Messenger RNA Decay
Module
4.6.4
SIDNEY R. KUSHNER
[SECTION EDITOR: GLEN BJÖRK]
June 1, 2007
Department of Genetics, University of Georgia, Athens, GA 30602-7223
Mailing address: Department of Genetics, University of Georgia, Athens, GA 30602-7223. Phone: (706) 542-8000, Fax: (706) 542-3910, E-mail:
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For many years the analysis of gene expression in Escherichia coli focused on either the process of making mRNAs (transcription) or converting the encoded information into proteins (translation). In contrast, only limited attention was given to understanding the processes by which mRNAs were functionally inactivated and nucleolytically degraded. However, over the past 15 years it has become apparent that the stability of an mRNA can play a very important role in the overall expression of its coding sequence. This has led to a renewed interest in understanding the molecular mechanisms of mRNA decay. As such, a great deal of progress has been made in understanding the enzymes involved in mRNA decay as well as important structural features of RNA molecules that can affect their stability.
This chapter discusses several topics relating to the mechanisms of mRNA decay. These topics include the following: important physical properties of mRNA molecules that can alter their stability; methods for determining mRNA half-lives; the genetics and biochemistry of proteins and enzymes involved in mRNA decay; posttranscriptional modification of mRNAs; the cellular location of the mRNA decay apparatus; regulation of mRNA decay; the relationships among mRNA decay, tRNA maturation, and ribosomal RNA processing; and biochemical models for mRNA decay. Taken together it is clear that E. coli has multiple pathways for ensuring the effective decay of mRNAs and that mRNA decay is closely linked to the cell’s overall RNA metabolism. Note that the discussion of biochemical models for mRNA decay reflects this author’s perspective on the subject and in some cases is highly speculative. For additional and sometimes different viewpoints on the subject, the reader is encouraged to examine review articles that have been authored by other investigators in this exciting field (85, 101, 103, 108, 144, 327, 332, 364). Finally, important unanswered questions in mRNA decay will be highlighted.
A unique feature in prokaryotes is the existence of both monocistronic and polycistronic transcripts. While the lengths of mRNAs can vary from as few as 325 nucleotides for the lpp transcript to many kilobases for operons such as lac or rplN, size is not a good predictor of mRNA stability. For example, the 325-nucleotide (nt) lpp mRNA has a measured half-life on the order of 10 to 15 min (159, 264, 303), while the trxA transcript of 493 nt has a half-life on the order of 1.9 to 2.5 min (14, 264, 303). Similarly, the ompA mRNA (1,215 nt) has a half-life of 15 min (28, 264, 303, 390), while the rne transcript (3,200 nt) has a half-life of 30 to 50 s (264, 305). In addition, studies of mRNA stability have shown that individual portions of a polycistronic mRNA can have significantly different half-lives (37, 212, 292). These examples demonstrate that the length of an mRNA is not a rate-limiting step in its decay.
Unlike in eukaryotes, in which the 5' terminus of each mRNA is capped with a 5'-methyl G cap structure, in E. coli the only unusual aspect at the 5' end is the triphosphate moiety. As will be discussed later, there do not appear to be 5' to 3' exonucleases in E. coli (98, 100, 101, 103, 415), nor is there currently any direct evidence for the specific enzymatic removal of the 5'-triphosphate (a prokaryotic equivalent of decapping). Since RNase E and its homologue RNase G bind to 5' termini and are inhibited by a 5'-triphosphate (240, 241, 380), as will be discussed later in more detail, it is likely that the 5' end and, in many cases, the entire 5' untranslated region (5' UTR) of an mRNA function as a stability element.
The analysis of the ompA transcript clearly indicates that a relatively short 5' UTR can dramatically affect mRNA stability. For example, if the 133-nt 5' UTR is left intact, the ompA gene has a half-life of approximately 13 to 17 min (114, 390) versus 3.6 to 4.4 min if it is removed (114). Moreover, if the 5' UTR is transferred to the 5' end of a short-lived message such as bla, the half-life of the chimeric mRNA increases from 3 to 4 min to more than 13 min (29). Emory et al. (114) have shown that the 5' UTR of ompA exists in a highly ordered structure consisting of three stem-loops. Significantly, the 5' end of the mRNA is fully paired in a double-stranded structure. As few as four unpaired nucleotides at the 5' end of the transcript are sufficient to overcome the stabilization afforded by the 5' UTR (114). More recently it has been demonstrated that there are two separate elements within the leader region that are responsible for increased mRNA stability (13).
However, not every 5' UTR serves as a stability element. The rne mRNA (encoding RNase E) contains a 361-nt 5' UTR (82). In wild-type cells the half-life of the rne mRNA is 30 to 50 s (264, 305). In the absence of a functional RNase E protein the half-life increases to more than 20 min (154). Furthermore, the presence of the 5' UTR is essential for this significant difference in stability and, in part, controls the autoregulation of intracellular RNase E levels (164) (see "Regulation of mRNA Decay," below). Detailed analysis of the 361-nt sequence has demonstrated a series of conserved stem-loop structures that facilitate the ability of RNase E to regulate its own intracellular levels (105, 168).
The results with these two mRNAs suggest that a 5' UTR can either stabilize the downstream transcript (ompA) or destabilize it (rne). In one case (ompA), it appears that stabilization is most likely caused by an unusual secondary structure that prevents access to the nuclease(s) that normally initiate decay by binding to the 5' terminus (see the subsequent discussion of RNase E and RNase G). With the rne transcript the opposite appears to be the case. Specifically, the presence of specific secondary structures appears to serve as an RNase E binding site, resulting in the rapid decay of the transcript (105).
In addition to the preceding examples, there are other mechanisms of gene regulation, such as the pnp (polynucleotide phosphorylase [PNPase]) and rnc (RNase III) mRNAs, in which cleavage at stem-loop structures by RNase III within their respective 5' UTRs significantly alters their stability (27, 320, 321). (See Fig. 1B for an example of stem-loop structure recognized by RNase III.) In these two examples, RNase III activity does not affect translation because the cleavages are well upstream of the respective ribosome binding sites; instead, it decreases the stability of the downstream mRNA. Remember that the RNase III cleavage generates a transcript with a 5'-phosphomonoester terminus.
There are also cases in which the effect of the 5' UTR may in fact relate to an inhibition of translation initiation through the masking of the ribosome binding site within a base-paired region (43, 403). Alternatively, with both the rpoS and ompA genes the 5' UTR contains a binding site for the Hfq protein (281, 391), a riboregulator that will be discussed later in more detail. In situations where translation initiation is reduced, mRNAs are more rapidly decayed (127, 161, 194, 326, 391, 399).
Many transcripts contain stem-loop structures in their 3' UTRs. Two sources of stem-loops are Rho-independent transcription termination events (277) and the presence of repetitive extragenic palindromic (REP) sequences (35, 130, 157, 365). In Rho-independent transcription terminators, a stable stem-loop structure is normally followed by a short single-stranded extension containing 1 to 8 U residues (Fig. 1A). In some cases there is no apparent single-stranded extension. This type of secondary structure can serve as a stability element because the three major 3' → 5' exonucleases known to be involved in mRNA decay (PNPase, RNase II, and RNase R, to be discussed later) all have difficulties binding to short single-stranded regions and, in the case of RNase II and PNPase, are inhibited by secondary structures (73, 75, 83, 132, 145, 147, 226, 254, 295, 360). Bioinformatic analysis suggests that almost 50% of the 2,592 protein-encoding transcription units and 71% of noncoding transcription units contain putative or reported Rho-independent transcription terminators (206).
In REP elements, the stem-loop structures are considerably longer but, in general, fall within the 3' UTR and not at the immediate terminus. However, there are cases in which these elements increase the stability of the upstream transcript (158, 286, 287).
Although poly(A) polymerase was first identified in E. coli (19), early reports of polyadenylation of mRNAs in the bacterium (135, 347, 375) were not taken seriously until the structural gene for poly(A) polymerase I (pcnB [227]) was identified in 1992 (59). It was subsequently shown that the lpp mRNA was polyadenylated after its Rho-independent transcription terminator (60). It is now generally accepted that polyadenylation stimulates mRNA decay (83, 150, 296). Furthermore, mRNA decay rates have been shown to vary as a function of intracellular poly(A) levels (264), and RNase E and PNPase levels are regulated in part by the intracellular poly(A) content (268). Recent experiments indicate that Rho-independent transcription terminators can function as polyadenylation signals (270, 271).
Another interesting aspect of polyadenylation in E. coli is the ability of PNPase to posttranscriptionally synthesize heteropolymeric tails on mRNA decay products (266, 270). What makes this reaction of interest is that, as will be discussed later, PNPase is thought to be one of the primary 3' → 5' exonucleases involved in the breakdown of mRNA decay intermediates (269). This phenomenon and other biochemical aspects of polyadenylation will be discussed in more detail later (see "Posttranscriptional Modification of mRNAs," below).
At the present time, specific nucleotide sequences, comparable to either type I or type II DNA restriction enzyme recognition sites, for example, do not appear to play any significant role in the decay of mRNAs. Thus, cleavage site specificity for endonucleases involved in mRNA decay (see "Genetics and Biochemistry of Proteins and Enzymes Involved in mRNA Decay," below) mostly depends on a combination of single-stranded regions, some of which are A/U rich, secondary, and possibly tertiary, structures. Furthermore, as will be discussed later, secondary structures are an impediment to many of the E. coli 3' → 5' exonucleases. Thus, secondary, and most likely tertiary, structures play an integral part in mRNA stability.
