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Polyadenylation and Degradation of RNA in Prokaryotes

and .

Summary

Polyadenylation is a postranscriptional modification of RNA found in all cells and in organelles. In bacteria, a small fraction of RNA harbors oligo(A) tails which are mostly shorter than 20 As. Poly(A) polymerase I of Escherichia coli can adenylate mRNAs, and small RNA regulators originating from the chromosome, from plasmids and from bacteriophages and also precursors and mature forms of tRNAs and rRNAs which have an accessible 3' extremity. The model of poly(A) metabolism presented in this chapter proposes that the ratio of adenylated to nonadenylated RNAs and the length of oligo(A) tails represent the equilibrium of the antagonistic activities of poly(A) polymerase I, which synthesizes poly(A) at the 3' end of all RNAs, and of exoribonucleases which shorten or completely remove the tails. Poly(A) tails provide toeholds where polynucleotide phosphorylase can initiate exonucleolytic degradation of tightly folded RNAs protected from exoribonucleases by 3' stable secondary structures. Polyadenylation promotes degradation of mRNA fragments and controls the intracellular concentration of regulatory RNAs.

Introduction

Poly(A) polymerase, the enzyme responsible for the synthesis of poly(A) tails of RNA was discovered in bacteria long before the identification of polyadenylated mRNAs in eukaryotic cells.1 The length of poly(A) tails (60 to 200 As) and the abundance of polydenylated mRNA in eukaryotic cells (nearly all mRNAs are adenylated) compared to the situation in bacteria where only a small fraction of RNA harbors short tails led to the assumption that polyadenylation was specific for eukaryotic cells and that it has only a minor function in the metabolism of bacterial mRNA. Eukaryotic polyadenylation is described by T. Preiss in chapter 12 . More recently, several important discoveries such as the characterization of the Escherichia coli poly(A) polymerase (PAP I), its gene (pcnB) and its implication in the control of mRNA stability renewed the interest of researchers for polyadenylation of RNAs in bacteria.2 The literature relating the development of this topic has been recently reviewed.3,4 This chapter focuses primarily on the metabolism of poly(A) tails and on the role of polyadenylation in the control of mRNA stability in E. coli.

Characterization of Poly(A) Tails

The repeated observation that pulse labeled RNAs isolated from E. coli and several other eubacteria and archaea could be retained on oligo(dT) cellulose was the first indication that polyadenylation takes place in prokaryotes.3 However, the fraction of polyadenylated molecules was usually very low (less than 3% of total labeled E. coli RNA was retained on oligo(dT) cellulose)5,6 and attempts to determine the length of tails gave very heterogeneous data. Digestion of polyadenylated RNA by RNase T1 and RNase A cleaving, respectively, the GpN, CpN and UpN phosphodiester bonds generates poly(A) tracts whose length can be determined by electrophoresis or from the adenosine:AMP ratio after alkaline hydrolysis. Poly(A) tails ranging from 1 7,8 to 50 As5,9 were detected in E. coli. It must be pointed out, however, that many of these data, based on the binding of polyadenylated RNA to oligo(dT),3 probably only give approximate idea of the polyadenylation status of bacterial RNA since they did not take in account RNAs harboring tails of less than 20 As that are not retained on oligo(dT). Investigations of specific transcripts confirmed that only a small fraction of bacterial RNAs harbors poly(A) tails which are mostly less than 20 As in length whereas the majority of molecules are not adenylated. Based on the fraction of RNA retained on oligo(dT), it was estimated that only 0.011% of bacteriophage f1 mRNA is polyadenylated 10 whereas 40% of lpp and trpA mRNAs harbor poly(A) tails of 10–20 As.11,12 On the other hand, analysis of mRNAs on Northern blot showed that the majority of rpsO primary transcripts are not adenylated with only 10% of these molecules harboring short tails of 1–3 nucleotides (P. Marujo, E. Hajnsdorf and P. Régnier, unpublished).8 In contrast to eukaryotic cells, polyadenylation is not restricted to mRNA in bacteria (Table 1).1 Reverse transcription experiments using a primer complementary to an oligonucleotide ligated to the 3' end of RNA molecules demonstrated that a few As were post-transcriptionally added at the 3' end of RNA I, the regulator of Col E1 type plasmid replication.14 Genomic RNA from bacteriophage MS2,15 the small oop RNA of bacteriophage λ,16 5S rRNA17 and precursors of 16S and 23S rRNAs18 are also polyadenylated under normal physiological conditions. Moreover, many other RNAs including the mature 23S rRNA,18 tRNATyr,17 the tmRNA,17 the small sok regulatory RNA19 and the 6S, 4.5S and M1 small RNAs17 can be polyadenylated in strains overproducing PAP I or lacking exonucleases able to degrade single-stranded 3' ends of RNAs (Table 1) (see below). These RNAs have very short tails (Table 1). Not only are stable, regulatory and messenger RNAs polyadenylated but most of them can be adenylated at many different sites resulting from transcription termination, endonucleolytic processing by RNase E and RNase III and digestion by exonucleases engaged in trimming of stable RNAs or degradation of mRNAs.4,10, 18,20 Finally, some regulatory and messenger RNAs (copA, ompA, trxA) stabilized upon PAP I inactivation are also presumed to be polyadenylated (Table 1) (see below).9, 21 It therefore appears that PAP I can polyadenylate all classes of bacterial RNA but that only a small fraction of molecules, that may be different from an RNA species to another, is actually polyadenylated.

