from polynucleotide phosphorylase. Proc. Natl. Acad. Sci. USA 85:4710–4714. [CrossRef]
84. Diwa, A., A. L. Bricker, C. Jain, and J. G. Belasco. 2000. An evolutionarily conserved RNA stem-loop functions as a sensor that directs feedback regulation of RNase E gene expression. Genes Dev. 14:1249–1260.
85. Diwa, A. A., X. Jiang, M. Schapira, and J. G. Belasco. 2002. Two distinct regions on the surface of an RNA-binding domain are crucial for RNase E function. Mol. Microbiol. 46:959–969. (Erratum, 47:1183.) [CrossRef]
86. Dong, H., L. A. Kirsebom, and L. Nilsson. 1996. Growth rate regulation of 4.5 S RNA and M1 RNA the catalytic subunit of Escherichia coli RNase P. J. Mol. Biol. 261:303–308. [CrossRef]
87. Donovan, W. P., and S. R. Kushner. 1983. Amplification of ribonuclease II (rnb) activity in Escherichia coli K-12. Nucleic Acids Res. 11:265–275. [CrossRef]
88. Donovan, W. P., and S. R. Kushner. 1986. Polynucleotide phosphorylase and ribonuclease II are required for cell viability and mRNA turnover in Escherichia coli K-12. Proc. Natl. Acad. Sci. USA 83:120–124. [CrossRef]
89. Doolittle, R. F., D. F. Feng, M. S. Johnson, and M. A. McClure. 1989. Origins and evolutionary relationships of retroviruses. Q. Rev. Biol. 64:1–30. [CrossRef]
90. Drolet, M., P. Phoenix, R. Menzel, E. Masse, L. F. Liu, and R. J. Crouch. 1995. Overexpression of RNase H partially complements the growth defect of an Escherichia coli delta topA mutant: R-loop formation is a major problem in the absence of DNA topoisomerase I. Proc. Natl. Acad. Sci. USA 92:3526–3530. [CrossRef]
91. Dunn, J. J. 1976. RNase III cleavage of single-stranded RNA. Effect of ionic strength on the fidelity of cleavage. J. Biol. Chem. 251:3807–3814.
92. Dunn, J. J. 1982. Ribonuclease III, p. 485–499. In P. D. Boyer (ed.), The Enzymes, vol. 15. Academic Press, New York, N.Y.
93. Ehretsmann, C. P., A. J. Carpousis, and H. M. Krisch. 1992. Specificity of Escherichia coli endoribonuclease RNase E: in vivo and in vitro analysis of mutants in a bacteriophage T4 mRNA processing site. Genes Dev. 6:149–159. [CrossRef]
94. Evans, S., and P. P. Dennis. 1985. Promoter activity and transcript mapping in the regulatory region for genes encoding ribosomal protein S15 and polynucleotide phosphorylase of Escherichia coli. Gene 40:15–22. [CrossRef]
95. Faubladier, M., K. Cam, and J. P. Bouche. 1990. Escherichia coli cell division inhibitor DicF-RNA of the dicB operon. Evidence for its generation in vivo by transcription termination and by RNase III and RNase E-dependent processing. J. Mol. Biol. 212:461–471. [CrossRef]
96. Fedoroff, O. Y., M. Salazar, and B. R. Reid. 1993. Structure of a DNA:RNA hybrid duplex. Why RNase H does not cleave pure RNA. J. Mol. Biol. 233:509–523. [CrossRef]
97. Feng, Y, T. A. Vickers, and S. N. Cohen. 2002. The catalytic domain of RNase E shows inherent 3' to 5' directionality in cleavage site selection. Proc. Natl. Acad. Sci. USA 99:14746–14751. [CrossRef]
98. Feunteun, J., B. R. Jordan, and R. Monier. 1972. Study of the maturation of 5 s RNA precursors in Escherichia coli. J. Mol. Biol. 70:465–474. [CrossRef]
98a. Forster, A. C., and S. Altman. 1990. External guide sequences for an RNA enzyme. Science 249:783–786. [CrossRef]
99. Frank, D. N., and N. R. Pace. 1998. Ribonuclease P: unity and diversity in a tRNA processing ribozyme. Annu. Rev. Biochem. 67:153–180. [CrossRef]
100. Fujiwara, M., T. Kato, T. Yamazaki, K. Yamasaki, and K. Nagayam. 2000. NMR structure of ribonuclease HI from Escherichia coli. Biol. Pharm. Bull. 23:1147–1152.
101. Galvani, C., J. Terry, and E. E. Ishiguro. 2001. Purification of the RelB and RelE proteins of Escherichia coli: RelE binds to RelB and to ribosomes. J. Bacteriol. 183:2700–2703. [CrossRef]
102. Gegenheimer, P., N. Watson, and D. Apirion. 1977. Multiple pathways for primary processing of ribosomal RNA in Escherichia coli. J. Biol. Chem. 252:3064–3073.
103. Gesteland, R. F. 1966. Isolation and characterization of ribonuclease I mutants of Escherichia coli. J. Mol. Biol. 16:67–84. [CrossRef]
104. Ghora, B. K., and D. Apirion. 1978. Structural analysis and in vitro processing to p5 rRNA of a 9S RNA molecule isolated from an rne mutant of E. coli. Cell 15:1055–1066. [CrossRef]
105. Ghosh, R. K., and M. P. Deutscher. 1978. Identification of an Escherichia coli nuclease acting on structurally altered transfer RNA molecules. J. Biol. Chem. 253:997–1000.
106. Ghosh, S., and M. P. Deutscher. 1999. Oligoribonuclease is an essential component of the mRNA decay pathway. Proc. Natl. Acad. Sci. USA 96:4372–4377. [CrossRef]
107. Goedken, E. R., and S. Marqusee. 2001. Co-crystal of Escherichia coli RNase HI with Mn2+ ions reveals two divalent metals bound in the active site. J. Biol. Chem. 276:7266–7271. [CrossRef]
108. Gopalan, V., H. Kuhne, R. Biswas, H. Li, G. W. Brudvig, and S. Altman. 1999. Mapping RNA-protein interactions in ribonuclease P from Escherichia coli using electron paramagnetic resonance spectroscopy. Biochemistry 38:1705–1714. [CrossRef]
109. Grunberg-Manago, M. 1999. Messenger RNA stability and its role in control of gene expression in bacteria and phages. Annu. Rev. Genet. 33:193–227. [CrossRef]
110. Guerrier-Takada, C., K. Gardiner, T. Marsh, N. Pace, and S. Altman. 1983. The RNA moiety of ribonuclease P is the catalytic subunit of the enzyme. Cell 35:849–857. [CrossRef]
111. Gurevitz, M., N. Watson, and D. Apirion. 1982. A cleavage site of ribonuclease F. A putative processing endoribonuclease from Escherichia coli. Eur. J. Biochem. 124:553–559.
112. Haas, E.S., and J. W. Brown. 1998. Evolutionary variation in bacterial RNase P RNAs. Nucleic Acids Res. 26:4093–4099. [CrossRef]
112a. Hansen, F. G., E. B. Hansen, and T. Atlung. 1985. Physical mapping and nucleotide sequence of the rnpA gene that encodes the protein component of ribonuclease P in Escherichia coli. Gene 38:85–93. [CrossRef]
113. Harinarayanan, R., and J. Gowrishankar. 2003. Host factor titration by chromosomal R-loops as a mechanism for runaway plasmid replication in transcription termination-defective mutants of Escherichia coli. J. Mol. Biol. 332:31–46. [CrossRef]
114. Hartmann, R. K., J. Heinrich, J. Schlegl, and H. Schuster. 1995. Precursor of C4 antisense RNA of bacteriophages P1 and P7 is a substrate for RNase P of Escherichia coli. Proc. Natl. Acad. Sci. USA 92:5822–5826. [CrossRef]
115. Haruki, M., E. Noguchi, C. Nakai, Y. Y. Liu, M. Oobatake, M. Itaya, and S. Kanaya. 1994. Investigating the role of conserved residue Asp134 in Escherichia coli ribonuclease HI by site-directed random mutagenesis. Eur. J. Biochem. 220:623–631. [CrossRef]
116. Hayes, C. S., and R T. Sauer. 2003. Cleavage of the A site mRNA codon during ribosome pausing provides a mechanism for translational quality control. Mol. Cell 12:903–911. [CrossRef]
117. Hayes, F, and M. Vasseur. 1976. Processing of the 17-S Escherichia coli precursor RNA in the 27-S pre-ribosomal particle. Eur. J. Biochem. 61:433–442. [CrossRef]
118. Hostomsky, Z., Z. Hostomska, and D. A. Matthews. 1993. Ribonucleases H, p. 341–376. In S. M. Linn, R. S. Lloyd, and R. J. Roberts (ed.), Nucleases. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
119. Huang, H. W., and J. A. Cowan. 1994. Metallobiochemistry of the magnesium ion. Characterization of the essential metal-binding site in Escherichia coli ribonuclease H. Eur. J. Biochem. 219:253–260. [CrossRef]
120. Huang, S., and M. P. Deutscher. 1992. Sequence and transcriptional analysis of the Escherichia coli rnt gene encoding RNase T. J. Biol. Chem. 267:25609–25613.