Most published mRNA half-life determinations measure the loss of either total pulse-labeled RNA or, more likely, a particular RNA sequence. These approaches result in the term "chemical half-life" because the numbers presented have nothing to do with the actual translatability of a given mRNA molecule. In one case, 3H-labeled uracil or uridine is used to pulse label an exponentially growing culture. After the addition of rifampin to stop new transcription initiation (152), samples are taken at various times. The loss of radioactivity is then measured by acid precipitation (to determine bulk half-lives), RNA-DNA or RNA-RNA hybridization techniques (to determine specific mRNAs half-lives), RNA-DNA protection analysis (390), or primer extension approaches (134).
In determining bulk RNA half-lives, a problem with the technique has been what to use as the baseline for 3H incorporation into stable RNA species. With wild-type E. coli, the published numbers are probably a reasonable estimate because, by 60 min after the cessation of new transcription, most of the remaining acid-insoluble radioactivity will be associated with ribosomal and transfer RNA species. However, with many of the mutants that have much slower mRNA decay rates (14, 139, 296), choosing an appropriate baseline is more difficult.
As such, this method has largely been replaced by the use of Northern blots to determine the half-lives of either individual full-length transcripts or RNA-protected DNA fragments. In this approach, total RNA samples are resolved on either polyacrylamide/urea or agarose/glyoxal gels. Species of up to 1,000 nt can be easily resolved on polyacrylamide gels (14, 303), while larger mRNAs can be separated on agarose gels, although the resolution is not as good. In addition, with larger RNA molecules there may be problems associated with the transfer from the gel to the membrane as well as interference from either 16S or 23S rRNA.
The additional technical problems associated with most half-life experiments need to be considered. For one, although adding rifampin is thought to specifically block transcription initiation, it may affect the cell in some other way that, in turn, indirectly affects mRNA decay. Second, in many E. coli mutants mRNA decay curves frequently appear biphasic. In one case an initial rapid burst of decay is followed by a much slower turnover of the remainder of the transcript. Alternatively, mRNAs may initially appear very stable and then decay rapidly. Until these types of decay curves are better understood at the biochemical level, it may not be possible to accurately determine a true chemical half-life for certain mRNAs.
Another problem associated with chemical decay studies is that measurements for the same mRNA may vary considerably between laboratories and even within laboratories. These differences most likely arise from the various different methods used for isolating RNA. Thus, it is probably best to view chemical half-lives as diagnostic rather than as absolute numbers. Comparisons, however, do appear valid if half-lives have been determined within the same experiment. For example, the difference between a half-life of 4 min and 5.5 min may be significant if the two numbers were determined side by side in several experiments. If, however, these two numbers arose from two determinations done in separate laboratories, the difference may not be statistically significant.
Another caution is worth mentioning. Even if the half-life of a full-length transcript increases significantly in a particular E. coli mutant, it does not necessarily mean that the functional half-life of the transcript has increased. For example, Ow et al. (305) showed that while the chemical half-life of the rne transcript increased in several mutants, there was actually a reduction in the level of RNase E protein. Thus, an increased chemical half-life, even for a full-length transcript, does not necessarily correlate with an increased protein level.
Finally, note that efforts have recently been undertaken to obtain a global view of mRNA decay by employing microarrays (30, 31, 355). In this approach, open reading frames, or portions of them, are spotted on microarray chips. Total RNA is isolated from cells after rifampin treatment and converted to labeled cDNAs using either gene-specific or random primers. However, there are several inherent limitations with this method. If only a portion of each open reading frame is on the chip, the half-life measurement will not be for the full-length mRNA. In addition, because of the method used for converting the RNA into labeled cDNAs, mRNA decay intermediates may hybridize along with full-length species. Thus the data obtained from arrays are best viewed as qualitative rather than quantitative. For more accurate measurements, Northern blot analysis of an individual mRNA is absolutely essential.
A functional half-life measures how long an mRNA remains sufficiently intact such that it can be translated to produce a protein that is biologically active. These measurements are far more difficult than examining chemical decay, in general, requiring either an immunological (287) or enzymatic assay (37) for the protein product of a given mRNA. The technique is particularly suited for inducible transcripts such as gal (1) or lac.
Alternatively, Pedersen et al. (311) used a completely different approach to measure functional half-lives. They assumed that the potential to synthesize a particular protein was equal to the functional decay rate of the corresponding mRNA. Synthetic potential was determined using two-dimensional polyacrylamide gel electrophoresis of 35S-labeled proteins following inhibition of transcription with either rifampin or streptolygidin. In this fashion they determined half-lives ranging from 40 s for ribosomal protein S1 to 20 min for an unidentified protein (311). Their data also suggested that mRNAs within a single transcriptional unit could have different functional half-lives (311).
Remember that functional half-lives measure a completely different type of mRNA decay. All that is necessary to functionally inactivate an mRNA is a single phosphodiester bond cleavage (to remove the ribosome binding site, for example). A similar cleavage within the coding sequence could also lead to functional inactivation, but only if the truncated polypeptide produced is no longer biochemically active. It is possible that mRNAs with short functional half-lives can have long chemical half-lives. It is thus important when reading the mRNA decay literature to distinguish between chemical and functional half-life determinations.
As initially envisioned by Apirion (9) in 1973, mRNA decay in E. coli involved the combined action of endo- and exonucleases to convert full-length transcripts into mononucleotides. However, at the time his paper was published there were few genetic or biochemical data to support his hypothesis. In the ensuing years considerable progress was made in understanding the complex events involved in mRNA decay. Deutscher (98, 100, 101) (see also Chapter 4.6.3, Exoribonucleases and Endoribonucleases) has summarized the biochemical properties of many of the E. coli ribonucleases on several occasions. I have chosen here to try to integrate the genetic and biochemical evidence that indicates a role for a particular protein in mRNA decay. In addition, some interesting genes are also discussed even though their relevant biochemistry is not currently complete.
As historically defined, exonucleases degrade RNA substrates one nucleotide at a time from either the 3' or 5' terminus. In E. coli there are examples of both hydrolytic decay, which releases mononucleotides (RNase II, RNase R, oligoribonuclease), and phosphorolytic cleavage, which generates nucleoside diphosphates (PNPase and RNase PH). In addition, there are two ribonucleases (RNase E and RNase G) whose catalytic activities are stimulated by binding to a 5' terminus but that cleave phosphodiester bonds at a distance (104, 169, 240, 241, 380). While these enzymes are considered endonucleases, note that deoxyribonucleases with similar specificities (e.g., RecBCD nuclease [308, 357]) are defined as exonucleases.
RNase II (rnb).
RNase II is a hydrolytic 3'-5' exonuclease that is highly processive but is sensitive to secondary structure (295, 358, 360). The enzyme requires both magnesium and potassium for full activity (147). In cell extracts, it accounts for 90% of the nucleolytic activity (99). The first rnb mutation was identified in 1976 (289). Some seven years later the structural gene for RNase II (rnb) was cloned and shown to encode a 72,360-dalton protein (106) (Table 1). Overexpression of RNase II activity did not change mRNA decay rates (106). More recent studies have shown that RNase II levels are regulated in part by a protein called Gmr (46). Furthermore, the intracellular level of RNase II increases in the absence of PNPase (414) and can vary as a function of growth conditions (46). In the past year the three-dimensional structure of the protein has been determined (123, 417).
Table 1Enzymes, proteins, and RNAs involved in mRNA decay |
While Apirion and coworkers suggested that RNase II was involved in mRNA decay (178, 193), it was not until 1986 that Donovan and Kushner (107) demonstrated directly that RNase II was both involved in mRNA decay and essential for cell viability in the absence of polynucleotide phosphorylase. RNase II has also been shown to participate in a limited way in tRNA processing (335). It has also been argued that RNase II plays a significant role in the degradation of poly(A) tails (244, 267). Other studies have suggested that in some cases RNase II may actually protect mRNAs from decay (149, 269, 313) by either blocking access to the 3' terminus through the rapid degradation of a poly(A) tail (84) or failure to dissociate from a substrate containing a stable stem-loop structure (Fig. 2). This argument is based on the fact that the enzyme is strongly inhibited by secondary structures (84, 360), such as those associated with Rho-independent transcription terminators.
Polynucleotide phosphorylase (pnp).
In the presence of millimolar levels of inorganic phosphate, polynucleotide phosphorylase catalyzes the processive 3' → 5' phosphorolytic degradation of RNA to nucleotide 5'-diphosphates (226). It was first identified in Azotobacter vinelandii (142, 143) and shortly thereafter in E. coli (225). However, it took a long time after the first pnp mutations were isolated by Reiner (334) in 1969 for its a role in mRNA decay to be clearly demonstrated by Donovan and Kushner (107). Early purification studies suggested that the enzyme consisted of two subunits of 86,000 and 48,000 daltons (318, 319). When the structural gene for the catalytic subunit (pnp) was cloned and sequenced (320, 330), it became clear that the molecular weight was actually 79,715 daltons (Table 1). Although it was thought that the second subunit was the glycolytic protein enolase (45,534 daltons, Table 1) (259, 324), recent work suggests that in fact this protein is the RhlB RNA helicase (46,995 daltons, Table 1) encoded by the rhlB gene (221). X-ray crystallographic analysis of the catalytic subunit from Streptococcus antibioticus has demonstrated a duplicated fold as the structural basis for catalytic activity (370). Because of the high degree of sequence homology between the E. coli and S. antibioticus proteins, Duran-Figueroa et al. (111) have modeled the E. coli PNPase and mapped its interaction domains with RNase E.
Deutscher (100) has pointed out that phosphorolysis has the advantage of capturing the energy from phosphodiester bond cleavage in the form of a nucleotide diphosphate. Data now indicate that polynucleotide phosphorylase is the primary exonuclease involved in the degradation of poly(A) tails (267, 296) and in mRNA decay (269). Furthermore, as will be discussed later in more detail, the enzyme also functions under certain circumstances as a poly(A) polymerase (266). Note that PNPase is inhibited by secondary structures (360) but has been shown to be capable of degrading certain stem-loop structures in combination with poly(A) polymerase (84). In addition, its interactions with RNase E (63, 323) and possibly with the RhlB RNA helicase may facilitate its ability to degrade mRNAs or decay intermediates including those containing secondary structures (221, 224).