Table 1. Polyadenylated bacterial RNAs.

Table 1

Polyadenylated bacterial RNAs.

Enzymes of Poly(A) Metabolism

In E. coli, the main enzymes of poly(A) metabolism are PAP I, which accounts for the synthesis of most, if not all, poly(A) tails and exonucleases of 3' to 5' polarity, capable of degrading singlestranded stretches of nucleotides found at the 3' end of RNAs.23,23PAP I is encoded by the pcnB gene and is dispensable for growth.2426 It is a monomer of approximatively 53 kDa 24,27 with characteristic features of the nucleotidyltransferase superfamily including eukaryotic poly(A) polymerases as well as eukaryotic and prokaryotic tRNA nucleotidyltransferases, which generate the CCA 3' extremity of tRNAs.28,29 PAP I requires a divalent cation, Mg2– or Mn2–, to be active.1,30 The enzyme uses ATP as substrate to polymerize AMP residues at the 3' end of RNA primers. CTP and UTP can also be used by PAP I but at a much lower rate (at about 5% the rate of ATP incorporation).30,31 Synthesis of poly(G) tails by PAP I has also been reported31but this reaction is probably very slow.30 These properties may explain why Cs are sometime incorporated in poly(A) tails (E. Hajnsdorf and P. Régnier, unpublished). The Km values for ATP and tRNA primers are 50 μM and 0.2 μM, respectively.30 The fact that poly(A) tails are synthesized at the 3' ends of many primary transcripts, processed RNAs and intermediary products of exonucleolytic degradation none of which share common 3' terminal features, suggests that PAP I does not specifically recognize structural motifs of particular RNAs.10,18,20 However, the variability of the length of tails mentioned above suggests that all RNA species are not adenylated with equal efficiency. Moreover, it has been shown that PAP I preferentially adenylates RNAs primers harboring a 5' monophosphorylated extremity or single-stranded stretches of nucleotides at the 3' or at the 5' end.31,32 The influence of 5' structures on polyadenylation efficiency could explain why truncation at the 5' end of RNA increases sensitivity to poly(A)-dependent degradation.33 PAP I is a distributive enzyme (it dissociates from RNAs after addition of each or a few nucleotides) which exhibits a preference for poly(A) primers.34 Its activity is modified by protein Hfq which interacts with A-rich regions of RNAs35,36 and is involved in replication of the single-stranded genomic RNA of bacteriophage Qβ37 and in the translation of the ompA messenger38 and of the ss subunit of RNA polymerase, specific of stationary phase and osmotic upshift.39 Hfq stimulates poly(A) synthesis in vivo and in vitro and converts PAP I into a processive enzyme which remains associated to the molecules that it elongates.40 Other potential cofactors interacting with PAP I are RNase E, which is involved in RNA decay, and DEAD box RNA helicases, which may cooperate with PAP I for poly(A) synthesis.41 The fact that PAP I competes with exonucleases capable of degrading poly(A) tails may explain why the long tails of 500–1000As that can be synthesized by PAP I in vitro are not detected in the cell.40,42