121. Huang, H., J. Liao, and S. N. Cohen. 1998. Poly(A)- and poly(U)-specific RNA 3' tail shortening by E. coli ribonuclease E. Nature 391:99–102. [CrossRef]
122. Irie, M. 1997. RNase T1/RNase T2 family RNases, p. 101–124. In G. D’Alessio and J. F. Riordan (ed.), Ribonucleases: Structure and Functions. Academic Press, Inc., San Diego, Calif.
123. Ishii, R., O. Nureki, and S. Yokoyama. 2003. Crystal structure of the tRNA processing enzyme RNase PH from Aquifex aeolicus. J. Biol. Chem. 278:32397–32404. [CrossRef]
124. Itaya, M. 1990. Isolation and characterization of a second RNase H (RNase HII) of Escherichia coli K-12 encoded by the rnhB gene. Proc. Natl. Acad. Sci. USA 87:8587–8591. [CrossRef]
125. Itaya, M., and R. J. Crouch. 1991. A combination of RNase H (rnh) and recBCD or sbcB mutations in Escherichia coli K12 adversely affects growth. Mol. Gen. Genet. 227:424–432. [CrossRef]
126. Itaya, M., A. Omori, S. Kanaya, R. J. Crouch, T. Tanaka, and K. Kondo. 1999. Isolation of RNase H genes that are essential for growth of Bacillus subtilis 168. J. Bacteriol. 181:2118–2123.
127. Itoh, T., and J. Tomizawa. 1980. Formation of an RNA primer for initiation of replication of ColE1 DNA by ribonuclease H. Proc. Natl. Acad. Sci. USA 77:2450–2454. [CrossRef]
128. Itoh, T., and J. Tomizawa. 1982. Purification of ribonuclease H as a factor required for initiation of in vitro Co1E1 DNA replication. Nucleic Acids Res. 10:5949–5965. [CrossRef]
129. Jain, C., and J. G. Belasco. 1995. RNase E autoregulates its synthesis by controlling the degradation rate of its own mRNA in Escherichia coli: unusual sensitivity of the rne transcript to RNase E activity. Genes Dev. 9:84–96. [CrossRef]
130. Jain, C., A. Deana, and J. G. Belasco. 2002. Consequences of RNase E scarcity in Escherichia coli. Mol. Microbiol. 43:1053–1064. [CrossRef]
131. Jain, S. K., B. Pragai, and D. Apirion. 1982. A possible complex containing RNA processing enzymes. Biochem. Biophys. Res. Commun. 106:768–778. [CrossRef]
132. Jarrige, A-C., D. Bréchemier-Baey, N. Mathy, O. Duché, and C. Portier. 2002. Mutational analysis of polynucleotide phosphorylase from Escherichia coli. J. Mol. Biol. 321:397–409. [CrossRef]
133. Jarrige, A-C., N. Mathy, and C. Portier. 2001. PNPase autoregulates its expression by degrading a double-stranded structure in the pnp mRNA leader. EMBO J. 20:6845–6855. [CrossRef]
134. Jensen, K. F. 1993. The Escherichia coli K-12 "wild type" W3110 and MG1655 have an rph frameshift mutation that leads to pyrimidine starvation due to low pyrE expression levels. J. Bacteriol. 175:3401–3407.
135. Jensen, K. F., J. T. Anderson, and P. Poulsen. 1992. Overexpresion and rapid purification of the orfE/rph gene product, RNase PH of Escherichia coli. J. Biol. Chem. 267:17147–17152.
136. Jerome, L. J., T. van Biesen, and L. S. Frost. 1999. Degradation of FinP antisense RNA from F-like plasmids: the RNA-binding protein, FinO, protects FinP from ribonuclease E. J. Mol. Biol. 285:1457–1473. [CrossRef]
137. Jiang, X., A. Diwa, and J. G. Belasco. 2000. Regions of RNase E important for 5'-end-dependent RNA cleavage and autoregulated synthesis. J. Bacteriol. 182:2468–2475. [CrossRef]
138. Jones, P. G., R. A. VanBogelen, and F. C. Neidhardt. 1987. Induction of proteins in response to low temperature in Escherichia coli. J. Bacteriol. 169:2092–2095.
139. Jung, Y. H., and Y. Lee. 1997. Escherichia coli rnpB promoter mutants altered in stringent response. Biochem. Biophys. Res. Commun. 230:582–586. [CrossRef]
140. Kaberdin, V. R., Y. H. Chao, and S. Lin-Chao. 1996. RNase E cleaves at multiple sites in bubble regions of RNA I stem loops yielding products that dissociate differentially from the enzyme. J. Biol. Chem. 271:13103–13109. [CrossRef]
141. Kaberdin, V. R., A. Miczak, J. S. Jakobsen, S. Lin-Chao, K. J. McDowall, and A. von Gabain. 1998.The endoribonucleolytic N-terminal half of Escherichia coli RNase E is evolutionarily conserved in Synechocystis sp. and other bacteria but not the C-terminal half, which is sufficient for degradosome assembly. Proc. Natl. Acad. Sci. USA 95:11637–11642. [CrossRef]
142. Kaga, N., G. Umitsuki, K. Nagai, and M. Wachi. 2002. RNase G-dependent degradation of the eno mRNA encoding a glycolysis enzyme enolase in Escherichia coli. Biosci. Biotechnol. Biochem. 66:2216–2220. [CrossRef]
142a. Kamada, K., F. Hanaoka, and S. K. Burley. 2003. Crystal structure of the MazE/MazF complex: molecular bases of antidote-toxin recognition. Mol. Cell 11:875–884. [CrossRef]
143. Kanaya, S., and R. J. Crouch. 1983. DNA sequence of the gene coding for Escherichia coli ribonuclease H. J. Biol. Chem. 258:1276–1281.
144. Kanaya, S., and M. Ikehara. 1995. Functions and structures of ribonuclease H enzymes. Subcell. Biochem. 24:377–422.
145. Kanaya, S., C. Katsuda-Nakai, and M. Ikehara. 1991. Importance of the positive charge cluster in Escherichia coli ribonuclease HI for the effective binding of the substrate. J. Biol. Chem. 266:11621–11627.
146. Kanaya, S., A. Kohara, Y. Miura, A. Sekiguchi, S. Iwai, H. Inoue, E. Ohtsuka, and M. Ikehara. 1990. Identification of the amino acid residues involved in an active site of Escherichia coli ribonuclease H by site-directed mutagenesis. J. Biol. Chem. 265:4615–4621.
147. Kasai, T., R. S. Gupta, and D. Schlessinger. 1977. Exoribonucleases in wild type Escherichia coli and RNase II-deficient mutants. J. Biol. Chem. 252:8950–8956.
148. Katayanagi K, M. Ishikawa, M. Okumura, M. Ariyoshi, S. Kanaya, Y. Kawano, M. Suzuki, I. Tanaka, and K. Morikawa. 1993. Crystal structures of ribonuclease HI active site mutants from Escherichia coli. J. Biol. Chem. 268:22092–22099.