RNase R (rnr/vacB).
The vacB locus was first identified based on its role in virulence in Shigella flexneri and E. coli (74, 379). Subsequently, it was shown to encode a 3' → 5' exonuclease now called RNase R (74). RNase R single mutants have little if any demonstrable phenotype. In contrast, RNase R PNPase double mutants are inviable (74), in a fashion similar to that observed for RNase II PNPase double mutants (107). However, the loss of cell viability in PNPase RNase R double mutants has been attributed to a defect in the degradation of rRNAs, not mRNAs (76). While the enzyme shows considerable sequence identity and similarity with RNase II, RNase II RNase R double mutants only show a small alteration in growth compared with either single mutant (B. Mohanty and S. R. Kushner, unpublished results). However, even in the absence of both RNase II and PNPase, the degradation of mRNA decay intermediates is slowed but not completely inhibited (14, 16, 17, 296).
More detailed analysis of RNase R has demonstrated that the enzyme can processively degrade a variety of RNAs containing extensive secondary structures, including rRNA (75). However, the enzyme needs between a 7- and 10-nt single-strand extension to bind to a substrate (388). For example, the addition of a poly(A) tail to an RNA transcript with a very short single-stranded 3' terminus makes this molecule a good substrate of RNase R even if it contains a stem-loop structure (73). Thus RNase R, under certain circumstances, clearly plays a role in mRNA decay, in particular, in the degradation of mRNAs containing REP elements (73) and nonstop RNAs (336).
Analysis of the regulation of RNase R synthesis has shown that its levels increase up to 10-fold under a variety of stress conditions, including entry into stationary phase (8, 72). In addition, the rnr gene is cotranscribed as part of a larger polycistronic operon that also encodes yjeB, rlmB, and yjfI (47). This large mRNA is rapidly degraded into smaller fragments by RNase E and possibly RNase G, suggesting that in vivo levels of the protein are partially a function of the stability of the rnr transcript (47).
Oligoribonuclease (orn).
Oligoribonuclease is a 3' → 5' exonuclease (Table 1) that, unlike any of the other enzymes described in this section, is specific for short oligoribonucleotides (294, 408). In fact, it has been shown that it is essential for cell viability and that oligonucleotides between 2 and 5 nt in length accumulate under conditions where the intracellular level of the enzyme is significantly reduced (129). Thus, oligoribonuclease appears to be responsible for the degradation of the terminal degradation products that are no longer substrates for PNPase, RNase II, and RNase R.
Other exonucleases.
Deutscher and his colleagues (53, 182, 183, 213, 335, 416) have carefully studied a variety of 3' → 5' exonucleases (RNase PH, RNase D, RNase BN, RNase T) that are involved in tRNA maturation. Because of their very limited substrate specificities, RNase D and RNase T do not seem likely to participate in mRNA decay (54, 406, 416). In the case of RNase PH, direct analysis of rph mutants has indicated that this enzyme has no role in mRNA decay (B. Mohanty and S. R. Kushner, unpublished results). RNase BN (18, 53, 54, 115; Table 1) will be discussed later, since it appears to be identical to the endoribonuclease RNase Z (53, 54, 115, 316).
Riboendonucleases are defined as enzymes that cleave at internal locations. As will be noted in the discussion that follows, the substrate specificities associated with the known E. coli enzymes indicate a significant role for higher-order structures rather than actual nucleotide sequence.
RNase I/RNase I* (rna).
RNase I was originally identified as a nonspecific endonuclease located in the periplasmic space of E. coli (285). The structural gene for RNase I (rna) has been cloned and sequenced (255, 412) and shown to encode a 29,487-dalton polypeptide (Table 1). Meador et al. (255) demonstrated that RNase I could degrade mRNA as well as the four ribonucleotide homopolymers.
While RNase I is a potential candidate for an mRNA decay enzyme, its location in the periplasmic space (285) and the absence of any demonstrable phenotype in rna deletion mutants (412) seem to rule it out as a major player in mRNA decay. Hautala et al. (153) did note, however, that the rna-19 allele had a small effect on the expression of the Neurospora crassa catabolic dehydroquinase gene in E. coli. This observation suggests that RNase I might play some role in mRNA decay. In fact, Cannistraro and Kennell (57) have reported an altered form of RNase I, called RNase I*, which is associated with the inner membrane. Based on what is now known about the intracellular location of the RNase E-based degradosome (see "Cellular Localization of the mRNA Decay Apparatus," below), this observation takes on added significance. They have suggested that RNase I* has a small role in the cleavage of larger RNA oligonucleotides but that its primary function is the degradation of small oligonucleotides (58).
RNase III (rnc).
RNase III was first identified as an endoribonuclease that cleaves double-stranded RNA molecules (339, 340). Since then, it has been extensively studied to determine its substrate specificity, as reviewed by Court (92) and Nicholson (288). The structural gene for RNase III (rnc) encodes a 25,419-dalton polypeptide (Table 1) that functions as a homodimer. In vivo, the enzyme specifically recognizes certain stem-loop structures (Fig. 1B) and can cleave either on one or both sides of the stem, usually within an internal unpaired region, to yield a two-base 3' overhang (38, 234, 307, 354, 402). By comparing 34 RNase III cleavage sites, Krinke and Wulff (198) found a consensus sequence of A/UNAGA/UGNNCA/UUNN within one arm of the stem. However, the conserved CUU/GAA base-paired sequence immediately adjacent to the unpaired region where cleavage occurs was not required for accuracy or selectivity (71). It is now thought that the specificity of RNase III resides in the exclusion of certain base pairs from the stem structure and specific local tertiary structures (208, 407). Furthermore, a minimal substrate for RNase III has been determined to contain two discrete double-helical elements (314).
Following the 1973 identification of the first rnc mutation (192), RNase III was subsequently shown to participate in the processing of 30S ribosomal RNA (110). Although the major phenotype associated with the rnc-105 allele is the limited accumulation of unprocessed 30S rRNA transcripts, approximately 10% of all cellular proteins are either under- or overproduced in a rnc-105 mutant (131, 372). This observation suggests that RNase III plays a direct role in the half-lives of a discrete subset of E. coli mRNAs, even though the half-lives of the majority of cellular mRNAs are essentially unaffected (12, 372). This hypothesis has been supported by observations that the enzyme directly affects the stability of several mRNAs, including rnc (27), pnp (321), dicB (117), and metY (331). Takiff et al. (374) and Babitzke et al. (21) have demonstrated that RNase III deletion mutants remain viable.
RNase E (rne/ams).
Apirion and his coworkers originally identified RNase E (rne) as an enzyme involved in the processing of the 5S rRNA from a 9S rRNA precursor (11, 128). The genetic proof that RNase E was involved in mRNA decay took a somewhat more convoluted path. In 1977, Kuwano et al. (200) isolated a mutation called ams (altered messenger RNA stability) that led to inviability at elevated temperatures and two- to threefold increases in the chemical half-life of total E. coli pulse-labeled RNA. It was also noted that while mRNA decay appeared to be altered, protein synthesis continued normally for several hours after shift to the nonpermissive temperature (299).
Subsequently, Arraiano et al. (14) showed that a triple mutant containing mutations in the ams pnp and rnb genes led to a significant increase in the half-life of total pulse-labeled RNA at 44oC (11 to 15 minutes in the triple mutant versus 3 minutes in a wild type control). Additionally, they demonstrated that discrete decay intermediates of the trxA (thioredoxin) and cat (chloramphenicol acetyltransferase) mRNAs were observed in such mutants following shift to the nonpermissive temperature (14). Chemical half-life measurements for specific full-length mRNAs showed increases from two- to fourfold (14).
Further studies with the ams-1 pnp-7 rnb-500 triple mutant indicated that the decay intermediates observed with trxA (16) and pyrF (17) arose from multiple endonucleolytic cleavages, many of which left the 5' termini of both mRNAs intact. Microarray analysis now indicates that RNase E is the primary endoribonuclease that initiates mRNA decay in E. coli (31).
The cloning and initial sequencing of the ams gene (81, 82) assisted researchers in demonstrating that ams and rne encoded the same protein (20, 257, 279, 376). Further studies by Casagerola et al. (65, 66) and Cormack et al. (91), showed that the original ams/rne clones were missing DNA sequences encoding more than 200 amino acids from the carboxy terminus of the product. The truncated clones were sufficient, however, to complement the ams-1 and rne-3071 mutations in vivo (81). These two alleles map at amino acids 66 and 68, within the S1 RNA binding domain of the protein (Fig. 3) (250).
rne encodes a 1,061-amino-acid protein whose calculated molecular mass is 118,066 daltons (Table 1) but which migrates on sodium dodecyl sulfate (SDS) gels with an apparent size of 180,000 daltons. The protein is unusual because it contains more than 25% charged amino acids, has an abundance of proline residues in the central region, and is much larger than all the other characterized E. coli ribonucleases. The enzyme requires either Mg2+ or Mn2+ and a monovalent cation for activity (263). Of considerable interest is the observation that RNase E appears to be a 5' end-dependent endonuclease (240, 241, 380), although under certain circumstances the enzyme can cleave at internal sites without binding to the 5' terminus (24). Furthermore, the amino-terminal portion of the protein forms multimers that in vitro are more active than monomers (169). In fact, structural studies suggest that the catalytic domain of RNase E is a homotetramer that is stabilized in part by Zn ions (49, 52). The determination of the three-dimensional structure of the amino-terminal 529 amino acids has led to the identification of five subdomains within the catalytic region of the protein: an S1 RNA binding domain (45), a 5' sensor region, an RNase H domain, a DNase 1 domain, and a Zn2+ link (51) (Fig. 3).
