Institute of Cell and Molecular Biology, University of Edinburgh, The Kings Buildings, Mayfield Road, Edinburgh EH9 3JR, UK1
Author for correspondence: Millicent Masters. Tel: +44 131 650 5355. Fax: +44 131 650 8650. e-mail: M.Masters{at}ed.ac.uk
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ABSTRACT |
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Keywords: Escherichia coli, plasmid replication, RNA degradation, ribonuclease III, RNAI
Abbreviations: PAP I, poly(A) polymerase; PNPase, polynucleotide phosphorylase; RIF, rifampicin
a Present address: Inveresk Research International Ltd, Tranent, East Lothian, UK.
b Present address: Asthma Genetics Group, Nuffield Department of Clinical Medicine, University of Oxford, John Radcliffe Hospital, Oxford OX3 9DU, UK.
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INTRODUCTION |
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RNAI breakdown is rapid, and, like that of mRNAs (reviewed by Nierlich & Murakawa, 1996 ; Kushner, 1996
), is accomplished by a combination of endonucleases and exonucleases. For these reasons, it has been studied as a model for mRNA decay. Major roles in RNAI decay have been demonstrated for a number of enzymes. These enzymes are listed in Table 1
. RNase E (Tomcsányi & Apirion, 1985
) is an essential enzyme with endonucleolytic activity important in messenger decay (Kuwano et al., 1977
) and rRNA processing (Ghora & Apirion, 1978
). Lin-Chao & Cohen (1991)
showed that it has a major role in RNAI decay. Polynucleotide phosphorylase (PNPase) is one of the two exonucleases implicated in mRNA degradation (Donovan & Kushner, 1986
). Xu & Cohen (1995)
reported on its role in RNAI decay. Poly(A) polymerase (PAP I), discovered as a consequence of its role in plasmid copy number maintenance (Lopilato et al., 1986
; March et al., 1989
; Liu & Parkinson, 1989
; also reviewed by Sarkar, 1996
, and Cohen, 1995
), has since been implicated in mRNA decay (OHara et al., 1995
; Hajnsdorf et al., 1995
). Its central role in RNAI decay was demonstrated by both Cohens group and our own (Xu et al., 1993
; He et al., 1993
).
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RNase E and PNPase (but not PAP I) have been shown to be associated, with others, in a complex termed the degradosome (Carpousis et al., 1994 ; Py et al., 1994
; Miczak et al., 1996
). We show here that these two enzymes can work separately in RNAI decay. Finally, we propose a pathway for the early stages of RNAI decay based on the half-lives of the primary and processed forms of RNAI in strains lacking one or more decay enzymes.
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METHODS |
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The following plasmids were used: pJF118EH (Furste et al., 1986 ), pBR325 (Bolivar, 1978
), pCML108 (Lin-Chao et al., 1994
), pLH1c (He et al., 1993
) and pBAD-PCN. pBR325 and pJF118EH, closely similar pBR322-based replicons, were used interchangeably to produce RNAI. pCML108 is a pSC101-based replicon from which RNAI, but no other ColE1-derived product, is expressed. In pLH1c, pcnB is cloned downstream of ptac (which is leaky); however, it is also constitutively expressed from an upstream chromosomal promoter which is weaker than the native pcnB promoter; the native promoter is not present in this construct (N. Binns, unpublished). pBAD-PCN is a plasmid in which pcnB expression is under the control of the arabinose-inducible pBAD promoter. Arabinose (0·2 or 0·4%) was added to induce the production or increase the concentration of PAP I during the course of an experiment. Expression from this promoter is almost completely inhibited by growth in the presence of 0·2% glucose. pBAD-PCN was constructed by cloning 1460 bp PCR-amplified pcnB DNA (from pJM513; March et al., 1989
) into the EcoRI and HindIII sites of pBAD18 (Guzman et al., 1995
). The primers used were (upstream) 5'-GCTAT GATTA GCCGG AATTC TTTTG TCCTG-3' and (downstream) 5'-CTGCC TATGG CAAGC TTCGC CACTG TCATG-3'.
RNAI half-life measurements.
Cells were grown at 37 °C in L-broth, treated with rifampicin (RIF; 0·25 mg ml-1; Sigma) to stop further RNA synthesis and sampled at intervals. RNA was extracted, separated by size on polyacrylamide gels, transferred to a nylon membrane and Northern blots were made by hybridizing radioactively labelled oligonucleotide probes, specific for RNAI or for a tRNA control, to the membrane, all as described by He et al. (1993) , except that RIF was added when the OD600 reached 0·4. Strains with temperature-sensitive mutations (rne-1 and rne-3071) were grown at 30 °C to an OD600 of 0·3 and were then transferred to 44 °C and grown for a further 60 min prior to the addition of RIF. tRNA is assumed to be stable and acts as a loading standard. The time-dependent change in RNAI/tRNA was used to calculate RNAI half-life graphically.