E. coli contains eight 3' to 5' exonucleases capable of degrading single-stranded RNAs or oligonucleotides that may all be involved in degradation of poly(A).43,44 The fact that poly(A) is much more abundant when two of these enzymes, RNase II and polynucleotide phosphorylase (PNPase), are inactive while inactivation of only one of them does not have such an effect suggests that each of these enzymes can degrade poly(A) tails.8,4548 RNase II is a processive hydrolase (it remains bound to the molecules that it degrades) which generates ribonucleotides monophosphate.49 The abundance of this enzyme, which represents 90% of the poly(A) degrading activity measured in E. coli cellular extracts, suggests that it plays a major role in poly(A) catabolism.50 Consistent with this idea, it has been shown that RNase II removes the poly(A) tails synthesized by PAP I at the 3' end of rpsO primary transcripts almost completely thus explaining why 90% of these molecules are unadenylated P. Marujo and P. Régnier, unpublished).8 Processive removal of nucleotides by RNase II is blocked by stable secondary structures which cause dissociation of the enzyme from the RNA.5155 This property explains why RNase II removes single-stranded stretches of nucleotides and poly(A) tails lying downstream of transcription terminators or of REP sequences but fails to degrade RNAs upstream of these hairpins.8,52,5659 RNAs devoid of 3' secondary structures are probably completely degraded by RNase II.33,60

PNPase is a phosphorylase which attacks single-stranded 3' end of RNAs that it degrades processively into monoribonucleotides diphosphates (NDP).61 In the reverse reaction, which takes place at low phosphate concentration, PNPase processively incorporates NDP into heterogenous polynucleotides of random sequences.61 However, its primary role in the cell is the 3' to 5' degradation of RNA molecules62 and it is one of the components of a multienzymatic complex refered to as the RNA degradosome which participates in mRNA decay (see Chapter 9, by RK Beran et al).63,64 Although PNPase is blocked by secondary structures, like RNase II, it is capable of degrading highly structured RNA that are resistant to this latter enzyme.5254,56,65 RNAs whose 3' terminal nucleotides are sequestered in a secondary structure are poorly attacked by PNPase free or associated with the degradosome.6668 In contrast, this enzyme can use single-stranded stretches of nucleotides lying downstream of stable 3' terminal secondary structures (for example transcriptional terminators) as a toehold to begin the exonucleolytic degradation of such RNAs.66, 67,69 A sequence of five As downstream of a hairpin is sufficient to initiate degradation.66 The mechanism of degradation of folded RNA by PNPase is described below. Beside its primary role in poly(A) and RNA degradation, it has been proposed that PNPase might account for the synthesis of the few poly(A) tracts which have been detected in PAP I deficient cells.48 It has also been suggested that PNPase synthesizes heterogenous tails (containing As, Us, Cs and Gs) in vivo in the absence of PAP I and incorporates U and C residues in poly(A) tails.48 Conversion of PNPase activity from phosphorolysis to polymerization and the reverse might be governed by transient modifications of local phosphate concentration at the 3' end of RNAs. The physiological significance of these alternate phases of RNA elongation and shortening remains mysterious.