The substrate specificity of the enzyme is still not completely understood. Initially, Mudd et al. (278) proposed a 10-nt recognition sequence based on the observations of Ghora and Apirion (128) and their own experience with processed bacteriophage T4 mRNAs (278). Subsequently, Ehretsmann et al. (112) suggested that the consensus recognition sequence was A/GAUU/AU, a subset of the 10-nt site. Additional work from the laboratory of S. Cohen, however, suggested that RNase E has few primary structural constraints other than a preference for cleaving 5' to an AU dinucleotide (219, 251, 252). However, a more detailed analysis by Kaberdin (172) has indicated a preferred cleavage that contains considerable redundancy, G/AC/A N G G/U/A ↓A/UC/U N C/AC/A. However, he noted that alterations in the efficiency of cleavage at certain sites could be mediated by interactions with adjacent regions of secondary structure (172). For example, RNase E cleaves within its own leader region at the sequence (ACCC↓↓A↓UUUUG) (Q. Liu, M. Ow, and S. R. Kushner, manuscript in preparation), which is significantly different than the site proposed by Kaberdin (172). As such, caution should be observed when trying to identify RNase E cleavage sites strictly on the basis of sequence scanning.
Another question regarding RNase E is the nature of the active protein in vivo. The enzyme is very susceptible to proteolysis, and several truncated forms have been detected that are still enzymatically active. While the full-length protein has catalytic activity (91, 377), a variety of C-terminal truncation mutants have been characterized that not only support cell viability but are relatively normal regarding mRNA decay at a variety of physiological temperatures (205, 233, 303, 304, 387). In fact, experiments by Caruthers et al. (64) suggest that a protein of only 395 amino acids, lacking the Zn link described by Callaghan et al. (51, 52), can support cell viability.
Of great interest is that RNase E copurifies with at least three other proteins, the 3' → 5' exonuclease PNPase, an RNA helicase called RhlB, and the glycolytic enzyme enolase, in a complex now called the degradosome (63, 259, 323, 324). There have also been reports of weak interactions between RNase E and polyphosphate kinase (36) and possibly poly(A) polymerase (329). The function of the degradosome will be discussed in more detail in "Biochemical Mechanisms for mRNA Decay," below. Detailed analysis of the scaffolding region of RNase E suggests that it is rather flexible and has little structure (50).
Finally, note that, besides its very important role in mRNA decay, RNase E is involved with most if not all aspects of RNA metabolism in E. coli. It has already been noted that the enzyme is required for the processing of both 9S RNA (128) and 16S rRNA (215, 393). In addition, the enzyme also plays a critical role in the maturation of tRNAs in the bacterium (216, 218, 304, 328). In fact, it has been argued that the essential function of RNase E is its requirement for the initiation of tRNA maturation (304). The enzyme is also required for the maturation of the RNA subunit of RNase P (236), the maturation of the ssrA RNA (tmRNA; see Chapter 4.2.7 for a detailed discussion of this molecule and its role in trans-translation) (220), and the decay of many small noncoding RNAs (247, 405). Since there is some indication that there is a hierarchy among RNase E substrates (303), at this point it is worth remembering that the phenotype associated with any rne mutant is probably a composite of defects in multiple RNA decay/processing/maturation pathways. For example, the rne-1 alleles to a significant defect in 9S rRNA processing at the nonpermissive temperature (128, 303), but the rneΔ610 does not, even though strains carrying either allele are defective in mRNA decay (303).
RNase K (rne).
RNase K was initially identified based on a series of site-specific cleavages in the 5' UTR of the ompA mRNA that seemed to regulate the stability of the transcript (235, 291). The enzyme was partially purified and shown to have a molecular weight between 60,000 and 62,000 daltons. At the time that this work was published there were conflicting reports regarding the size of RNase E, ranging from 66 kDa (344) to what turned out to be its true molecular weight of 118 kDa (65, 66). With the observation that RNase E was quite susceptible to proteolysis (63, 91) and the demonstration that RNase E could also cleave in the 5' UTR of the ompA mRNA (280), it became clear that RNase K was a proteolysis product of RNase E, not a unique enzyme (237).
RNase G (rng/cafA).
RNase G was initially identified as a protein involved in the formation of cytoplasmic axial filaments (298). It was subsequently noted that there was extensive sequence identity (34.1% over the first 489 amino acids, Fig. 3) between the CafA protein and the N terminus of RNase E (392). It has now been shown that the cafA gene encodes a 5' end-dependent endoribonuclease whose catalytic activity is similar to that of RNase E (215, 245, 380) (Table 1). Both RNase G and RNase E participate in the maturation of the 5' terminus of 16S rRNA, but they cleave at distinct sites and RNase G is responsible for generating the mature 5' end of the 16S rRNA (215, 380, 393).
Although inactivation of RNase G does not lead to any significant phenotypic alterations in the cell (298, 392), it has been observed that the stability of the adhE and eno mRNAs increases in rng mutants (174, 394). In addition, microarray analysis of the E. coli transcriptosome showed that inactivation of RNase G led to a significant increase (>twofold) in the steady-state level of 11 mRNAs (202).
Detailed genetic analysis has shown that RNase G apparently serves as a backup for RNase E in both the processing of 9S rRNA precursors and the decay of a variety of mRNAs but probably does not participate significantly in tRNA maturation (306). However, there are now conflicting data as to whether RNase G can stably complement RNase E mutations under conditions where the enzyme is dramatically overproduced (95, 202, 306, 392) (D. Chung and S. R. Kushner, manuscript in preparation). What is clear is that native RNase G cannot complement RNase E mutants under normal physiological conditions (95, 306) and that there are very small amounts of the protein in the cell (42).
RNase M.
In 1989, Cannistraro and Kennell (56) characterized a 26,000-dalton protein they called RNase M (Table 1), which they argued was involved in the decay of the lac mRNA. The enzyme appeared to have a target specificity of Pyr-A and cleaved RNA to give 5'-OH termini (56). Based on tryptic fingerprints, the protein seemed to resemble RNase I (255), but its activity was still present in an rna mutant strain (MRE600) (362). While Kennell has argued that RNase M is the primary endoribonuclease for mRNA degradation in E. coli (58, 185), biochemical data now suggest that the RNase M is a multiply mutated form of RNase I and is probably found only in the strain originally used for its purification (367).
RNase N.
In 1976, Misra et al. (261) reported the identification of a new endoribonuclease that could cleave rRNA as well as homopolymers into small oligonucleotides and 5' mononucleotides. Subsequently, this protein was examined in more detail and was shown to be a homodimer with a native molecular mass of 120 kDa (262). Additionally, they noted that, unlike RNase III, RNase N could digest single-stranded and double-stranded RNAs with equal efficiency (262). Unfortunately, no further work has been done on this enzyme, although it was reported as an unpublished observation that Subbarayan and Deutscher (102) were not able reproduce the original observations. Despite this report, it is worth noting that RNase N was purified from an RNase III-deficient strain (261) and that its catalytic properties and molecular weight make it distinct from RNase E, RNase G, RNase Z, and RNase LS. It would seem, therefore, that there is a distinct possibility that E. coli may have at least one additional endoribonuclease.
RNase P.
RNase P is an unusual ribonuclease in that it contains both a protein and catalytic RNA subunit (363) (Table 1). The enzyme is essential for cell viability, presumably because it produces the mature 5' terminus on all tRNA species in the bacterium (7). The first indication that it might be involved in mRNA decay was the observation by Alifano et al. (6) that the polycistronic his mRNA in Salmonella enterica serovar Typhimurium was cleaved by RNase P. Subsequent microarray analysis has shown a limited number of RNase P cleavage sites within the intercistronic regions of the tna, secG, rbs, and his operons in E. coli (210). It has also been demonstrated that an RNase P cleavage site exists between the lacY and lacA genes, which may help explain some of the polarity effects observed with this operon (212). The authors showed that cleavage of mRNA transcripts occurred at a rate ten times slower than that observed with tRNA precursors (212). The implications of this observation will be discussed below, in "Biochemical Mechanisms for mRNA Decay."
RNase LS (rnlA/yfjN).
It was reported in 2003 that a chromosomal mutation in E. coli, initially called std-2, led to the stabilization of bacteriophage T4 mRNAs (301). Subsequently, the std-2 mutation was shown to map to the yfjN locus, which has now been renamed rnlA; the protein it encodes is RNase LS (302) (Table 1). While inactivation of RNase LS appeared to partially stabilize the rpsO mRNA (302), its primary role seems to be as an antagonist of bacteriophage T4 infection (398). Very few homologues of RNase LS are found in other prokaryotes.
RNase Z (rnz/elaC).
The elaC gene of E. coli encodes a binuclear zinc phosphodiester protein (351) (Table 1) that belongs to a large superfamily, whose representatives are found in prokaryotes, eukaryotes, and archaea (109, 260, 312, 349, 350). In eukaryotes and Bacillus subtilis the enzyme called RNase Z has been shown to be involved in the maturation of the 3' termini of tRNA precursors that do not include a chromosomally encoded CCA determinant (312, 350). Since all of the tRNA precursors in E. coli contain encoded CCA determinants (35), it was not clear what function, if any, this protein had in the bacterium. However, recent experiments have demonstrated that RNase Z is active in E. coli but is not involved in tRNA maturation (316). Rather it serves as backup endonuclease in mRNA decay (316). Furthermore, it has also been demonstrated that, at high protein levels, in vitro RNase Z has limited 3' → 5' exonuclease activity, which was previously called RNase BN (18, 54, 115, 352). Based on its recently determined crystal structure, RNase Z in E. coli appears to function as a homodimer (197).