149. Katayanagi, K., M. Miyagawa, M. Matsushima, M. Ishikawa, S. Kanaya, M. Ikehara, T. Matsuzaki, and K. Morikawa. 1990. Three-dimensional structure of ribonuclease H from E. coli. Nature 347:306–309. [CrossRef]
150. Katayanagi, K., M. Miyagawa, M. Matsushima, M. Ishikawa, S. Kanaya, H. Nakamura, M. Ikehara, T. Matsuzaki, and K. Morikawa. 1992. Structural details of ribonuclease H from Escherichia coli as refined to an atomic resolution. J. Mol. Biol. 223:1029–1052. [CrossRef]
151. Katayanagi, K., M. Okumura, and K. Morikawa. 1993. Crystal structure of Escherichia coli RNase HI in complex with Mg2+ at 2.8 A resolution: proof for a single Mg(2+)-binding site. Proteins 17:337–346. [CrossRef]
152. Keck, J. L., E. R. Goedken, and S. Marqusee. 1998. Activation/attenuation model for RNase H. A one-metal mechanism with second-metal inhibition. J. Biol. Chem. 273:34128–34133. [CrossRef]
153. Kelly, K. O., and M. P. Deutscher. 1992. The presence of only one of five exoribonucleases is sufficient to support the growth of Escherichia coli. J. Bacteriol. 174:6682–6684.
154. Kelly, K. O., and M. P. Deutscher. 1992. Characterization of Escherichia coli RNase PH. J. Biol. Chem. 267:17153–17158.
155. Kelly, K. O., N. B. Reuven, Z. Li, and M. P. Deutscher. 1992. RNase PH is essential for tRNA processing and viability in RNase-deficient Escherichia coli cells. J. Biol. Chem. 267:16015–16018.
156. Kennell, D. 2002. Processing endoribonucleases and mRNA degradation in bacteria. J. Bacteriol. 184:4645–4657. [CrossRef]
157. Kharrat, A., M. J. Macias, T. J. Gibson, M. Nilges, and A. Pastore. 1995. Structure of the dsRNA binding domain of E. coli RNase III. EMBO J. 14:3572–3584.
158. King, T. C., R. Sirdeshmukh, and D. Schlessinger. 1984. RNase III cleavage is obligate for maturation but not for function of Escherichia coli pre-23S rRNA. Proc. Natl. Acad. Sci. USA 81:185–188. [CrossRef]
159. Kirsebom, L. A. 2002. RNase P RNA-mediated catalysis. Biochem. Soc. Trans. 30:1153–1158. [CrossRef]
160. Kogoma, T. 1997. Stable DNA replication: interplay between DNA replication, homologous recombination, and transcription. Microbiol. Mol. Biol. Rev. 61:212–238.
161. Kogoma, T., X. Hong, G. W Cadwell., K. G. Barnard, and T. Asai. 1993. Requirement of homologous recombination functions for viability of the Escherichia coli cell that lacks RNase HI and exonuclease V activities. Biochimie 75:89–99. [CrossRef]
162. Kole, R., and S. Altman. 1981. Properties of purified ribonuclease P from Escherichia coli. Biochemistry 20:1902–1906. [CrossRef]
163. Komine, Y., M. Kitabatake, T. Yokogawa, K. Nishikawa, and H. Inokuchi. 1994. A tRNA-like structure is present in 10Sa RNA, a small stable RNA from Escherichia coli. Proc. Natl. Acad. Sci. USA 91:9223–9227. [CrossRef]
164. Krasilnikov, A. S., X. Yang, T. Pan, and A. Mondragon. 2003. Crystal structure of the specificity domain of ribonuclease P. Nature 421:760–764. [CrossRef]
165. Kurz, J. C., and C. A. Fierke. 2000. Ribonuclease P: a ribonucleoprotein enzyme. Curr. Opin. Chem. Biol. 4:553–558. [CrossRef]
166. Kurz, J. C., S. Niranjanakumari, and C. A. Fierke. 1998. Protein component of Bacillus subtilis RNase P specifically enhances the affinity for precursor-tRNAAsp. Biochemistry 37:2393–2400. [CrossRef]
167. Lai, L., H. Yokota H, L. W. Hung, R. Kim, and S. H. Kim. 2000. Crystal structure of archaeal RNase HII: a homologue of human major RNase H. Struct. Fold. Des. 8:897–904. [CrossRef]
168. Le Derout, J., P. Regnier, and E. Hajnsdorf. 2002. Both temperature and medium composition regulate RNase E processing efficiency of the rpsO mRNA coding for ribosomal protein S15 of Escherichia coli. J. Mol. Biol. 319:341–349. [CrossRef]
169. Lee, K., J. A. Bernstein, and S. N. Cohen. 2002. RNase G complementation of rne null mutation identifies functional interrelationships with RNase E in Escherichia coli. Mol. Microbiol. 43:1445–1456. (Erratum, 46:295.)
170. Lee, K., X. Zhan, J. Gao, J. Qiu, Y. Feng, R. Meganathan, S. N. Cohen, and G. Georgiou. 2003. RraA. a protein inhibitor of RNase E activity that globally modulates RNA abundance in E. coli. Cell 114:623–634. [CrossRef]
171. Li, H. L., B. S. Chelladurai, K. Zhang, and A. W. Nicholson. 1993. Ribonuclease III cleavage of a bacteriophage T7 processing signal. Divalent cation specificity, and specific anion effects. Nucleic Acids Res. 21:1919–1925. [CrossRef]
172. Li, H., and A. W. Nicholson. 1996. Defining the enzyme binding domain of a ribonuclease III processing signal. Ethylation interference and hydroxyl radical footprinting using catalytically inactive RNase III mutants. EMBO J. 15:1421–1433.
173. Li, J., and R. M. Wartell. 1998. RNase H1 can catalyze RNA/DNA hybrid formation and cleavage with stable hairpin or duplex DNA oligomers. Biochemistry 37:5154–5161. [CrossRef]
174. Li, Y., and S. Altman. 2003. A specific endoribonuclease, RNase P, affects gene expression of polycistronic operon mRNAs. Proc. Natl. Acad. Sci. USA 100:13213–13218. [CrossRef]
175. Li, Z., and M. P. Deutscher. 1995. The tRNA processing enzyme RNase T is essential for maturation of 5S RNA. Proc. Natl. Acad. Sci. USA 92:6883–6886. [CrossRef]
176. Li, Z., and M. P. Deutscher. 1996. Maturation pathways for E. coli tRNA precursors: a random multienzyme process in vivo. Cell 86:503–512. [CrossRef]
177. Li, Z., and M. P. Deutscher. 2002. RNase E plays an essential role in the maturation of Escherichia coli tRNA precursors. RNA 8:97–109. [CrossRef]
178. Li., Z., S. Pandit, and M. P. Deutscher. 1998. 3’ exoribonucleolytic trimming is a common feature of the maturation of small, stable RNAs in Escherichia coli. Proc. Natl. Acad. Sci. USA 95:2856–2861. [CrossRef]
179. Li, Z., S. Pandit, and M. P. Deutscher. 1999. Maturation of 23S ribosomal RNA requires the exoribonuclease RNase T. RNA 5:139–146. [CrossRef]
180. Li, Z., S. Pandit, and M. P. Deutscher. 1999. RNase G (CafA protein) and RNase E are both required for the 5' maturation of 16S ribosomal RNA. EMBO J. 18:2878–2885. [CrossRef]
181. Li, Z., S. Reimers, S. Pandit, and M. P. Deutscher. 2002. RNA quality control: degradation of defective transfer RNA. EMBO J. 21:1132–1138. [CrossRef]
182. Li, Z., L. Zhan, and M. P. Deutscher. 1996. The role of individual cysteine residues in the activity of Escherichia coli RNase T. J. Biol. Chem. 271:1127–1132. [CrossRef]
183. Li, Z., L. Zhan and M. P. Deutscher. 1996. Escherichia coli RNase T functions in vivo as a dimer dependent on cysteine 168. J. Biol. Chem. 271:1133–1137. [CrossRef]
184. Lima, W. F., and S. T. Crooke. 1997. Cleavage of single strand RNA adjacent to RNA-DNA duplex regions by Escherichia coli RNase H1. J. Biol. Chem. 272:27513–27516. [CrossRef]
185. Lima, W. F., V. Mohan, and S. T. Crooke. 1997. The influence of antisense oligonucleotide-induced RNA structure on Escherichia coli RNase H1 activity. J. Biol. Chem. 272:18191–18199. [CrossRef]
186. Lin-Chao, S., and S. N. Cohen. 1991. The rate of processing and degradation of antisense RNAI regulates the replication of ColE1-type plasmids in vivo. Cell 65:1233–1242. [CrossRef]
187. Lin-Chao, S., C. L. Wei, and Y. T. Lin. 1999. RNase E is required for the maturation of ssrA RNA and normal ssrA RNA peptide-tagging activity. Proc. Natl. Acad. Sci. USA 96:12406–12411. [CrossRef]
188. Lin-Chao, S., T. T. Wong, K. J. McDowall, and S. N. Cohen. 1994. Effects of nucleotide sequence on the specificity of rne-dependent and RNase E-mediated cleavages of RNA I encoded by the pBR322 plasmid. J. Biol. Chem. 269:10797–10803.