Preparation of RNA and Northern blot analysis.
The protocols used for the extraction of RNA and Northern blot analysis were as described by He et al. (1993) with some changes. RNA was separated on 8% polyacrylamide/7 M urea denaturing gels and transferred to positively charged nylon membranes (Boehringer Mannheim) and fixed to the membrane by UV cross-linking, using a Stratagene UV Stratalinker at 1200 µJ, 254 nm. Oligonucleotide probes were end-labelled with 32P using New England Biolabs polynucleotide kinase and following the manufacturers protocol; hybridization was carried out according to Church & Gilbert (1984)
. Hybridization signals were quantified using a Molecular Dynamics Phosphor Imager 400S (Molecular Dynamics) and Image-Quant version 3.22 software. Autoradiographs were made using Kodak X-Omat film. Hybridizations using the UB6 oligonucleotide to detect 3' polyadenylation were done at 40 °C; all others were done at 50 °C. Filters requiring a second hybridization were stripped overnight in 100 ml 50 mM Tris/HCl (pH 8·0), 0·1 mM EDTA, 0·1% SDS at 65 °C.
Oligodeoxyribonucleotides.
Oligonucleotides were purchased from Oswel DNA Service. The probe used to detect RNAI and for primer extension analysis was UB2 (5'-GATCA AGAGC TACCA ACTCT T-3'). The probe, SS2 (He et al., 1993
), was 5'-CCGGT AGAGT TGCCC CTACT CCGGT TTTAG-3'. Primers for the RT-PCR reaction were UB2 and UB1 (5'-ACAGT ATTTG GTATC TGCGC TCTGC-3') for the control reaction and UB6 (5'-TTTTT TTTTA ACAAA AAAAC CACC-3') and UB1 to establish the presence of adenylation.
Primer extension analysis.
Primer extension analysis/reverse transcription was performed on total RNA extracted from cells harbouring the pBR325 plasmid. The method used was from Current Protocols in Molecular Biology (Triezenberg, 1992 ), and used 24 units AMV reverse transcriptase (Promega AMV; 510 units µl-1) and 10 pmol 32P-kinase end-labelled primer (T4 kinase; New England Biolabs), labelled as described by the manufacturer. Unincorporated nucleotide was removed using NAP-5 columns (Pharmacia). One-third of the reaction mix was loaded on an 8% polyacrylamide/7 M urea gel. Primer extension products were sized using a DNA sequencing ladder generated with a Pharmacia T7 Sequencing kit and the supplied M13 template.
RT-PCR.
cDNA was generated from total RNA extracts using the UB6 or UB2 primer and AMV reverse transcriptase at 40 °C using the same methods as described for primer extension except that the primers were not labelled. Twenty-six picomoles of additional primer (UB2 and UB1 for the control reaction or UB6 and UB1 to amplify polyadenylated sequences) was added to 10 µl of the cDNA reaction product along with 200 µmol dNTPs, 1·5 mM MgCl2 and 25 units Taq polymerase (Promega Taq 5 units µl-1) in a final volume of 100 µl Taq buffer. A Hybaid Omnigene thermocycler and the following cycling conditions were used: 1 cycle at 94 °C for 3 min, 50 °C for 30 s and elongation at 72 °C for 1 min followed by 30 cycles of 94 °C for 1 min, 50 °C for 30 s and elongation at 72 °C for 1 min. This was followed by a final elongation step at 72 °C for 10 min. The PCR products were separated on a 1·25% agarose gel, in TBE buffer (0·089 M Tris/borate, 0·002 M EDTA). The amount of ethidium-stained product was measured directly using a Transilluminator and a UVP camera gel analysis system or a Southern blot was analysed using the Phosphor-Imager. The ratio of PCR products generated by the two primer pairs from a single RNA sample was taken as a measure of relative polyadenylation.
In vitro RNase III cleavage.
In vitro cleavage of total RNA samples prepared as described above was performed using His-tagged RNase III and the method described by Li et al. (1993) , with or without potassium glutamate. The purified RNase III, the generous gift of A. Nicholson (Wayne State University, Detroit, MI, USA), had a concentration of 0·14 µg µl-1. The final reaction volume of 20 µl contained 510 µg RNA and 3 µl RNase III. The reaction was incubated at 37 °C for 0, 10, 20 and 30 min and stopped with stop buffer (50% deionized formamide, 20 mM EDTA, 89 mM Tris/HCl, pH 7·5, 89 mM boric acid, 20% sucrose and 0·1% each of bromophenol blue and xylene cyanol). Half of each reaction mix was loaded onto an 8% polyacrylamide/7 M urea gel and transferred to nylon as described above. Hybridization with an RNAI-specific probe was used to reveal the RNase III cleavage pattern of RNAI synthesized from pBR325 or pCML108.