RNase E plays a major role in mRNA decay in addition to being involved in the maturation of 5S and 16S rRNA.70,71 It is the scaffold for the association of PNPase, the RhlB RNA helicase and enolase, a glycolytic enzyme, in the RNA degradosome.63,64,67,72 It is a 5' end dependent endoribonuclease which preferentially cleaves RNAs harboring single-stranded monophosphorylated 5' extremities.73 This enzyme, which frequently initiates RNA decay, cleaves molecules in singlestranded A-U rich sequences70,71 and removes tails of polyadenylated molecules in vitro.74,75

In addition to PNPase and RNase II, E. coli contains five additional 3' to 5' exoribonucleases, RNase T, RNase PH, RNase D, RNase BN and RNase R, capable of removing single-stranded nucleotides from the 3' extremity of RNAs, that have been implicated in the maturation of many stable and small RNAs.44,7678 These enzymes, which are also blocked by secondary structures, could contribute to the degradation of oligo(A) tails added at the 3' ends of mRNAs, regulatory RNAs and precursors of stable RNAs. It appears however that they cannot counteract the synthesis of long poly(A) tails by PAP I.8,46,47

A Model of Poly(A) Metabolism

The appearance of long poly(A) tails, not detected under normal conditions, in cells either lacking the two 3' to 5' exoribonucleases PNPase and RNase II or overproducing PAP I, led to the conclusion that the length of bacterial poly(A) tails is determined by a dynamic equilibrium between the opposing activities of PAP I and exoribonucleases.8,17,22,56 Most, if not all, 3' RNA extremities accessible to PAP I can likely be adenylated. Stable RNAs such as tRNAs and rRNAs are nearly never subject to polyadenylation probably because their 3' termini are masked by aminoacylation or burial within the ribonucleoparticles.17 In contrast, PAP I polyadenylates the 3' extremities of abnormal tRNA precursors (M. Deutscher, personnal communication) that cannot be aminoacylated and of precursors of the 16S and 23S rRNAs that might emerge from the surface of the ribosome.18 Moreover, tightly folded RNAs with a triphosphorylated 5' extremity and lacking terminal single-stranded stretches of nucleotides may be poorly adenylated.31,32 The fact that PAP I is a distributive enzyme, which dissociates from the RNA primer after addition of one or a few adenosine residue(s), implies that 3' ends of tails become accessible to exoribonucleases after each (or a few) step(s) of adenylation. If long tails have been synthesized, RNase II or PNPase can carry out rapid processive degradation of poly(A) until it encounters stable stem-loops which cause stalling and dissociation of both ribonucleases (Fig. 1A). The processive degradation probably produces RNAs with single-stranded stretches of about 9 nucleotides downstream of the hairpin.51 These molecules and other RNA harboring short poly(A) tails are probably degraded distributively, at a slower rate, by RNase II and PNPase to generate short tails of 1–3 As such as those found in the RNA I and rpsO transcripts.8,14 In the case of the rpsO mRNA, RNase II can completely remove the tail while PNPase produces RNAs harboring tails of 2–4 As (Fig. 1A).8 More generally, RNase II can remove single-stranded nucleotides very close to the base of secondary structures that are not accessible to PNPase.52 3' terminal hairpins of different structures and stabilities may block 3' to 5' exoribonucleases at different positions and thus determine the number of As left at the 3' end of the RNA.53 For example, exonucleases can probably remove As very close to hairpins containing breathing A-U base pairs at the base of the stem while these As may not be accessible if the hairpin is closed by G-C base pairs.8,53 Such exoribonuclease-hairpin interactions could explain why the lpp transcripts harbor tails of 10–15 As.12 Moreover, the other 3' to 5' exonucleases that are involved in 3' trimming of stable RNA precursors may also contribute to the nibbling of messengers and generate tails of different lengths depending on their capability to approach secondary structures.77 On the other hand, it has been suggested that the poly(A) tails of some RNAs may be specifically degraded by PNPase or RNase II.23

Figure 1. Metabolism of poly(A) in bacteria.

Figure 1

Metabolism of poly(A) in bacteria. A) This model accounts for the synthesis of oligo(A) tails at the 3' end of the rpsO mRNA coding for ribosomal protein S15 in vivo. Transcription terminates in the UCA sequence at the bottom of the hairpin of the transcription (more...)