189. Liou, G. G., W. N. Jane, S. N. Cohen, N. S. Lin, and S. Lin-Chao. 2001. RNA degradosomes exist in vivo in Escherichia coli as multicomponent complexes associated with the cytoplasmic membrane via the N-terminal region of ribonuclease E. Proc. Natl. Acad. Sci. USA 98:63–68. (Erratum, 98:8157.) [CrossRef]
190. Littauer, U. Z, and H. Soreq. 1982. Polynucleotide phosphorylase, p. 517–553. In P. D. Boyer (ed.), The Enzymes, vol. XV, part B. Academic Press, Inc., New York, N.Y.
191. Loria, A., and T. Pan. 2001. Modular construction for function of a ribonucleoprotein enzyme: the catalytic domain of Bacillus subtilis RNase P complexed with B. subtilis RNase P protein. Nucleic Acids Res. 29:1892–1897. [CrossRef]
192. Lundberg, U., and S. Altman. 1995. Processing of the precursor to the catalytic RNA subunit of RNase P from Escherichia coli. RNA 1:327–334.
193. Mackie, G. A. 1991. Specific endonucleolytic cleavage of the mRNA for ribosomal protein S20 of Escherichia coli requires the product of the ams gene in vivo and in vitro. J. Bacteriol. 173:2488–2497.
194. Mackie, G. A. 1992. Secondary structure of the mRNA for ribosomal protein S20. Implications for cleavage by ribonuclease E. J. Biol. Chem. 267:1054–1061. (Erratum, 267:8702.)
195. Mackie, G. A. 1998. Ribonuclease E is a 5'-end-dependent endonuclease. Nature 395:720–723. [CrossRef]
196. Mackie, G. A. 2000. Stabilization of circular rpsT mRNA demonstrates the 5'-end dependence of RNase E action in vivo. J. Biol. Chem. 275:25069–25072. [CrossRef]
197. Mackie, G. A., and J. L. Genereaux. 1993. The role of RNA structure in determining RNase E-dependent cleavage sites in the mRNA for ribosomal protein S20 in vitro. J. Mol. Biol. 234:998–1012. [CrossRef]
198. Maisurian, A. N., and E. A. Buyanovskaya. 1973. Isolation of an Escherichia coli strain restricting bacteriophage suppressor. Mol. Gen. Genet. 120:227–229. [CrossRef]
199. Maki, H., T. Horiuchi, and M. Sekiguchi. 1983. Structure and expression of the dnaQ mutator and the RNase H genes of Escherichia coli: overlap of the promoter regions. Proc. Natl. Acad. Sci. USA 80:7137–7141. [CrossRef]
200. March, P. E., J. Ahnn, and M. Inouye. 1985. The DNA sequence of the gene (rnc) encoding ribonuclease III of Escherichia coli. Nucleic Acids Res. 13:4677–4685. [CrossRef]
201. March, P. E., and M. A. Gonzalez. 1990. Characterization of the biochemical properties of recombinant ribonuclease III. Nucleic Acids Res. 18:3293–3298. [CrossRef]
202. Marianovsky, I., E. Aizenman, H. Engelberg-Kulka, and G. Glaser. 2001. The regulation of the Escherichia coli mazEF promoter involves an unusual alternating palindrome. J. Biol. Chem. 276:5975–5984. [CrossRef]
203. Marujo, P. E., E. Hajnsdorf, J. Le Derout, R. Andrade, C. M. Arraiano, and P. Regnier. 2000. RNase II removes the oligo(A) tails that destabilize the rpsO mRNA of Escherichia coli. RNA 6:1185–1193. [CrossRef]
204. Masse, E., F. E. Escorcia, and S. Gottesman. 2003. Coupled degradation of a small regulatory RNA and its mRNA targets in Escherichia coli. Genes Dev. 17:2374–2383. [CrossRef]
204a. Massire C., L. Jaeger, and E. Westhof. 1998. Derivation of the three-dimensional architecture of bacterial ribonuclease P RNAs from comparative sequence analysis. J. Mol. Biol. 279:773–ss793. [CrossRef]
205. Mathy, N., A.-C. Jarrige, M. Robert-LeMeur, and C. Portier. 2001. Increased expression of Escherichia coli polynucleotide phosphorylase at low temperatures is linked to a decrease in the efficiency of autocontrol. J. Bacteriol. 183:3848–3854.[PubMed] [CrossRef]
206. Matsunaga, J., M. Dyer, E. L. Simons, and R. W. Simons. 1996. Expression and regulation of the rnc and pdxJ operons of Escherichia coli. Mol. Microbiol. 22:977–989. [CrossRef]
207. Matsunaga, J., E. L. Simons, and R. W. Simons. 1996. RNase III autoregulation: structure and function of rncO, the posttranscriptional "operator". RNA 2:1228–1240.
208. Matsunaga, J., E. L. Simons, and R. W. Simons. 1997. Escherichia coli RNase III (rnc) autoregulation occurs independently of rnc gene translation. Mol. Microbiol. 26:1125–1135. [CrossRef]
209. Mattatall, N. R., and K. E. Sanderson. 1998. RNase III deficient Salmonella typhimurium LT2 contains intervening sequences (IVSs) in its 23S rRNA. FEMS Microbiol. Lett. 159:179–185. [CrossRef]
210. Mayer, J. E., and M. Schweiger. 1983. RNase III is positively regulated by T7 protein kinase. J. Biol. Chem. 258:5340–5343.
211. McClain, W. H., C. Guerrier-Takada, and S. Altman. 1987. Model substrates for an RNA enzyme. Science 238:527–530. [CrossRef]
212. McDowall, K. J., and S. N. Cohen. 1996. The N-terminal domain of the rne gene product has RNase E activity and is non-overlapping with the arginine-rich RNA-binding site. J. Mol. Biol. 255:349–355. [CrossRef]
213. McDowall, K. J., R. G. Hernandez, S. Lin-Chao, and S. N. Cohen. 1993. The ams-1 and rne-3071 temperature-sensitive mutations in the ams gene are in close proximity to each other and cause substitutions within a domain that resembles a product of the Escherichia coli mre locus. J. Bacteriol. 175:4245–4249.
214. McDowall, K. J., V. R. Kaberdin, S. W. Wu, S. N. Cohen, and S. Lin-Chao. 1995. Site-specific RNase E cleavage of oligonucleotides and inhibition by stem-loops. Nature 374:287–290. [CrossRef]
215. McDowall, K. J., S. Lin-Chao, and S. N. Cohen. 1994. A+U content rather than a particular nucleotide order determines the specificity of RNase E cleavage. J. Biol. Chem. 269:10790–10796.
216. McLaren, R. S., S. F. Newbury, G. S. Dance, H. C. Causton, and C. F. Higgins. 1991. mRNA degradation by processive 3’-5’exoribonucleases in vitro and the implications for prokaryotic mRNA decay in vivo. J. Mol. Biol. 221:81–95. [CrossRef]
217. McMurry, L. M., and S. B. Levy. 1987. Tn5 insertion in the polynucleotide phosphorylase (pnp) gene in Escherichia coli increases susceptibility to antibiotics. J. Bacteriol. 169:1321–1324.
218. Meador, J., III, B. Cannon, V. J. Cannistraro, and D. Kennell. 1990. Purification and characterization of Escherichia coli RNase I. Comparisons with RNase M. Eur. J. Biochem. 187:549–553. [CrossRef]
219. Meader, J., III, and D. Kennell. 1990. Cloning and sequencing the gene encoding Escherichia coli ribonuclease I: exact physical mapping using the genome library. Gene 95:1–7. [CrossRef]
220. Mian, I. S. 1997. Comparative sequence analysis of ribonucleases HII, III, II, PH and D. Nucleic Acids Res. 25:3187–3195. [CrossRef]
221. Miczak, A., V. R. Kaberdin, C. L. Wei, and S. Lin-Chao. Proc. Natl. Acad. Sci USA 93:3865–3869. [CrossRef]
222. Minasov, G., M. Teplova, P. Nielsen, J. Wengel, and M. Egli. 2000. Structural basis of cleavage by RNase H of hybrids of arabinonucleic acids and RNA. Biochemistry 39:3525–3532. [CrossRef]
223. Misra, T. K., and D. Apirion. 1979. RNase E, an RNA processing enzyme from Escherichia coli. J. Biol. Chem. 254:11154–11159.