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RESULTS |
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To test whether RNase II has any significant role in RNAI decay we measured the half-lives of RNAI108 in MM38 (Fig. 2d) and RNAI103 in MM38 pcnB rnb (Northern blot not shown). RNAI108 decays at about the same rate in the presence or absence of RNase II (Fig. 2d
; and Table 2
, rows 1 and 5); the rate of RNAI103 decay is also unchanged in the absence of RNase II (Table 2
, rows 3 and 7).
A second, RNase-E-independent, path of RNAI decay involves RNase III
Fig. 3 shows Northern blot analyses of RNAI decay in rne and rnc strains. Fig. 3(b)
(top curve) shows that RNAI108 decays in MM38 rne-3071 with a half-life of ~8 min when the enzyme is inactivated at 44 °C, fourfold slower than in non-mutant cells (see also Lin-Chao & Cohen, 1991
). Although some RNAI108 remains even 60 min after RIF addition (Fig. 3a
), it is clear that RNAI can be degraded independently of RNase E. To test whether this might require RNase III, the second principal endonuclease with a known role in mRNA processing, we constructed an rnc rne-3071 double mutant. Fig. 3(c)
shows that, in MM38 rnc rne-3071, significant amounts of RNAI remain at 60 min after RIF addition, but principally as a novel form of increased length (RNAIex), rather than as RNAI108. The half-life of total RNAI is increased to 33 min in the double mutant. In an rne+ rnc strain RNAIex also accumulates (Fig. 3d
), suggesting that it is not a substrate for RNase E. We conclude that RNase III is required either to remove, or to prevent the formation of, RNAIex.
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Closer examination of the kinetics of RNAI decay in rnc strains shows that the fates of RNAI108 and RNAIex are very different. This is shown in Fig. 3(b) in which the decay of RNAI108+RNAIex (middle curve) is plotted. The complex shape of this curve results from the initial (most likely RNase E mediated) rapid decay of RNAI108 combined with the conversion of a substantial fraction of RNAI108 into the much more stable RNAIex. RNAIex has an approximate half-life of 45 min in this experiment.
RNAIex is likely to be PAP I adenylated
The longer form of RNAI, RNAIex, which appears in rnc strains is likely to be adenylated RNAI108. Observations which suggest this are shown in the Northern blots in Fig. 4(ac). Firstly, extended RNAI108 is not observed in an rnc pcnB strain (Fig. 4a
), indicating that PAP I activity is required for it to be made. Conversely, if PAP I is overexpressed in an rnc strain an increasing proportion of extended RNAI is observed as PAP I concentration increases (Fig. 4b
). In the experiment shown, the production of PAP I was induced by addition of arabinose to MM38 rnc (pBAD-PCN), a strain in which PAP I production is under the control of the araBAD promoter. Overproduction of PAP I also promotes the production of extended RNAI in rnc+ strains; this material is not long-lived (Fig. 4c
).
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Direct Northern blotting of RNAs from different mutant strains was also done using 32P-labelled UB6 and UB2 oligonucleotides as probes. Fig. 4(e) shows that the UB6 probe, designed to detect adenylated RNAI, hybridized preferentially to the RNAI extracted from the rnc mutant or from the PAP I overproducer. The control probe hybridized to all forms of RNAI in each strain. This is the result expected if UB6 is specific for polyadenylated RNAI and if, amongst the strains tested here, RNAI is only extensively polyadenylated in the RNase-III-deficient and PAP-I-overproducing strains. Taking all of the above evidence together, we conclude that RNAIex is likely to be a 3' adenylated form of RNAI108 (to be referred to as RNAI108An) which is produced by PAP I adenylation of RNAI108.
Is RNAI a substrate for RNase III cutting?
The experiments described above demonstrate that a stable, adenylated form of RNAI108 accumulates in the absence of RNase III. This could mean that either RNase III itself cuts RNAI108An, or RNase III works indirectly by, for instance, facilitating the production of the actual degrading enzyme. To distinguish between these possibilities we attempted to identify possible products of RNase III cleavage in vivo and in vitro. Our results, although not conclusive, show that RNase III can cut RNAI, but cannot define an in vivo substrate.