The model described above, which postulates that 3' to 5' processive exonucleases can initiate removal of poly(A) tails after addition of each (or a few) A residue, implies that these enzymes counteract poly(A) tail elongation and therefore prevent the synthesis of long tails. It has therefore been proposed that cofactors similar to mammalian CPSF (Cleavage and Polyadenylation Specifity Factor) and PABP II (a poly(A)-binding protein)79 may cause processive synthesis and ensure the protection of the long poly(A) tails of up to 50 nucleotides in length that have been detected in E. coli.. 9 One possible candidate was the host factor Hfq which binds A-rich RNAs (see above). The fact that the poly(A) tails of the rpsO mRNAs are slightly shorter in Hfq deficient cells and that Hfq stimulates elongation of poly(A) and converts PAP I into a processive enzyme in vitro strongly suggests that it affects poly(A) synthesis in vivo (Fig. 1A).40 Two other poly(A)-binding proteins may also affect poly(A) metabolism. CspE prevents poly(A) removal in vitro and ribosomal protein S1 can interact physically with two enzymes of poly(A) metabolism, PNPase and RNase E.80 It has been proposed that an endonucleolytic cleavage by RNase E, close to the RNA-poly(A) junction may remove poly(A) tails protected by poly(A)-binding proteins (Fig. 1B).75 However there is no evidence that cofactors such as CspE and ribosomal protein S1 affects poly(A) metabolism in vivo. Moreover, the fact that tails longer than 20 As are unfrequently detected in E. coli suggests that they may not be many poly(A)-binding proteins which protect tails from exoribonucleases in prokaryotes. More likely, these long tails could result from the dynamic equilibrium between synthesis by PAP I and Hfq and degradation by 3' to 5' exoribonucleases and probably correspond to a limited number of molecules which can undergo many successive steps of elongation without being attacked by exoribonucleases. One could also imagine that only few RNA species that are preferentially polyadenylated can gain long poly(A) tails.

Functions of Polyadenylation

The intracellular concentration of several small RNA regulators of different physiological functions is poly(A)-dependent. Originally, a mutation in the pcnB gene coding for PAP I reduced the copy number of ColE 1 type plasmids whose replication is negatively controled by RNA I.25 RNA I is a 108 nucleotide antisense molecule which forms a duplex with the RNA II primer of DNA replication. Inhibition of pBR322 DNA replication in the absence of PAP I was shown to result from the accumulation of an RNase E cleavage product of RNA I, lacking the five 5' nucleotides of the primary transcript, which must be polyadenylated to be degraded.14 The other small regulatory RNAs whose stability and intracellular concentration also depend on polyadenylation, namely copA, sok and oop, control the replication and maintenance of plasmid R1 and the lysogeny of bacteriophage λ, respectively.19,21,81 As in the case of RNA I, polyadenylation destabilizes truncated copA and sok molecules generated by RNase E.

In addition to the small RNA regulators mentioned above, polyadenylation also contributes to the rapid degradation of fragments produced by endonucleolytic digestion of mRNAs.10,33,56 The mapping of mRNA-poly(A) junctions at multiple locations in several transcripts reinforces the notion that poly(A) polymerase I can polyadenylate and destabilize many different RNA fragments with different 3' extremities thought to be generated by endo and exoribonucleolytic cleavages.4,10,18,20

We mentioned above that precursors of stable RNAs, which accumulate in strains deficient for 3' trimming exoribonucleases, are also polyadenylated.17 It has been proposed that E. coli has developed a quality control system that eliminates abnormally folded RNAs which cannot be rapidly processed into active mature molecules. Consistent with this idea, there is experimental evidence that the precursor of a thermosensitive mutant tRNATrp is polyadenylated and degraded by PNPase (M. Deutscher, personnal communication).