Poly(A) polymerase (pcnB).
Poly(A) polymerase catalyzes the template-independent addition of AMP moieties onto the 3'-terminal hydroxyl groups of specific RNA molecules using ATP as a substrate. The enzyme was first identified in E. coli in 1962 (19) and purified and characterized in 1976 (325) (Table 1). It requires both Mg2+ and Mn2+ for optimal activity (59). In vitro the enzyme will also incorporate non-A residues (400) but not in vivo (266). The structural gene for poly(A) polymerase I (pcnB) has been cloned and sequenced (59, 227, 243). Work by Mohanty and Kushner (264) has demonstrated that the level of poly(A) polymerase is kept low in the cell in part by the inefficient translation of its mRNA. It is estimated that there are only between 30 and 50 molecules per cell (270). In fact, overproduction of the enzyme is very toxic to the bacterium (264). A role for poly(A) polymerase in mRNA decay has now been clearly demonstrated (83, 150, 264, 296). This evidence will be discussed in more detail below (see "Posttranscriptional Modification of mRNAs").
It has been reported that E. coli contains a second poly(A) polymerase (175) that accounts for the small amount of polyadenylation seen in pcnB mutants (296), but the protein ostensibly encoding this putative poly(A) polymerase (61) is unable to polyadenylate mRNAs (265). Rather it has now been shown that PNPase is the second poly(A) polymerase type activity in E. coli (266) and that it functions to add polynucleotide tails in both pcnB and wild-type strains (270). A mutant lacking both PNPase and PAP I has no detectable polyadenylation in exponentially growing cultures (266). New work has also demonstrated that Rho-independent transcript terminators serve as polyadenylation signals (270, 271) and that most transcripts in exponentially growing cells undergo some degree of polyadenylation (271).
Hfq (hfq).
Hfq is a small (11,035 daltons) RNA binding protein (Table 1) that was first identified based on its requirement for the replication of the bacteriophage Qβ (122). It belongs to the family of Sm and Sm-like proteins (348, 404). A variety of studies have shown that Hfq acts as a pleiotropic regulator of many genes, in part through its effects on the stability of specific mRNAs and small regulatory RNAs (282, 385, 386). Hfq appears to protect regulatory RNAs by binding to A/U-rich regions that are also RNase E cleavage sites (272, 405). Furthermore, it has been suggested that Hfq can interact directly with RNase E to form a complex that facilitates the degradation of small noncoding RNAs (275).
The first indication that Hfq might play some role in mRNA decay came from experiments showing that the absence of the protein affected the polyadenylation of the rpsO transcript (151, 201). Furthermore, it has also been demonstrated that Hfq is required for the polyadenylation of mRNAs containing Rho-independent transcription terminators through the formation of a multiprotein complex that seems to contain Hfq, PAP I, and PNPase (270) (Fig. 2). The exact role of Hfq in polyadenylation is still not fully understood because some evidence has been presented that the protein can bind equally well at terminal or internal stretches of poly(A) (119). However, these results are not supported by data that Hfq does not protect poly(A) tails from degradation from either PNPase or RNase II (270).
Since most RNAs contain secondary structures and these structures have been shown to inhibit the activity of both RNase II and PNPase (145, 360), it is not surprising that RNA helicase might be required for mRNA decay. RNA helicases in E. coli belong to the DExD/H family, which is related in sequence and structure to helicase superfamily 1 (137). E. coli contains five DEAD-box genes (csdA [formerly called deaD], dhpA, rhlB, rhlE, and srmB) and 13 DEx-H-box genes (162, 315). Of the DExH box genes, only the HrpA helicase apparently participates in RNA metabolism (195), as will be discussed below. The reader is directed to recent reviews for a more detailed discussion of RNA helicases (90, 162).
RhlB RNA helicase.
The RhlB RNA helicase was first identified as a member of the DEAD box family of helicases (176) (Table 1). The protein was subsequently shown to be associated with RNase E as part of the degradosome (259, 324). RhlB by itself is not a very effective RNA helicase but is significantly activated by its interaction with RNase E (86). In fact, in vitro experiments have shown that there may be two distinct binding sites on RNase E for RNA helicases (187). In this work RhlB could be replaced by the association of RhlE at a different site on the RNase E protein (187). The nature of the physical interaction of the RhlB protein with RNase E has also been determined (68). In addition, it has been reported that the RhlB protein is actually the second subunit of PNPase (224), suggesting a possible alternative complex that can promote the degradation of structured RNA intermediates by PNPase in the absence of RNase E. Genetic and microarray analysis suggests that the enzyme is required for the decay of certain specific mRNAs, RNA decay intermediates that contain REP elements, and untranslated mRNAs (31, 186, 188).
HrpA.
The hrpA gene was first identified based on its similarity to the DEAH family of RNA helicases from Saccharomyces cerevisiae (276) (Table 1). Subsequently, the putative HrpA RNA helicase was shown to be involved in the cleavage of a polycistronic mRNA that encodes a fimbrial adhesion designated F1845 (32). Specifically, when it was shown that this polycistronic mRNA was processed independent of both RNase III and RNase E (33), experiments were undertaken to identify other proteins that facilitated this reaction. Initially, it was shown that there was translational control of the mRNA-processing event (230) that involved a tripeptide sequence within the nascent DaaP protein (231). Subsequently, Koo et al. (195) isolated a hrpA mutant that was defective in the processing of the daaA-E polycistronic transcript. Exactly how this putative RNA helicase acts to promote the cleavage of the mRNA is not fully understood at this time. Another DEAD RNA helicase, SrmB, was shown not to substitute for HrpA (195).
SrmB.
The srmB locus was first isolated as a multicopy suppressor of a temperature-sensitive mutation in the ribosomal protein L24 (293) (Table 1). Mutations in this locus lead to defects in the biogenesis of ribosomes at 20oC (284) and more specifically of 50S ribosomal subunits (69). Iost and Dreyfus (160) showed that overproduction of the SrmA protein led to the in vivo stabilization of a lacZ mRNA generated from a bacteriophage T7 promoter. Additional evidence for a possible role for this RNA helicase in mRNA decay comes from the findings that the SrmB protein can bind to RNase E (187) and that based on Far Western analysis associates with PAP I (329).
RhlE.
The existence of the rhlE-encoded RNA helicase (Table 1) was first predicted as an RNA helicase in work that identified another DEAD-box enzyme, RhlB (176), which was discussed above. Subsequently, a deletion of the rhlE-coding sequence was constructed but the mutant had no discernable phenotype (297). However, some role for the rhlE-encoded RNA helicase been suggested by experiments in which the protein was able to substitute for RhlB in an in vitro degradosome assay and was shown to bind to RNase E at a site different from RhlB (187). In addition, the protein is also thought to interact with PAP I (329).
DeaD/CsdA.
The DeaD RNA helicase (Table 1) was also identified as a multicopy suppressor of a mutation in the ribosomal protein gene rpsB (383). Subsequently, it was shown that this gene was strongly induced during cold shock and was renamed csdA (170). Clear evidence has been presented that this RNA helicase is involved in the biogenesis of the 50S ribosomal subunit (70) as well as in the stabilization of mRNAs when they are overexpressed (39, 322). Further indication for a role of this protein in mRNA decay comes from the identification of a "cold shock degradosome" including RNase E and CsdA (322). Finally, it has been shown that CsdA can interact with PAP I (329).
Enolase.
The discovery of the glycolytic enzyme enolase (Table 1) as a component of the degradosome (63, 259, 323, 324) was somewhat surprising. The three-dimensional structure of enolase associated with its recognition site from RNase E has been determined (67). Although it was initially thought not to have any significant affect on mRNA decay, recent experiments suggest a definitive role for the protein in the decay of specific mRNAs (31, 274).
DnaK.
The DnaK protein (Table 1) is a well-described heat shock-inducible chaperone (26) that was originally identified as a minor component of the degradosome (259). By using a proteomic approach, it has been shown that while DnaK is not required for either degradosome assembly or structure it does associate with RNase E under certain stress conditions such as cold shock or in the absence of PNPase (333).
RraA and RraB.
Originally identified in a screen for proteins that could increase the level of disulfide bond isomerization in E. coli, RraA is a 17.4-kDa protein (Table 1) that inhibits RNase E activity through a direct protein-protein interaction with the enzyme (203). Crystallographic data of RraA have suggested that the protein contains a unique fold that represents an early structure, which has been adapted for a wide range of functions (273). Microarray analysis indicates that the stability of approximately 80 mRNAs is affected by inactivation of the rraA (regulator of ribonuclease activity A) gene (203). Further support for the interaction of RraA and RNase E comes from data that simultaneous overexpression of both proteins overcomes the toxicity associated with increased levels of the endonuclease (401). Furthermore, it has now been shown that a second inhibitor protein called RraB binds to RNase E at a different location and interferes with the decay of a different set of transcripts (124).
CsrA.
The csrA gene encodes a 61-amino-acid protein (Table 1) that is involved in the regulation of a variety of stationary-phase genes (228, 229, 341, 342, 345). The protein has been shown to interact with the regulatory regions of the mRNAs it controls, in some cases by preventing translation (22). Inhibition of translation leads to more rapid turnover of the transcripts. Antagonists for this protein are two nontranslated RNAs encoded by the csrB and csrC genes (228, 396) (Table 1). In addition, the CsrA protein has been shown to activate the transcription of the csrB RNA (146).
CsrD.
With the identification of proteins that specifically inhibit the activity of RNase E (124, 203), it was not entirely unexpected that there might be factors that actually stimulated its activity on particular substrates. This has, in fact, turned out to be the case with the CsrD protein (369) (Table 1). Specifically the CsrD protein somehow interacts with both the CsrB and CsrC regulatory RNAs or RNase E itself to convert the regulatory RNAs into good substrates for RNase E (369).