224. Mohanty, B. K., and S. R. Kushner. 2000. Polynucleotide phosphorylase, RNase II and RNase E play different roles in the in vivo modulation of polyadenylation in Escherichia coli. Mol. Microbiol. 36:982–994. [CrossRef]
225. Mohanty, B. K., and S. R. Kushner. 2000. Polynucleotide phosphorylase functions both as a 3’→5’ exonuclease and a poly(A) polymerase in Escherichia coli. Proc. Natl. Acad. Sci. USA 97:11966–11971. [CrossRef]
226. Mohanty, B. K., and S. R. Kushner. 2002. Polyadenylation of Escherichia coli transcripts plays an integral role in regulating intracellular levels of polynucleotide phosphorylase and RNase E. Mol. Microbiol. 45:1315–1324. [CrossRef]
227. Mohanty, B. K., and S. R. Kushner. 2003. Genomic analysis in Escherichia coli demonstrates differential roles for polynucleotide phosphorylase and RNase II in mRNA abundance and decay. Mol. Microbiol. 50:645–658. [CrossRef]
228. Moll, I., Afonyushkin T., O. Vytvytska, V. R. Kaberdin, and U. Blasi. 2003. Coincident Hfq binding and RNase E cleavage sites on mRNA and small regulatory RNAs. RNA 9:1308–1314. [CrossRef]
228a. Motamedi, H., Y. Lee, and F. J. Schmidt. 1984. Tandem promoters preceding the gene for the M1 RNA component of Escherichia coli ribonuclease P. Proc. Natl. Acad. Sci. USA 81:3959–3963. [CrossRef]
229. Mudd, E. A., and C. F. Higgins. 1993. Escherichia coli endoribonuclease RNase E: autoregulation of expression and site-specific cleavage of mRNA. Mol. Microbiol. 9:557–568. [CrossRef]
230. Mudd, E. A., H. M. Krisch, and C. F. Higgins. 1990. RNase E, an endoribonuclease, has a general role in the chemical decay of Escherichia coli mRNA: evidence that rne and ams are the same genetic locus. Mol. Microbiol. 4:2127–2135. [PubMed] [CrossRef]
231. Muroya, A., D. Tsuchiya, M. Ishikawa, M. Haruki, M. Morikawa, S. Kanaya, and K. Morikawa. 2001. Catalytic center of an archaeal type 2 ribonuclease H as revealed by X-ray crystallographic and mutational analyses. Protein Sci. 10:707–714. [CrossRef]
232. Nakamura, H., Y. Oda, S. Iwai, H. Inoue, E. Ohtsuka, S. Kanaya, S. Kimura, C. Katsuda, K. Katayanagi, K. Morikawa, H. Miyashiro, and M. Ikehara. 1991. How does RNase H recognize a DNA.RNA hybrid? Proc. Natl. Acad. Sci. USA 88:11535–11539. [CrossRef]
233. Nanbu-Wakao, R., S. Asoh, K. Nishimaki, K. Tanaka, and S. Ohta. 2000. Bacterial cell death induced by human pro-apoptotic Bax is blocked by an RNase E mutant that functions in an anti-oxidant pathway. Genes Cells 5:155–167. [CrossRef]
234. Nashimoto, H., A. Miura, H. Saito, and H. Uchida. 1985. Suppressors of temperature-sensitive mutations in a ribosomal protein gene, rpsL (S12), of Escherichia coli K12. Mol. Gen. Genet. 199:381–387. [CrossRef]
235. Nashimoto, H., and H. Uchida. 1985. DNA sequencing of the Escherichia coli ribonuclease III gene and its mutations. Mol. Gen. Genet. 201:25–29. [CrossRef]
236. Naureckiene, S., and B. E. Uhlin. 1996. In vitro analysis of mRNA processing by RNase E in the pap operon of Escherichia coli. Mol. Microbiol. 21:55–68. [CrossRef]
237. Nicholson, A. W. 1996. Structure, reactivity, and biology of double-stranded RNA. Prog. Nucleic Acids Res. Mol. Biol. 52:1–65. [CrossRef]
238. Nicholson, A. W. 1997. Escherichia coli ribonucleases: paradigms for understanding cellular RNA metabolism and regulation, p. 1–49. In G. D'Alessio & J. F. Riordan (ed.), Ribonucleases: Structures and Functions. Academic Press, Inc., New York, N.Y.
239. Nicholson, A. W. 1999. Function, mechanism and regulation of bacterial ribonucleases. FEMS Microbiol. Rev. 23:371–390. [CrossRef]
240. Niranjanakumari, S., T. Stams, S. M. Crary, D. W. Christianson, and C. A. Fierke. 1998. Protein component of the ribozyme ribonuclease P alters substrate recognition by directly contacting precursor tRNA. Proc. Natl. Acad. Sci. USA 95:15212–15217. [CrossRef]
241. Niyogi, S. K., and A. K. Datta. 1975. A novel oligoribonuclease of Escherichia coli. I. Isolation and properties. J. Biol. Chem. 250:7307–7312.
242. Nomura, T., and A. Ishihama. 1988. A novel function of RNase P from Escherichia coli: processing of a suppressor tRNA precursor. EMBO J. 7:3539–3545.
243. Oda, Y., S. Iwai, E. Ohtsuka, M. Ishikawa, M. Ikehara, and H. Nakamura. 1993a.Binding of nucleic acids to E. coli RNase HI observed by NMR and CD spectroscopy. Nucleic Acids Res. 21:4690–4695. [CrossRef]
244. Oda, Y., H. Nakamura, S. Kanaya, and M. Ikehara. 1991. Binding of metal ions to E. coli RNase HI observed by 1H-15N heteronuclear 2D NMR. J. Biomol. NMR 1:247–255. [CrossRef]
245. Oda, Y., M. Yoshida, and S. Kanaya. 1993. Role of histidine 124 in the catalytic function of ribonuclease HI from Escherichia coli. J. Biol. Chem. 268:88–92.
246. Ogawa, T., and T. Okazaki. 1984. Function of RNase H in DNA replication revealed by RNase H defective mutants of Escherichia coli. Mol. Gen. Genet. 193:231–237. [CrossRef]
247. Ohtani, N., M. Haruki, M. Morikawa, R. J. Crouch, M. Itaya, and S. Kanaya. 1999. Identification of the genes encoding Mn2+-dependent RNase HII and Mg2+-dependent RNase HIII from Bacillus subtilis: classification of RNases H into three families. Biochemistry 38:605–618. [CrossRef]
248. Ohtani, N., M. Haruki, A. Muroya, M. Morikawa, and S. Kanaya. 2000. Characterization of ribonuclease HII from Escherichia coli overproduced in a soluble form. J. Biochem. (Tokyo) 127:895–899.[PubMed]
249. Okada, Y., M. Wachi, A. Hirata, K. Suzuki, K. Nagai, and M. Matsuhashi. 1994. Cytoplasmic axial filaments in Escherichia coli cells: possible function in the mechanism of chromosome segregation and cell division. J. Bacteriol. 176:917–922.
250. Ono, M., and M. Kuwano. 1980. Chromosomal location of a gene for chemical longevity of messenger ribonculeic acid in a temperature-sensitive mutant of Escherichia coli. J. Bacteriol. 142:325–326.
251. Ost, K. A., and M. P. Deutscher. 1990. RNase PH catalyzes a synthetic reaction, the addition of nucleotides to the 3’ end of RNA. Biochimie 72:813–818. [CrossRef]
252. Ost, K. A., and M. P. Deutscher. 1991. Escherichia coli orfE (upstream of pyrE) encodes RNase PH. J. Bacteriol. 173:5589–5591.