In contrast to eukaryotic cells, where it has been firmy established that poly(A) tails play an important role in translation initiation, there are only very few data supporting the idea that polyadenylation may affect translation in prokaryotes. Based on the observation that ribosomal protein S1 can bind both poly(A) tails and ribosome-binding sites of messengers it has been proposed that S1 could establish a link between the 3' and the 5' ends of the molecule and possibly create a functional interaction between translation and mRNA stability.80 In this respect, ribosomal protein S1 would be the prokaryotic equivalent of the translation initiation factor/poly(A)-binding protein complex of eukaryotic cells. Moreover, if not fortuitous, the association of PAP I82 and of polyadenylated RNAs83 with polysomes could be an indication that poly(A)-dependent degradation affects the stability of mRNAs associated with ribosomes. Polyadenylation, however, does not affect protein synthesis in vitro.84

Polyadenylation also allows the rescue of mutant MS2 bacteriophages whose genome is degraded by RNase III. In this case, addition of A residues by PAP I at the 3' end of fragmented RNA genomes (or of the fragmented complementary strands used as template during replication) allows selection of RNase III resistent genomes containing insertions of stretches of As or Us.15

Although there is no indication that polyadenylation destabilizes fragments of MS2 RNA, most physiological functions that are poly(A)-dependent are based on the destabilization of small RNAs (Table 1). The stabilization of specific transcripts encoding decay enzymes, RNase E and PNPase, by polyadenylation18 may be an indirect effect due to the destabilization of another RNA (Table 1). On the other hand, the fact that five As at the 3' end of RNAs are able to promote exonucleolytic degradation66 suggests that long tails of 50 nucleotides or so may have a different function in mRNA metabolism (see below).

The Role of Polyadenylation in mRNA Decay

The first indication that poly(A) tails destabilize RNA came from the discovery that PAP I controls the stability of the small RNA (RNA I) which regulates the replication of ColE1 plasmids (see above).14 Further study led to the conclusion that PNPase does not bind RNA I, whose 3' end is sequestered in a secondary structure and that poly(A) tails provide sites where PNPase can bind and initiate the exonucleolytic degradation of RNA I.68,69 Polyadenylation has since been shown to be involved in the degradation of other RNA species and of RNA in general9 and it is admitted that the mechanism of degradation of RNA I can be extended to the poly(A)-dependent degradation of any RNA with a 3' secondary structure.9,10,19,21,47,56 The current idea is that PNPase can carry out the complete processive degradation of RNAs containing weak secondary structures but is blocked when it encounters stable hairpins which cause dissociation of the ribonuclease from its substrate.22,56,68 It has been proposed that an Exonucleolytic Impeding Factor, referred to as EIF, might provoke PNPase stalling at secondary structures.85 When blocked at secondary structures, PNPase releases RNAs devoid of a 3' single-stranded stretch of nucleotides that cannot be bound by exoribonucleases ( Fig. 2A). The current model of RNA decay postulates that these tail-less RNAs are readenylated thus allowing PNPase to reinitiate exonucleolytic decay. Again, PNPase can generate tail-less RNA or, possibly, continue to degrade the RNA upstream of the tail and remove few nucleotides at the bottom of the hairpin before to dissociate from the RNA (Fig. 2A).22 ,56 Localized melting of tightly folded secondary structures probably allows PNPase to invade and nibble the base of the structure. Several cycles of exoribonucleolytic degradation and polyadenylation would allow PNPase to progressively shorten the stem and eventually to completely degrade it. Consistent with this hypothesis, it has been shown in vitro that complete degradation of RNAs polyadenylated downstream of a stable hairpin can only be carried out by PNPase if the RNA is continuously adenylated by PAP I.56 It has been proposed that the most stable of 3' protecting hairpins which cannot be traversed by free PNPase are degraded by PNPase associated with degradosome whose activity is facilitated by the RNA helicase RhlB.72 This helicase, activated through its association with the Rne polypeptide is thought to break the hydrogen bonds between nucleotides of the hairpin, which are then removed by PNPase in the degradosome.67 An incremental mechanism of degradation similar to that described above for free PNPase probably accounts for the degradation of stably folded RNAs by degradosome.56 The clustering of the poly(A) sites in several transcripts observed in vivo is consistent with the idea that successive adenylation events are required for progression of PNPase.10,18,20 Because polyadenylation also causes destabilization of rpsO mRNAs in strains lacking PNPase,47 it has been proposed that other ribonucleases might be involved in poly(A)-dependent degradation of RNA.22 One could for example imagine that long tails interfering with internal sequences may induce transconformation of the RNA and expose structural motifs sensitive to endoribonucleases.