When Babitzke et al. (21) constructed an rne pnp rnb rnc quadruple mutant and examined mRNA decay compared with an rne pnp rnb triple mutant, they found that eliminating RNase III led to more rapid mRNA turnover. This surprising result was interpreted to mean that the absence of RNase III leads to an increased synthesis of enzyme(s) that can compensate for the loss of RNase E, RNase III, RNase II, and PNPase (21). That mRNA decay continued relatively normally even when four major ribonucleases were missing led to the search for additional genes that affect mRNA decay. In fact, as described above, it is now known that additional ribonucleases (RNase R, RNase G, RNase LS, RNase P, and RNase Z) play roles in mRNA decay.
However, the search for proteins involved in mRNA decay has been complicated by the fact that, although some mutations that affect mRNA decay also lead to cell inviability (rne, pnp rnb, rnpA, pnp rnr, orn), most do not (rnc, rng, rnb, rhlB, rraA, rraB, pcnB, rnlA, rnz). In addition, if a new mutant is identified, it is important to show that the gene product directly affects mRNA decay. Thus there have been no simple selection methods. Most of the new ribonucleases (RNase G, RNase Z, and RNase LS) and regulatory proteins (Hfq, RraA, RraB, CsrD) have been identified by using indirect approaches or have resulted from serendipity.
MrsC/HflB/FtsH.
An example of serendipity relates to an experiment in which a series of temperature-sensitive mutants were isolated in a screen for new pnp alleles (139). While no pnp alleles were identified, several new loci (messenger RNA stability) that affected mRNA decay were characterized. The first of these alleles (mrsC) encodes a Mg2+-dependent ATPase (395) (Table 1), whose gene was independently identified as a filamentation locus (ftsH [382]), as well as a protein involved in the transfer of proteins across the inner membrane (4) and high frequency of bacteriophage λ lysogenization (hflB [155]). Inactivating the MrsC/FtsH/HflB protein in the absence of RNase E (rne), RNase II (rnb), and polynucleotide phosphorylase (pnp) causes a rapid cessation of RNA synthesis and a dramatic increase in the stability of individual mRNAs such as trxA, cat, and kan (139). Of particular interest is that MrsC/FtsH/HflB protein has been shown to be membrane anchored through its amino terminus (381) and, in fact, is part of a very large multiprotein complex that also contains HflK and HflC (346). A variety of reports have demonstrated that the MrsC/FtsH/HflB protein functions as an ATP-dependent protease (5, 25, 140, 156, 190, 204). If this is the protein’s only catalytic activity, it suggests that proteolysis may be necessary to activate either an RNase or some other protein that is involved in mRNA decay and that this posttranscriptional processing occurs at the inner membrane.
MrsF/RrmJ.
A second locus identified in the experiment described in the previous section encoded a 23,204-dalton protein (Table 1) that was not essential for cell viability but was under heat shock control (44; F. Zheng and S. R. Kushner, unpublished results). It was called mrsF because strains carrying mutations in this protein showed alterations in mRNA stability. However, the mrsF locus, now called rrmJ, actually encodes a methyltransferase activity associated with the 2'-O-methylation of the universally conserved U2552 in the A loop of 23S rRNA (44, 48, 148). Thus the changes observed in mRNA half-lives probably arise as a secondary effect of a reduced rate of protein synthesis. Strains carrying a deletion of the rrmJ locus grow more slowly than wild-type controls and have significant defects in ribosome biogenesis (44, 48).
Even though a poly(A) polymerase activity had been identified in E. coli in 1962 (19), in general, it was assumed that polyadenylation was unique to eukaryotes (23, 207). The idea that E. coli mRNAs were modified posttranscriptionally first appeared in 1975 when two laboratories (283, 361) suggested that short poly(A) tails were on pulse-labeled RNA species in E. coli. In fact, from 1978 through 1992, Sarkar and her coworkers published a variety of reports on the existence of poly(A) tails on the RNAs isolated from E. coli (60, 135, 179, 347, 361) and B. subtilis (136). This work was ignored in general until cDNA sequencing showed that the lpp mRNA was polyadenylated (60). The poly(A) tails were attached either at the 3' end of the Rho-independent transcription termination or to a slightly shortened mRNA species in which the 3' stem-loop had been removed (60).
Experiments by O’Hara et al. (296) subsequently demonstrated that in wild-type E. coli the average poly(A) tail length was between 10 and 40 nt. In the absence of polynucleotide phosphorylase and RNase II, tail lengths increased to more than 100 nt (267, 296). Furthermore, more than 90% of the poly(A) tails were eliminated in the absence of poly(A) polymerase I (296), the product of the pcnB (59, 227, 243) gene.
Over the past decade, considerable progress has been made in understanding the mechanism of polyadenylation in E. coli. Mohanty and Kushner (264) showed that expression of the pcnB gene is controlled in part at the level of translation initiation, yielding very small amounts of the PAP I protein (estimated to be on the order of 30 molecules/cell [270]). The small number of PAP I molecules per cell in part explains the observation that only 1.3% of the total pulse-labeled RNA is polyadenylated at any given time (59). In fact, using a special construct in which the pcnB gene was under the control of the lac promoter, it was shown that in vivo poly(A) levels could be varied several hundredfold, leading to an increase in mRNA decay for a variety of transcripts (264). However, even though only a small percentage of E. coli mRNAs appear to be polyadenylated when tested using oligo(dT) analysis, macroarray experiments on the E. coli transcriptome have demonstrated that at least 90% of the transcripts in exponentially growing cells are polyadenylated to some extent (271). In fact, it seems likely that the fraction of mRNAs that are polyadenylated is much higher but that the tails are too short to detect by using standard procedures.
There is direct evidence that RNase II rapidly deadenylates the rpsO mRNA (120, 244). However, a more global analysis comparing the roles of PNPase and RNase II in the degradation of poly(A) tails has shown that PNPase is probably the more important enzyme (267). It has also been observed that poly(A) levels also play a role in the regulation of intracellular levels of PNPase and RNase E (268).
Other unusual aspects of E. coli polyadenylation have been identified. In the first place, rRNAs, in particular an immature 23S rRNA, are the primary target for polyadenylation in exponentially growing E. coli (264). Furthermore, polyadenylation not only is an important feature of mRNA decay but also appears to be involved in the turnover of defective stable RNA species (214, 217), thus serving as an important feature in RNA quality control. The addition of poly(A) tails to a variety of RNA substrates appears to function as a targeting signal for the binding of 3' → 5' exonucleases such as PNPase, RNase II, and RNase R. In fact, without polyadenylation, some naturally occurring transcripts, in particular ones containing stem-loop structures, are resistant to degradation (73, 186).
While it is now certain that polyadenylation is a targeting signal, other aspects of the polyadenylation story are still not entirely clear. For example, it was once thought that poly(A) polymerase acted alone and that almost any RNA molecule with a 3'-OH terminus could act as a substrate (118). However, recent experiments have suggested a more complicated story. While it was initially argued that the RNA binding protein Hfq reduced the preference of PAP I for transcripts terminated in a Rho-independent fashion (201), it has now been shown that Rho-independent transcription terminators (Fig. 1A) serve as polyadenylation signals, but only in the presence of the Hfq protein (270, 271) (Fig. 2). Furthermore, disagreement also exists regarding how Hfq functions in the process of polyadenylation. In one case, evidence has been presented that in vitro Hfq protects poly(A) tails from degradation by 3' → 5' exonucleases (151, 201), but the in vivo poly(A) sizing assays of Mohanty and Kushner (270) clearly show that this is not the case.
Another puzzling observation has been the nature of polyadenylation in the absence of PAP I. A variety of experiments have shown that PAP I is responsible for only 90% of the poly(A) observed in wild type E. coli (264). In fact, it was reported that E. coli contained a second poly(A) polymerase encoded by the f310 open reading frame (61, 175), but this work was placed in doubt by the observation that F310 protein did not have polyadenylation activity (265). Surprisingly, the residual polyadenylation has been accounted for by the biosynthetic activity associated with PNPase (266).
The tails synthesized by PNPase can be very long (>600 nt), contain a combination of all four nucleotides, and in general are found near the 5' end of mRNA species (266, 270) (B. Mohanty and S. R. Kushner, unpublished results). While it had been suggested from in vitro experiments that PNPase could be involved in the synthesis of poly(A) tails in E. coli (97, 325, 397), the prevailing view on the in vivo function of PNPase for many years was that it was involved in "...the salvage of nucleotides from RNA rather than to their polymerization to RNA." (196). This conclusion was supported by the fact that the intracellular PO4 concentration has been determined to be ~13 mM (356) and that the phosphorolysis reaction is optimal at 5 to 10 mM PO4 (133, 191). It now seems clear that phosphate concentrations low enough to facilitate the biosynthetic reaction of the enzyme can occur in vivo. Alternatively, the interactions of PNPase with other proteins such as RNase E or PAP I somehow alter its biosynthetic activity. Finally, note that the biosynthesis of polynucleotide tails by PNPase has now been observed in several other prokaryotes (343, 359), but it is not clear what role, if any, they play in mRNA decay.
While in general it has been thought that mRNA decay is a constitutively expressed salvage pathway in E. coli, the first evidence that this assumption was not correct was the discovery that RNase III regulates the levels of both PNPase and itself through cleavage of their respective mRNAs (27, 320, 321). In addition, RNase E regulates its own synthesis through the degradation of its mRNA (104, 105, 154, 164). Intracellular levels of RNase E are also controlled by transcription from multiple promoters (305) and intracellular poly(A) levels (268).