253. Ow, M. C., and S. R. Kushner. 2002. Initiation of tRNA maturation by RNase E is essential for cell viability in E. coli. Genes Dev. 16:1102–1115. [CrossRef]
254. Ow, M. C., Q. Liu, B. K. Mohanty, M. E. Andrew, V. F. Maples, and S. R. Kushner. 2002. RNase E levels in Escherichia coli are controlled by a complex regulatory system that involves transcription of the rne gene from three promoters. Mol. Microbiol. 43:159–171. [CrossRef]
255. Ow, M. C., T. Perwez, and S. R. Kushner. 2003. RNase G of Escherichia coli exhibits only limited functional overlap with its essential homologue, RNase E. Mol. Microbiol. 49:607–622. [CrossRef]
255a. Pace, N. R., and J. W. Brown. 1995. Evolutionary perspective on the structure and function of ribonuclease P, a ribozyme. J. Bacteriol. 177:1919–1928.
256. Pace, N. R., and D. Smith. 1990. Ribonuclease P: function and variation. J. Biol. Chem. 265:3587–3590.[PubMed]
257. Padmanabha, K. P., and M. P. Deutscher. 1991. RNase T affects Escherichia coli growth and recovery from metabolic stress. J. Bacteriol. 173:1376–1381.
258. Padmanabham, S., Z. Zhou, C. Y. Chu, R. W. Lim, and L. W. Lim. 2001. Overexpression, biophysical characterization and crystallization of ribonuclease I from Escherichia coli, a broad-specificity enzyme in the RNase T2 family. Arch. Biochem. Biophys. 390:42–50. [CrossRef]
259. Park, J. W., Y. Jung, S. J. Lee, D. J. Jin, Y. Lee, and S. J. Lee. 2002. Alteration of stringent response of the Escherichia coli rnpB promoter by mutations in the -35 region. Biochem. Biophys. Res. Commun. 290:1183–1187. [CrossRef]
260. Peck-Miller, K. A., and S. Altman. 1991. Kinetics of the processing of the precursor to 4.5 S RNA, a naturally occurring substrate for RNase P from Escherichia coli. J. Mol. Biol. 221:1–5. [CrossRef]
261. Pedersen, K., S. K. Christensen, and K. Gerdes. 2002. Rapid induction and reversal of a bacteriostatic condition by controlled expression of toxins and antitoxins. Mol. Microbiol. 45:501–510. [CrossRef]
262. Pedersen, K., A. V. Zavialov, M. Yu. Pavlov, J. Elf, K. Gerdes, and M. Ehrenberg. 2003. The bacterial toxin RelE displays codon-specific cleavage of mRNAs in the ribosomal A site. Cell 112:131–140. [CrossRef]
263. Plautz, G., and D. Apirion. 1981. Processing of RNA in Escherichia coli is limited in the absence of ribonuclease III, ribonuclease E and ribonuclease P. J. Mol. Biol. 149:813–819. [CrossRef]
264. Portier, C. 1975. Quaternary structure of Escherichia coli polynucleotide phosphorylase: new evidence for a trimeric structure. FEBS Lett. 50:79–81. [CrossRef]
265. Portier, C. 1975. Quanternary structure of polynucleotide phosphorylase from Escherichia coli: evidence of a complex between two types of polypeptide chains. Eur. J. Biochem. 55:573–582. [CrossRef]
266. Portier, C., L. Dondon, M. Grunberg-Manago, and P. Regnier. 1987. The first step in the functional inactivation of the Escherichia coli polynucleotide phosphorylase messenger is a ribonuclease II processing at the 5’ end. EMBO J. 6:2165–2170.
267. Poulson, P., F. Bonekamp, and K. F. Jensen. 1984. Structure of the Escherichia coli pyrE operon and control of pyrE expression by a UTP modulated intercistronic attenuation. EMBO J. 3:1783–1790.
268. Poulsen, P., K. F. Jensen, P. Valentin-Hansen, P. Carlsson, and L. G. Lundberg. 1983. Nucleotide sequence of the Escherichia coli pyrE gene and the DNA in front of the protein-coding region. Eur. J. Biochem. 135:223–229. [CrossRef]
269. Py, B., C. F. Higgins, H. M. Krisch, and A. J. Carpousis. 1996. A DEAD-box RNA helicase in the Escherichia coli RNA degradosome. Nature 381:169–172. [CrossRef]
270. Qin, H., T. R. Sosnick, and T. Pan. 2001. Modular construction of a tertiary RNA structure: the specificity domain of the Bacillus subtilis RNase P RNA. Biochemistry 40:11202–11210. [CrossRef]
271. Quinones, A., C. Kucherer, R. Piechocki, and W. Messer. 1987. Reduced transcription of the rnh gene in Escherichia coli mutants expressing the SOS regulon constitutively. Mol. Gen. Genet. 206:95–100. [CrossRef]
272. Regnier, P., and M. Grunberg-Manago. 1989. Cleavage by RNase III in the transcripts of the met Y-nus-A-infB operon of Escherichia coli releases the tRNA and initiates the decay of the downstream mRNA. J. Mol. Biol. 210:293–302. [CrossRef]
273. Regnier, P., and M. Grunberg-Manago. 1990. RNase III cleavages in non-coding leaders of Escherichia coli transcripts control mRNA stability and genetic expression. Biochimie 72:825–834. [CrossRef]
274. Regnier, P., M. Grunberg-Manago, and C. Portier. 1987. Nucleotide sequence of the pnp gene of Escherichia coli encoding polynucleotide phosphorylase. Homology of the primary structure of the protein with the RNA-binding domain of ribosomal protein S1. J. Biol. Chem. 262:63–68.
275. Reuven, N. B., and M. P. Deutscher. 1993. Multiple exoribonucleases are required for the 3’ processing of Escherichia coli tRNA precursors in vivo. FASEB J. 7:143–148.
276. Reuven, N. B., E. V. Koonin, K. E. Rudd, and M. P. Deutscher. 1995. The gene for the longest known Escherichia coli protein is a member of helicase superfamily II. J. Bacteriol. 177:5393–5400.
277. Reuven, N. B., Z. Zhou, and M. P. Deutscher. 1997. Functional overlap of tRNA nucleotidyltransferase, poly(A) polymerase I, and polynucleotide phosphorylase. J. Biol. Chem. 272:33255–33259. [CrossRef]
278. Robertson, H. D. 1982. Escherichia coli ribonuclease III cleavage sites. Cell 30:669–672. [CrossRef]
279. Robertson, H. D., R. E. Webster, and N. D. Zinder. 1968. Purification and properties of ribonuclease III from Escherichia coli. J. Biol. Chem. 243:82–91.
280. Robertson, H. D., S. Altman, and J. D. Smith. 1972. Purification and properties of a specific Escherichia coli ribonuclease which cleaves a tyrosine transfer ribonucleic acid presursor. J. Biol. Chem. 247:5243–5251.
281. Rotondo, G., and D. Frendewey. 1996. Purification and characterization of the Pac1 ribonuclease of Schizosaccharomyces pombe. Nucleic Acids Res. 24:2377–2386. [CrossRef]
282. Roy, P., Cudny, H. and M. P. Deutscher. 1982. The transfer RNA processing defect in Escherichia coli strains BN and CAN is not due to a mutation in RNase D or RNase II. J. Mol. Biol. 159:179–187. [CrossRef]
283. Rudd, K. E. 1998. Linkage map of Escherichia coli K-12, edition 10: the physical map. Microbiol. Mol. Biol. Rev. 62:985–1019.