Figure 2. Poly(A)-dependent degradation of RNA harboring 3' stable secondary structures.

Figure 2

Poly(A)-dependent degradation of RNA harboring 3' stable secondary structures. A) Poly(A) tails synthesized at the 3' ends of RNAs offer a toehold where exonucleases can initiate decay. RNase II removes the poly(A) tails before to be arrested by 3' hairpins. (more...)

The fact that RNase II can completely degrade the poly(A) tails used as a toehold by PNPase to initiate RNA decay implies that this enzyme antagonizes the degradation of polyadenylated RNAs by PNPase (Fig. 2A).8,22,56 Consistent with this idea, RNase II protects the polyadenylated rpsT transcript from PNPase mediated degradation in vitro.56 This protective role of RNase II explains why the rpsO transcript and the RNA OUT, which represses Tn10 transposition, are more rapidly degraded when RNase II is inactive.8,46,59

It is striking that the full length primary transcripts which have been examined thus far are not, or are only slightly, dependent on poly(A)-dependent degradation in vivo.9,47,56 Indeed, PAP I inactivation only causes appreciable stabilization of full length mRNAs in cells that are also deficient for RNase E and exoribonucleases.9,47 Moreover, poly(A)-dependent ribonucleases cannot participate in the degradation of long polycistronic transcripts which begin to be degraded whereas RNA polymerase has not synthesised yet the 3' end of the molecules.86 This argues against the idea that polyadenylation could initiate decay of primary transcripts.9 We propose that primary transcripts harbor preferential cleavage sites which allow RNase E to control the decay of the molecule (Fig. 2B). Degradation initiated by RNase E could mask the destabilizing effect of polyadenylation if the endonucleolytic pathway degrade mRNA more rapidly than the poly(A)-dependent process catalyzed by exoribonucleases.47 Moreover, elimination of poly(A) tails by RNase II could protect the primary transcript from the exonucleolytic activity of PNPase either free or in the degradosome.56 On the other hand, it is also possible that a degradosome bound simultaneously at the rate limiting processing site, recognized by RNase E, and at the 3' end of the polyadenylated RNA, through PNPase and RhlB, could impair poly(A)-dependent degradation of full length transcripts (Fig. 2B).75 Similar multi-site interactions could coordinate the RNase E cleavages which initiate the decay of RNA I and of the copA and sok RNAs and the subsequent poly(A)-dependent degradation of the processed RNAs.14,19,87 Indeed, it has been shown that the copA RNA must be cleaved by RNase E before becoming a target of poly(A)-dependent degradation21 and it was proposed that RNase E and PNPase interact functionally during degradation of RNA I.68 Initial cleavages of primary transcripts at preferential RNase E sites generate RNA fragments harboring 5' monophosphorylated extremities which promote RNase E processing73 and polyadenylation32 (Fig. 2C). RNase E probably cleaves these RNA fragments at subsequent sites unmasked by the refolding of the RNA fragment or by the run off of ribosomes. In contrast to full length transcripts, RNA fragments can be simultaneously degraded by RNase E and poly(A)-dependent ribonucleases.10,33 One can imagine that both pathways of decay are active because RNase E interacting simultaneously with the 5' monophosphorylated extremity and an internal processing site should not prevent the poly(A)-dependent exonucleolytic degradation, which takes place at the other extremity of the molecule (Fig. 2C). Cleavages by endonucleolytic enzymes will eventually generate RNA fragments devoid of RNase E sites that are exclusively degraded by poly(A)-dependent ribonucleases. This is probably the case for small folded RNAs or mRNA fragments that harbor single-stranded segments too short to contain A-U tracks recognized by RNase E.22,33,56 Although polyadenylation is only required for degradation of tightly folded fragments it seems that it also facilitates decay of unfolded and weakly folded RNAs that can be degraded by RNase II and PNPase even if they are not adenylated.31,33,60 Cooperation of RNase E or degradosome and poly(A)-dependent ribonucleases probably ensures the rapid degradation of RNA into ribonucleotides available for synthesis of macromolecules.