When coupled with the data showing an interrelationship between the intracellular levels of PNPase and RNase II (46, 414), autoregulation of PNPase (167, 248), and the control of RNase R levels by RNase E (47), it has become increasingly clear that the mRNA decay capacity of the cell is carefully regulated. Further support for this idea is derived from the fact that overexpression of either RNase E or PAP I results in the loss of cell viability.
Besides the control of many of the various mRNA decay enzymes by a variety of posttranscriptional mechanisms, the physical structure of each mRNA molecule at the 5' and 3' ends (as discussed in "Structural Properties of mRNA Molecules That Can Affect Stability," above) also serves to regulate decay. In fact, additional structures probably exist within the translated regions of mRNAs that are also important for decay.
Taken together, each mRNA most likely has programmed into it susceptibility to several possible decay pathways depending on the presence of recognition sites for the various endoribonucleases (RNase E, RNase G, RNase Z, RNase III, RNase P, and RNase LS). Based on microarray data and specific half-life measurements (30, 31, 202, 306, 316), it appears that each mRNA can be degraded by either a primary or possibly multiple secondary pathways. If a primary pathway is inactivated, one or more secondary mechanisms fill in. For example, it has been well documented that the rpsT mRNA decays in an RNase E-dependent fashion (239). However, the half-life of the rpsT mRNA increases significantly in both rne-1 rng::cat and rne-1 Δrnz mutants (306, 316). In most cases, the presence of alternative decay pathways is probably sufficient for the cell to function, thereby explaining why some mutants, in which mRNA decay is severely defective (e.g., rneΔ610, rneΔ645, and rne-131 [233, 304]), still remain viable. Most likely, only in special cases will the inability to degrade specific mRNAs be sufficiently deleterious to result in cell death. Finally, it seems unlikely that every molecule of a particular mRNA will be degraded in the exact same fashion.
Although many studies have suggested that growth rates do not influence the stability of most transcripts (40, 87, 125, 126, 309), a direct relationship between growth and the stability of the ompA and cat mRNAs has been reported (256, 290). In addition, Meyer and Schottel (258) showed that cat transcript stability was growth rate independent but varied with growth-medium composition. Nilsson et al. (290) also showed that the bla and lpp mRNAs were not growth medium dependent. Additional studies are apparently needed to determine how important growth rate and medium composition are to mRNA stability.
Since mRNAs were discovered in 1961 (41, 141, 163), it has been assumed that the process of translation would stabilize mRNAs by protecting them from nucleolytic attack. Several lines of evidence have served to support this view. If antibiotics are used to stall ribosomes on the mRNA, stability increases (93, 222, 353). When termination codons or feedback inhibition of ribosomal protein synthesis is used to reduce the number of ribosomes on an mRNA (88, 116, 290), stability decreases. Kennell and Reizman (184) also observed a correlation between translation efficiency and mRNA stability in the lac operon.
Conflicting evidence, however, has also been reported. Mackie (238) noted that translational feedback regulation of ribosomal protein S20 led to the stabilization of the mRNA. In addition, von Gabain et al. (390) showed no differences in the stability of translated and untranslated segments of both the bla and ompA transcripts. Results of experiments using mRNAs with mutations in the leader or proximal part of a gene to alter the protein output of the mRNA suggested no clear explanation. A correlation between translation efficiency and mRNA stability was found in some cases (78, 399) but not others (55, 113, 317).
McCormick et al. (249) reexamined the question of translation and mRNA stability. For the fusion mRNAs they tested (a combination of ribosomal protein S10 and β-gal), they concluded that, as translation decreased, mRNA degradation increased (249). Additional experiments employing the lac mRNA have provided a clearer indication that efficient translation effectively increases mRNA stability (161, 171, 194).
In another approach, Lopez et al. (232) examined mRNA stability in the presence of translation inhibitors by using bacteriophage T7 RNA polymerase to synthesize a lacZ mRNA. They argued that drug-induced translation arrest might lead to the saturation of the degradation machinery by increasing the levels of rRNAs that need to be degraded (232). This would in effect tie up both RNase E and PNPase under conditions in which their autoregulation could not lead to increased levels of either nuclease (232). For a more detailed analysis of this subject the reader is directed to two recent reviews on this subject (96, 173).
Note also that most studies on mRNA decay in E. coli have dealt strictly with exponentially growing cells. However, in nature, the organism is frequently subjected to stress conditions, including both nutrient starvation and cold shock. In turns out that under these circumstances specialized systems are activated to rapidly restrict the translatability of mRNAs by cleaving transcripts that are already associated with ribosomes. The initial cleavage events in these systems appear to be independent of the activity of the endoribonucleases that were described in "Genetics and Biochemistry of Proteins and Enzymes Involved in mRNA Decay," above.
This approach to mRNA decay has been called "shutdown decay" (89) and involves a series of so-called toxin-antitoxin systems that are thought to allow cells to rapidly respond to altered growth conditions. These systems work through the synthesis of two proteins, one designated as the toxin and the other the antitoxin. Under nonstress conditions, the two proteins interact with each other and the cells grow normally. Under stress conditions, the antitoxin is preferentially degraded, leading to a rapid, if temporal, activation of the toxin protein. The classic system in E. coli, associated with the RelE and RelB proteins (79), displays codon-specific cleavage of mRNAs in the ribosomal A site by RelE (310). Additional toxin-antitoxin systems that have been described include mazEF (409, 410), chpBK (411), and yoeB/yefM (80, 177). The ribonucleases encoded by these three systems seem to be able to cleave mRNAs either in between ribosomes or at upstream positions. In addition, it is not clear whether a nuclease activity exists that is an intrinsic part of the ribosome that becomes activated under certain circumstances. A cleavage event that occurs during translation will generate a nonstop RNA, which is probably a target for the tmRNA trans-translation system (see Chapter 4.2.7), leading to more rapid decay of the transcript (209, 336). A more detailed review of these systems by Condon (89) has recently been published.
Over twenty years ago Jain et al. (166) suggested the existence of a multiprotein complex that contained RNase E, RNase P, and RNase III. However, because of technical flaws associated with these experiments, it was not until the elegant work of Carpousis et al. (63) and others (323) that the RNase E-based degradosome was discovered. This complex has subsequently been characterized in more detail (50, 259, 324, 387). The presence of a multiprotein complex containing both endo- and exonucleolytic activities (RNase E and PNPase, respectively) along with a RNA helicase to unwind secondary structures (RhlB) provided a straightforward explanation for the rapidity with which many mRNAs were degraded (84, 327). The question of intracellular localization of this protein "machine" was first addressed using immunogold localization (223). These authors concluded that the degradosome was located near the inner cell membrane (223).
Although the technical limitations of this procedure made the results not totally compelling, recent proteomic analysis of the cell envelope fraction has found that RNase E, RhlB, enolase, PNPase, RNase II, and RNase III are all present (121). Although these experiments contained several internal controls and detected several proteins (HflB/MrsC, OmpA, OmpC, and OmpF) expected to be associated with the membrane, the presence of many ribosomal proteins and the LacI repressor protein (121) raised the concern that some of the proteins were present due to nonspecific interactions. However, new data suggest that the degradosome is organized as filamentous helical structures that coil around the length of the cell and act as components of the bacterial cytoskeleton (371). The authors hypothesized a compartmentalization of the major mRNA degrading apparatus at the inner membrane (371). They also noted that the RNase E protein encoded by the rneΔ645 allele (304) fails to form the helical structures (371). These new results also provide a potential explanation for how the HflB protease that is anchored in the inner membrane (395) plays a role in mRNA decay. Finally, note that there is one documented example of the membrane localization for the decay of a specific mRNA (180).
An exciting new area of research in E. coli is the role of small RNAs (sRNA) in control of a variety of genes. Several detailed reviews of this topic have recently appeared (138, 242). Many of the small RNAs function through direct RNA/RNA hybridization that leads to the rapid degradation of both the small RNA and its mRNA target (138). For example, the RhyB RNA is an antisense regulator whose expression is repressed by the Fur repressor and regulates the stability of at least six mRNAs (e.g., sdhCDAB, fumA, acnA, sodB, ftnA, and bfr) encoding Fe-binding or Fe-storage proteins in E. coli (246). The RyhB RNA is stabilized in the presence of Hfq and is degraded in an RNase E-dependent fashion. Under stress conditions the synthesis of RhyB increases and it pairs with its target mRNAs. Once the sRNA is paired with its target mRNA, both molecules are degraded in an RNase E-initiated reaction (247).
Similar results have been seen with other small RNA regulators (2, 272). While the exact mechanism of this reaction is not well understood, particularly in light of the preference of RNase E for single-stranded regions (see previous discussion of RNase E), the reported interaction between Hfq, which binds to a large number of small RNAs (138), and RNase E provides a possible mechanism of how the levels of some of these RNAs are controlled (275). It thus appears that this system provides another level of regulation that can promote the rapid degradation of specific mRNAs when it becomes necessary to adapt to changes in growth conditions and provides a prokaryotic parallel to the RNA interference systems observed in eukaryotes.
Over 30 years ago Apirion (9) hypothesized, long before most of the nucleases and proteins described in this review had been identified, that mRNA decay in E. coli proceeded by a series of endo- and exonucleolytic steps. It turns out that, although there are multiple decay pathways in E. coli, his original idea was correct. In fact, several different types of array experiments have suggested that only a small percentage of E. coli transcripts are degraded exclusively in an exonucleolytic fashion (31, 269). Thus the majority of mRNA decay is initiated via endonucleolytic cleavages carried out by at least six different endonucleases (RNase E, RNase G, RNase III, RNase P, RNase Z, and RNase LS) as well as the stress-specific enzymes such as RelE and MazF.