284. Rydberg, B., and J. Game. 2002. Excision of misincorporated ribonucleotides in DNA by RNase H (type 2) and FEN-1 in cell-free extracts. Proc. Natl. Acad. Sci. USA 99:16654–16659. [CrossRef]
285. Ryter, J.M., and S.C. Schultz. 1998. Molecular basis of double-stranded RNA-protein interactions: strsssucture of a dsRNA-binding domain complexed with dsRNA. EMBO J. 17:7505–7513. [CrossRef]
285a. Sakamoto, H., N. Kimura, and Y. Shimura. 1983. Processing of transcription products of the gene encoding the RNA component of RNase P. Proc. Natl. Acad. Sci. USA 80:6187–6191. [CrossRef]
286. Sakano, H., and Y. Shimura. 1978. Characterization and in vitro processing of transfer RNA precursors accumulated in a temperature-sensitive mutant of Escherichia coli. J. Mol. Biol. 123:287–326.[PubMed] [CrossRef]
286a. Sat, B., R. Hazan, T. Fisher, H. Khaner, G. Glaser, and H. Engelberg-Kulka. 2001. Programmed cell death in Escherichia coli: some antibiotics can trigger mazEF lethality. J Bacteriol. 183:2041–2045. [CrossRef]
287. Schedl, P., and P. Primakoff. 1973. Mutants of Escherichia coli thermosensitive for the synthesis of transfer RNA. Proc. Natl. Acad. Sci. USA 70:2091–2095.[PubMed] [CrossRef]
288. Schedl, P., J. Roberts, and P. Primakoff. 1976. In vitro processing of E. coli tRNA precursors. Cell 8:581–594. [CrossRef]
289. Schiffer, S., S. Rosch, and A. Marchfelder. 2002. Assigning a function to a conserved group of proteins: the tRNA 3'-processing enzymes. EMBO J. 21:2769–2777. [CrossRef]
290. Schmidt, M., and N. Delihas. 1995. micF RNA is a substrate for RNase E. FEMS Microbiol. Lett. 133:209–213. [CrossRef]
291. Seidman, J. G., F. J. Schmidt, K. Foss, and W. H. McClain. 1975. A mutant of Escherichia coli defective in removing 3’ terminal nucleotides from some transfer RNA precursor molecules. Cell 5:389–400. [CrossRef]
292. Sharkady, S. M., and J. M. Nolan. 2001. Bacterial ribonuclease P holoenzyme crosslinking analysis reveals protein interaction sites on the RNA subunit. Nucleic Acids Res. 29:3848–3856. [CrossRef]
293. Shen, V., and D. Schlessinger. 1982. RNase I, II and IV of Escherichia coli. p. 501–515. In P. D. Boyer (ed.), The Enzymes, vol. XV, part B. Academic Press, Inc., New York, N.Y.
294. Sim, S., K. S. Kim, and Y. Lee. 2002. 3'-end processing of precursor M1 RNA by the N-terminal half of RNase E. FEBS Lett. 529:225–231. [CrossRef]
295. Sirdeshmukh, R., and D. Schlessinger. 1985. Ordered processing of Escherichia coli 23S rRNA in vitro. Nucleic Acids Res. 13:5041–5054. [CrossRef]
296. Soderbom, F., U. Binnie, M. Masters, and E. G. Wagner. 1997. Regulation of plasmid R1 replication: PcnB and RNase E expedite the decay of the antisense RNA, CopA. Mol. Microbiol. 26:493–504. [CrossRef]
296a. Sousa, S., I. Marchand, and M. Dreyfus. 2001. Autoregulation allows Escherichia coli RNase E to adjust continuously its synthesis to that of its substrates. Mol. Microbiol. 42:867–878. [CrossRef]
297. Spahr, P. F., and R. F. Gesteland. 1968. Specific cleavage of bacteriophage R17 RNA by an endonuclease isolated from E. coli MRE-600. Proc. Natl. Acad. Sci. USA 59:876–883.[PubMed] [CrossRef]
298. Spickler, C., and G. A. Mackie. 2000. Action of RNase II and polynucleotide phosphorylase against RNAs containing stem-loops of defined structure. J. Bacteriol. 182:2422–2427. [CrossRef]
299. Spitzfaden, C., N. Nicholson, J. J. Jones, S. Guth, R. Lehr, C. D. Prescott, L. A. Hegg, and D. S. Eggleston. 2000. The structure of ribonuclease P protein from Staphylococcus aureus reveals a unique binding site for single-stranded RNA. J. Mol. Biol. 295:105–115. [CrossRef]
300. Srivastava, A. K., and D. Schlessinger. 1988. Coregulation of processing and translation: mature 5' termini of Escherichia coli 23S ribosomal RNA form in polysomes. Proc. Natl. Acad. Sci. USA 85:7144–7148. [CrossRef]
301. Srivastava, A. K., and D. Schlessinger. 1990. Mechanism and regulation of bacterial ribosomal RNA processing. Annu. Rev. Microbiol. 44:105–129. [CrossRef]
302. Srivastava, S. K., V. J. Cannistraro, and D. Kennell. 1992. Broad-specificity endoribonucleases and mRNA degradation in Escherichia coli. J. Bacteriol. 174:56–62.
303. St. Johnston, D., N. H. Brown, J. G. Gall, and M. Jantsch. 1992. A conserved double-stranded RNA-binding domain. Proc. Natl. Acad. Sci. USA 89:10979–10983. [CrossRef]
304. Stams, T., S. Niranjanakumari, C. A. Fierke, and D. W. Christianson. 1998. Ribonuclease P protein structure: evolutionary origins in the translational apparatus. Science 280:752–755. [CrossRef]
305. Stark, B. C., R. Kole, E. J. Bowman, and S. Altman. 1978. Ribonuclease P: an enzyme with an essential RNA component. Proc. Natl. Acad. Sci. USA 75:3717–3721. [CrossRef]
306. Steitz, T. A., and J. A. Steitz. 1993. A general two-metal-ion mechanism for catalytic RNA. Proc. Natl. Acad. Sci. USA 90:6498–6502. [CrossRef]
307. Subbarayan, P. R., and M. P. Deutscher. 2001. Escherichia coli RNase M is a multiply altered form of RNase I. RNA 7:1702–1707.
308. Sun, W., E. Jun, and A. W. Nicholson. 2001. Intrinsic double-stranded-RNA processing activity of Escherichia coli ribonuclease III lacking the dsRNA-binding domain. Biochemistry 40:14976–14984. [CrossRef]
309. Symmons, M. F., M. G. Williams, B. F. Luisi, G. H. Jones, and A. J. Carpousis. 2002. Running rings around RNA: a superfamily of phosphate-dependent RNases. Trends Biochem. Sci. 27:11–18. [CrossRef]
310. Takiff, H. E., S. M. Chen, and D. L. Court. 1989. Genetic analysis of the rnc operon of Escherichia coli. J. Bacteriol. 171:2581–2590.
311. Talbot, S. J., and S. Altman. 1994. Gel retardation analysis of the interaction between C5 protein and M1 RNA in the formation of the ribonuclease P holoenzyme from Escherichia coli. Biochemistry 33:1399–1405. [CrossRef]
312. Talkad, V., D. Achord, and D. Kennell. 1978. Altered mRNA metabolism in ribonuclease III-deficient strains of Escherichia coli. J. Bacteriol. 135:528–541.
313. Taraseviciene, L., G. R. Bjork, and B. E. Uhlin. 1995. Evidence for an RNA binding region in the Escherichia coli processing endoribonuclease RNase E. J. Biol. Chem. 270:26391–26398. [CrossRef]
314. Tian, B., and M. B. Mathews. 2003. Phylogenetics and functions of the double-stranded RNA-binding motif: a genomic survey. Prog. Nucleic Acids Res. Mol. Biol. 74:123–158. [CrossRef]
315. Tock, M. R., A. P. Walsh, G. Carroll, and K. J. McDowall. 2000. The CafA protein required for the 5'-maturation of 16S rRNA is a 5'-end-dependent ribonuclease that has context-dependent broad sequence specificity. J. Biol. Chem. 275:8726–8732. [CrossRef]
316. Tsai, H. Y., B. Masquida, R. Biswas, E. Westhof, and V. Gopalan. 2003. Molecular modeling of the three-dimensional structure of the bacterial RNase P holoenzyme. J. Mol. Biol. 325:661–675. [CrossRef]
317. Tsunaka, Y., M. Haruki, M. Morikawa, M. Oobatake, and S. Kanaya. 2003. Dispensability of glutamic acid 48 and aspartic acid 134 for Mn2+-dependent activity of Escherichia coli ribonuclease HI. Biochemistry 42:3366–3374. [CrossRef]
318. Umitsuki, G., M. Wachi, A. Takada, T. Hikichi, and K. Nagai. 2001. Involvement of RNase G in in vivo mRNA metabolism in Escherichia coli. Genes Cells 6:403–410. [CrossRef]
319. Vanzo, N. F., Y. S. Li, B. Py, E. Blum, C. F. Higgins, L. C. Raynal, H. M. Krisch, and A. J. Carpousis. 1998. Ribonuclease E organizes the protein interactions in the Escherichia coli RNA degradosome. Genes Dev. 12:2770–2781. [CrossRef]
320. Vioque, A., J. Arnez, and S. Altman. 1988. Protein-RNA interactions in the RNase P holoenzyme from Escherichia coli. J. Mol. Biol. 202:835–848. [CrossRef]
321. Viswanathan, M., K. W. Dower, and S. T. Lovett. 1998. Identification of a potent DNase activity associated with RNase T of Escherichia coli. J. Biol. Chem. 273:35126–35131. [CrossRef]
322. Viswanathan, M., A. Lanjuin, and S. T. Lovett. 1999. Identification of RNase T as a high-copy suppressor of the UV sensitivity associated with single-strand DNA exonuclease deficiency in Escherichia coli. Genetics 151:929–934.