Conclusions and Perspectives

An important recent improvement of our knowledge of RNA metabolism is the discovery that polyadenylation is a ubiquitous mechanism which takes place not only in nucleus and cytoplasm of eukaryotic cells but also in eubacteria and archaea and in chloroplasts88 and mitochondria89 considered as endosymbiotic organelles of prokaryotic origin. Although polyadenylation machineries of eukaryotic cells and bacteria probably derived from a common ancestor,47 it is striking that both the mechanism and the role of polyadenylation are different in prokaryotes and eukaryotes. In contrast to eukaryotic cells, where the vast majority of mRNAs harbors long poly(A) tails that are covered and protected by poly(A)-binding proteins, most RNAs are not adenylated and harbor short poly(A) tails that are accessible to exoribonucleases in bacteria. An intriguing property of bacterial polyadenylation is its lack of specifity. Indeed, current data suggest that any accessible 3' extremity of any RNA species (tRNA, mRNA, rRNA, regulatory RNAs) can be polyadenylated by PAP I. Since poly(A) tails promote RNA degradation, this implies that poly(A)-dependent decay is a global mechanism which can carry out degradation of all RNAs which exhibit an exposed 3' end. Consistent with this idea, polyadenylation promotes degradation of mRNA fragments, regulatory RNAs and nonfunctional tRNAs. Stable RNAs such as functional tRNAs, rRNAs and full length mRNAs whose stability is controled by RNase E are presumably protected from poly(A)-dependent degradation by aminoacylation, ribosomal proteins, RNA binding proteins and RNA folding. On the other hand, phages and plasmids use this mechanism of degradation of cellular RNAs to control the intracellular concentration of regulatory RNAs. Interestingly, in organelles as in prokaryotes polyadenylation promotes mRNA degradation.88,89 Poly(A) tails of up to 270 nucleotides in length containing Gs and few Us and Cs have been characterized in chloroplasts.88 As in bacteria, mRNA, rRNAs and tRNAs are polyadenylated in mitochondria.90 The function of polyadenylation in bacteria and organelles is therefore very different from that of eukaryotic poly(A) tails which stabilize mRNAs. It is worth pointing out, however, that removal of poly(A) tails is the first step of RNA decay pathways initiated at the 3' end of the molecules in both prokaryotes and eukaryotes.

Bacterial polyadenylation and its function in RNA metabolism have only recently begun to be investigated and despite of the fact that several players in poly(A) metabolism have already been discovered, our current view of this mechanism is probably still very naïve. If one takes eukaryotic polyadenylation as a model, it is likely that bacteria contain factors affecting polyadenylation that have yet to be identified. Cofactors and structural features of RNA could affect either the recognition of RNAs by PAP I or that of poly(A) tails by exoribonucleases and thus affect poly(A) synthesis and nibbling. The interaction between PAP I and degradosome components suggests that RNase E mediated RNA decay and poly(A) synthesis could be functionally related. Such interactions might afford a better understanding of why RNA fragments and full length transcripts harboring similar terminal hairpins are not equally sensitive to poly(A)-dependent decay.10,33,56 It is also likely that poly(A)-dependent ribonucleases still have to be identified in E. coli.47 Recent improvements of methodologies of RNA structure determination and the availability of mutants affected in genes coding for enzymes of poly(A) metabolism should allow comparison of the fraction of polyadenylated molecules and the lengths of poly(A) tails of different RNA species. Moreover, utilization of cell-free polyadenylation/RNA degradation assays to reconstitute the polyadenylation machinery and its interactions with enzymes involved in RNA degradation will be required to understand completely this important aspect of RNA metabolism.

Acknowledgements

We are grateful to Ciaran Condon for careful critical reading of the manuscript. The authors are supported by grants from CNRS, the Ministère de l'Education Nationale de la Recherche et de la Technologie and Denis Diderot-Paris 7 University. P.E.M. is recipient of a Ph D grant from Praxis XXI (Portugal).

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