In addition, since E. coli and, presumably, its close cousin serovar Typhimurium apparently do not have 5'-3' riboexonucleases like yeast (62, 366), RNase II, PNPase, and RNase R (all 3' → 5' enzymes) are the primary means for generating mononucleotides and nucleoside diphosphates that can either be recycled into new RNA molecules or converted into deoxynucleotides by nucleotide reductase and then funneled into DNA synthesis. In this regard the involvement of a phosphorolytic activity in mRNA decay has advantages because it generates mononucleotide diphosphates (99). Danchin (94) has even argued that a major role for PNPase is to generate CDP that is subsequently converted into dCTP for use in DNA synthesis. The following paragraphs outline an attempt to generate a series of comprehensive models of mRNA decay that integrate the current knowledge in the field.
A variety of array data clearly indicate that RNase E is the primary endonuclease involved in the initiation of mRNA decay for numerous E. coli mRNAs (31, 202). However, genetic analysis has also shown that five other endoribonucleases (RNase G, RNase P, RNase III, RNase Z, and RNase LS) are also involved (210, 302, 306, 316, 338) but primarily in backup roles. One model that can explain these observations is shown in Fig. 4A and B. A major assumption in this model is that RNase E is the most abundant endonuclease in the cell. In fact, it appears that E. coli actually has excess RNase E in that changes in mRNA decay rates are only observed when enzyme levels are reduced by more than 80% (165, 305). Additional assumptions are that RNase III has the most stringent cleavage specificity, requiring the presence of specific stem-loop structures (Fig. 1B), and RNase P has a very limited role because presumably only a very small subset of mRNAs can fit into the active site of the catalytic M1 RNA subunit.
Thus the model assumes that RNase E will normally initiate the decay of both monocistronic (Fig. 4A) and polycistronic mRNAs (Fig. 4B) by binding to the 5' terminus (240, 380). While its homologue RNase G can also do this (380), it is much less likely because of the relatively low levels of the protein within the cell (42) and the fact that it binds very poorly to a 5'-triphosphate (380). Even though RNase E is also inhibited by a 5'-triphosphate (241), it has been hypothesized that nonspecific phosphodiesterases may remove the 5'-triphosphate or, if not, that this is the rate-limiting step in mRNA decay. The ability of the 5' UTR of ompA to stabilize transcripts (113, 114) through a double-stranded structure at the 5' end provides further support for the binding of RNase E at the terminus.
Once RNase E binds, its first cleavage reaction takes place at a downstream site, generating a new 5' terminus with a phosphomonoester (Fig. 4A and B). Based on the proposed multimeric nature of RNase E, a second RNase E subunit binds the new downstream 5' terminus (Fig. 5). Since the enzyme is stimulated by 5'-phosphomonoesters (169, 240) the degradation process will be accelerated. In some cases, the binding of RNase E to a particular mRNA may preclude another endonuclease from binding due to the proximity of the cleavage sites and the physical size of the RNase E-based degradosome or its actual location within the cell (Fig. 4A and B). If, however, an RNase E-generated decay intermediate contains an RNase G, RNase Z, or RNase LS site, these enzymes, at this point, may cleave to generate even shorter decay intermediates (Fig. 4A and B). In the absence of RNase E, message decay is slowed but is not completely blocked because the presence of RNase G, RNase LS, and RNase Z can initiate decay at reduced efficiency (Fig. 4A and B). The end result of the action of the six endonucleases is a series of decay intermediates that are rapidly degraded by a combination of PNPase, RNase II, and RNase R (Fig. 6). Those intermediates or sometimes full-length transcripts containing secondary structures at their 3' termini are polyadenylated by PAP I (Fig. 2).
An interesting question raised by the characterization of the RNase E-based degradosome is whether mRNAs are degraded simultaneously from both the 5' and 3' ends. This highly speculative model invokes the recent finding of degradosome filaments (371) and multimeric forms of RNase E (50, 52, 169). In this model, RNase E would bind to the 5' terminus while the PNPase component of the degradosome would bind to the 3' end (Fig. 5). After the first cleavage event, the upstream RNA fragment would be released and the new 5' terminus would be transferred to an adjacent RNA binding site in the filament (Fig. 5). By passing an mRNA around the filament all of the RNase E cleavages could be rapidly completed. It is not clear whether PNPase or RNase E would bind to the mRNA first, but such a model could explain the very rapid disappearance of full-length mRNAs with the concomitant appearance of decay intermediates. In fact, decay intermediates have in general been visualized only in multiple mutants lacking RNase E, PNPase, and RNase II (14, 16, 17).
Note, however, that mRNA decay is almost as efficient even in the absence of a functional degradosome (303) but that mutants containing very short forms of the protein are very defective in mRNA decay (304). Thus while the degradosome clearly functions in wild-type cells, mRNA decay is still relatively efficient in its absence. In addition, many gram-negative bacteria contain only the catalytic portion of RNase E.
While it is accepted that polyadenylation plays a role in the decay of mRNAs (108), there is still considerable debate whether polyadenylation is required for the initiation of decay of full-length transcripts. In the case of mRNAs that end in a Rho-independent transcription termination such as lpp, the data are quite clear that polyadenylation plays a role in the initiation of mRNA decay (264), in particular, since it does not appear that this transcript is a substrate for RNase E (303). Since it is also known that all three exonucleases (PNPase, RNase II, and RNase R) require unstructured 3' termini for binding (73), polyadenylation in this case would be a critical first step for the initiation of decay (Fig. 2). In addition, the binding of PNPase may actually bring RNase E to a substrate that it might otherwise be blocked from binding initially because of a paired 5' terminus (ompA). Besides this role in the initiation of decay, polyadenylation clearly is required for the degradation of decay intermediates (see next section) that contain extensive secondary structures (73, 186).
A variety of genetic evidence indicates that mRNA decay intermediates generated by the endonucleases discussed above are degraded by a combination of PNPase, RNase II, and RNase R (73, 74, 107, 269). As shown in Fig. 6, in combination with PAP I, the decay intermediates are rapidly degraded into mononucleotides and nucleoside diphosphates. The short oligonucleotides (2 to 5 nt in length) that are no longer substrates for these enzymes are substrates for oligoribonuclease. The posttranscriptional addition of polynucleotide tails on short mRNA decay products (266, 270) results from the biosynthetic activity of PNPase. The function of these polynucleotide tails is currently not understood.
Since some of the endonucleases (RNase E, RNase III, RNase P, and perhaps others) involved in mRNA decay also participate in ribosomal RNA processing and tRNA maturation, it is worth considering the interrelationship among these three processes. Ribosomal RNAs and many tRNAs are transcribed from seven ribosomal RNA operons as polycistronic molecules. The action of RNase III, RNase E, RNase P, and many exoribonucleases (see references 103 and 213 for reviews and a comparison of mRNA decay and tRNA processing) convert the 30S transcripts into mature 5S, 16S, 23S, and tRNA species. Those tRNAs that are independently transcribed are also matured through the combined action of RNase E, RNase P, and several exoribonucleases (216, 304). While the absence of RNase III does not affect cell viability (21, 373), inactivating RNase E by using either the rne-1 or rne-3071 alleles leads to the accumulation of 9S rRNA precursors, immature tRNA precursors, and subsequent cell death (10, 11, 216, 304).
The take home lesson is that E. coli has a finite number of ribonucleases that can both process and mature noncoding RNAs and, under slightly different circumstances, degrade mRNAs. In fact, the high demand for functional ribosomes during exponential growth would indicate that tRNA maturation and rRNA processing must occur more rapidly than the decay of most mRNAs. Not surprisingly, the half-lives of tRNA precursors are so short that they cannot be measured in wild-type E. coli (304) and have been hypothesized to be less than 30 s (S. R. Kushner, unpublished results). Likewise, Li and Altman (212) have estimated that RNase P processes tRNA substrates 10-fold faster than it cleaves in the lacY lacA intercistronic region. Furthermore, the assembly of functional ribosomes may also be relatively flexible, allowing altered forms of 16S, 23S, and 5S structural RNAs to be incorporated. In contrast, the accumulation of partially degraded mRNA species may lead to problems by tying up ribosomes synthesizing nonfunctional truncated proteins. Thus it is important to remember that perturbation of any aspect of E. coli RNA metabolism is going to have global consequences.
Many of the ribonucleases found in E. coli have homologues in a large number of prokaryotes. However, some enzymes very important in E. coli (RNase E) are mainly found in gram-negative organisms. Others such as PNPase are ubiquitous in both gram-negative and gram-positive bacteria. While the actual participants in mRNA decay may vary from one prokaryote to another, the general features of a combination of endo- and exonucleolytic decay seem to hold. For a more detailed discussion of this topic see the recent review by Kushner (199).
There are still many unanswered questions regarding both the mechanism and importance of mRNA decay. Some of these are listed below.
Are there additional ribonucleases involved in mRNA decay?
Is there a specific enzyme that converts the 5'-terminal triphosphate into a monophosphate, thereby facilitating the binding of either RNase E or RNase G?
What is the exact role of polyadenylation in mRNA decay?
What is the complete composition of the PNPase/RNase E multiprotein complex?
What is the biochemical mechanism of degradosome-initiated mRNA decay?
How important are RNA/RNA interactions in mRNA decay?
Is there a second multiprotein complex involved in poly(A)-dependent mRNA decay?
What is the exact relationship of the inner membrane and mRNA decay?
How is mRNA decay regulated, in particular, during various stages of the cell cycle?
Why is oligoribonuclease an essential enzyme?
Is mRNA decay really essential for cell viability?
How general is the phenomenon of RNA/RNA interactions in the control of mRNA decay?
These questions represent only a few of the important issues yet to be resolved before mRNA decay in E. coli and serovar Typhimurium can be truly understood.
This work was supported in part by a grant from the National Institute of General Medical Sciences (GM57220) to S.R.K.
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Module4.6.4