323. von Meyenburg, K., E. Boye, K. Skarstad, L. Koppes, and T. Kogoma. 1987. Mode of initiation of constitutive stable DNA replication in RNase H-defective mutants of Escherichia coli K-12. J. Bacteriol. 169:2650–2658.
324. Wachi, M., M. Doi, T. Ueda, M. Ueki, K. Tsuritani, K. Nagai, and M. Matsuhashi. 1991. Sequence of the downstream flanking region of the shape-determining genes mreBCD of Escherichia coli. Gene 106:135–136. [CrossRef]
325. Wachi, M., N. Kaga, G. Umitsuki, D. P. Clark, and K. Nagai. 2001. A novel RNase G mutant that is defective in degradation of adhE mRNA but proficient in the processing of 16S rRNA precursor. Biochem. Biophys. Res. Commun. 289:1301–1306. [CrossRef]
326. Wachi, M., G. Umitsuki, and K. Nagai. 1997. Functional relationship between Escherichia coli RNase E and the CafA protein. Mol. Gen. Genet. 253:515–519. [CrossRef]
327. Wachi, M., G. Umitsuki, M. Shimizu, A. Takada, and K. Nagai. 1999. Escherichia coli cafA gene encodes a novel RNase, designated as RNase G, involved in processing of the 5' end of 16S rRNA. Biochem. Biophys. Res. Commun. 259:483–488. [CrossRef]
328. Wehr, C. T. 1973. Isolation and properties of a ribonuclease-deficient mutant of Salmonella typhimurium. J. Bacteriol. 114:96–102.
329. Westhof, E., and S. Altman. 1994. Three-dimensional working model of M1 RNA, the catalytic RNA subunit of ribonuclease P from Escherichia coli. Proc. Natl. Acad. Sci. USA 91:5133–5137. [CrossRef]
330. Wilson, H. R., D. Yu, H. K. Peters III, J. G. Zhou, and D. L. Court. 2002. The global regulator RNase III modulates translation repression by the transcription elongation factor N. EMBO J. 21:4154–4161. [CrossRef]
331. Xu, F., and S. N. Cohen. 1995. RNA degradation in Escherichia coli regulated by 3’ adenylation and 5’ phosphorylation. Nature 374:180–183. [CrossRef]
332. Yamanaka, K., and M. Inouye. 2001. Selective mRNA degradation by polynucleotide phosphorylase in cold shock adaptation in Escherichia coli. J. Bacteriol. 183:2808–2816. [CrossRef]
333. Yang, W., W. A. Hendrickson, R. J. Crouch, and Y. Satow. 1990. Structure of ribonuclease H phased at 2 A resolution by MAD analysis of the selenomethionyl protein. Science 249:1398–1405. [CrossRef]
334. Yazbeck, D. R., K. L. Min, and M. J. Damha. 2002. Molecular requirements for degradation of a modified sense RNA strand by Escherichia coli ribonuclease H1. Nucleic Acids Res. 30:3015–3025. [CrossRef]
335. Young, R. A., and J. A. Steitz. 1978. Complementary sequences 1700 nucleotides apart form a ribonuclease III cleavage site in Escherichia coli ribosomal precursor RNA. Proc. Natl. Acad. Sci. USA 75:3593–3597. [CrossRef]
336. Yu, D., and M. P. Deutscher. 1995. Oligoribonuclease is distinct from the other known exoribonucleases of Escherichia coli. J. Bacteriol. 177:4137–4139.
337. Zaniewski, R., and M. P. Deutscher. 1982. Genetic mapping of a mutation in Escherichia coli leading to a temperature-sensitive RNase D. Mol. Gen. Genet. 185:142–147. [CrossRef]
338. Zhang, J., and M. P. Deutscher. 1988. Escherichia coli RNase D: sequencing of the rnd structural gene and purification of the overexpressed protein. Nucleic Acids Res. 16:6265–6278. [CrossRef]
339. Zhang, J., and M. P. Deutscher. 1988. Transfer RNA is a substrate for RNase D in vivo. J. Biol. Chem. 263:17909–17912.
340. Zhang, J., and M. P. Deutscher. 1988. Cloning, characterization, and effects of overexpression of the Escherichia coli rnd gene encoding RNase D. J. Bacteriol. 170:522–527.
341. Zhang, J., and M. P. Deutscher. 1989. Analysis of the upstream region of the Escherichia coli rnd gene encoding RNase D. Evidence for translational regulation of a putative tRNA processing enzyme. J. Biol. Chem. 264:18228–18233.
342. Zhang, J., and M. P. Deutscher. 1992. A uridine-rich sequence required for translation of prokaryotic mRNA. Proc. Natl. Acad. Sci. USA 89:2605–2609. [CrossRef]
343. Zhang, K., and A. W. Nicholson. 1997. Regulation of ribonuclease III processing by double-helical sequence antideterminants. Proc. Natl. Acad. Sci. USA 94:13437–13441. [CrossRef]
344. Zhang, Y., J. Zhang, K. P. Hoeflich, M. Ikura, G. Qing, and M. Inouye. 2003. MazF cleaves cellular mRNA specifically at ACA to block protein synthesis in Eschericha coli. Mol. Cell. 12:913–923. [CrossRef]
345. Zhang, X., L. Zhu, and M. P. Deutscher. 1998. Oligoribonuclease is encoded by a highly conserved gene in the 3’-5’ exonuclease superfamily. J. Bacteriol. 180:2779–2781.
346. Zhou, Z. and M. P. Deutscher. 1997. An essential function for the phosphate-dependent exoribonucleases RNase PH and polynucleotide phosphorylase. J. Bacteriol. 179:4391–4395.
347. Zhu, L., and M. P. Deutscher. 1992. The Escherichia coli rna gene encoding RNase I: Sequence and unusual promoter structure. Gene 119:101–106. [CrossRef]
348. Zhu, L., T. Gangopadhyay, K. P. Padmanabha, and M. P. Deutscher. 1990. Escherichia coli rna gene encoding RNase I: Cloning, overexpression, subcellular distribution of the enzyme, and use of an rna deletion to identify additional RNases. J. Bacteriol. 172:3146–3151.
349. Zilhao, R., L. Camelo, and C. M. Arraiano. 1993. DNA sequencing and expression of the gene rnb encoding Escherichia coli ribonuclease II. Mol. Microbiol. 8:43–51. [CrossRef]
350. Zilhao, R., J. Plumbridge, E. Hajnsdorf, P. Regnier, and C. M. Arraiano. 1996. Escherichia coli RNase II: characterization of the promoters involved in the transcription of rnb. Microbiology 142:367–375. [CrossRef]
351. Zilhao, R., P. Regnier, and C. M. Arraiano. 1995. The role of endonucleases in the expression of ribonuclease II in Escherichia coli. FEMS Microbiol. Lett. 130:237–244. [CrossRef]
352. Zuo, Y., and M. P. Deutscher. 1999. The DNase activity of RNase T and its application to DNA cloning. Nucleic Acids Res. 27:4077–4082. [CrossRef]
353. Zuo, Y., and M. P. Deutscher. 2001. Exoribonuclease superfamilies: structural analysis and phylogenetic distribution. Nucleic Acids Res. 29:1017–1026. [CrossRef]
354. Zuo, Y., and M. P. Deutscher. 2002. Mechanism of action of RNase T. I. Identification of residues required for catalysis, substrate binding, and dimerization. J. Biol. Chem. 277:50155–50159. [CrossRef]
355. Zuo, Y., and M. P. Deutscher. 2002. Mechanism of action of RNase T. II. A structural and functional model of the enzyme. J. Biol. Chem. 277:50160–50164. [CrossRef]
356. Zuo, Y., and M. P. Deutscher. 2002. The physiological role of RNase T can be explained by its unusual substrate specificity. J. Biol. Chem. 277:29654–29661. [CrossRef]
Module4.6.3