Possible products of RNase III cutting in vivo
Since the degradation products of adenylated RNAI disappear too rapidly to be observed on Northern blots we examined the more stable processed forms of unadenylated RNAI extracted from pcnB strains to see whether their presence is RNase III dependent. Fig. 5(a) shows that two bands of ~93 nt and ~83 nt in size, visible in pcnB extracts, are absent from pcnB rnc extracts, and are thus candidate products of RNase III cutting. We also examined pcnB rne extracts since they lack both major enzymes thought to initiate RNAI decay. The Northern blot in Fig. 5(b)
shows extracts of MM38 rne-1
pcnB sampled after RIF inhibition of RNA synthesis. As RNAI decays, a
95 nt degradation product of RNAI appears. The band is broad and could contain a mixture of the products of RNAI103 and RNAI108 cut at or near nt 98. The kinetics of disappearance of RNAI103 and RNAI108 and of the appearance of the
95 nt form (data not shown) is consistent with this interpretation.
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RNase III can cut RNAI in vitro
If RNase III were able to cut RNAI in vitro, in the absence of any other enzymes of RNA metabolism, to yield products of the same size(s) as the RNase-III-dependent bands identified in vivo this would support the hypothesis that RNase III can cut RNAI in the cell. Protein-free extracts of RNA were prepared from MM38 pcnB (pBAD-PCN) grown with either arabinose (to induce PAP I production) or glucose (to repress production) and digested with purified RNase III using a standard protocol (Li et al., 1993 ). No cutting of RNAI was observed (data not shown). However, when the salt (potassium glutamate), generally considered to be required for specificity of cutting, was omitted from the reaction mixture, cutting was observed. Fig. 6(a)
shows a Northern blot of samples taken during RNase III treatment. All forms of RNAI present are fully and specifically processed to yield two major products each. Since the 103 nt and 108 nt forms are cut to yield products differing in size by 5 bases, the sites of cutting must be near the 3' rather than near the 5' end of RNAI, with the longer product resulting from a cut near nt 98 and the shorter from a cut near nt 82 (indicated in Fig. 1
). When RNAI is folded these bases are close together in a double-stranded region flanking two GC pairs which are in turn flanked by AU pairs. This putative cutting site resembles that identified in rncO (Matsunaga et al., 1996
; Bardwell et al., 1989
). The material in the 73/71 doublet band prominent in the absence of PAP I is also rapidly processed by RNase III.
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RNase-III-dependent cutting of RNAI does not require RNAII
RNase III cuts both double-stranded RNA and folded single-stranded RNA. Since RNAI forms a hybrid with RNAII it is possible that this hybrid, rather than RNAI, is the primary substrate for RNase III. Indeed, Tomizawa & Itoh (1981) reported that this was the case in vitro. As there is less than 10% as much RNAII as RNAI in broth-grown cells, most RNAI will be unhybridized, making it unlikely that only RNAI:RNAII hybrid is being cut. However, to confirm that RNAII is not required for RNase III cutting of RNAI, we repeated several of the experiments described above using pCML108 (Lin-Chao et al., 1994
), a pSC101 derivative which produces RNAI but not RNAII, to supply RNAI. RNAI extracted from pCML108 strains can also be cut by RNase III in vitro to yield fragments of the sizes described above. Extended RNAI also accumulates in an rnc strain with this plasmid (data not shown), showing that hybridization with RNAII is not required to produce material which requires RNase III for its removal.
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DISCUSSION |
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Distinguishing between (2) and (3) depends on determining whether RNAI (or RNAI108An) is a bona fide in vivo substrate of RNase III. We have not been able to do this here. What we have shown is that lack of RNase III leads to the accumulation of a novel form of RNAI (a possible substrate?) and that certain RNAI decay products are not observed when RNase III is absent (possible products?).
An in vitro demonstration that RNase III could cut RNAI to produce the products observed in vivo would support the hypothesis that RNAI is an RNase III substrate in the cell. Our in vitro results show very specific cutting by RNase III, but only at what are normally regarded as low-specificity sites (Li et al., 1993 ; Dunn, 1976
), that is, sites cut only when ionic strength is low. However, since RNase III cleavage of RNAI is not a preferred reaction in vivo, and indeed appears to occur principally on adenylated substrates or in the absence of RNase E or PAP I, inefficient cutting could ensure that RNase III processing does not take precedence over that carried out by RNase E. It is interesting to note that RNase III, which cuts p10Sa RNA in vivo, only cuts this substrate in vitro in low salt in a reaction requiring Mn2+ but inhibited by Mg2+ (Srivastava et al., 1992
). This shows that special conditions can be required for RNase III cutting of particular substrates and that the existence of a cellular factor that promotes RNase III cutting of adenylated RNAI cannot be excluded. Taken together our results are consistent with the possibility, but do not prove, that RNAI is a substrate for RNase III.
A loose consensus sequence has been suggested for RNase III recognition (Krinke & Wulff, 1990 ) but it remains difficult to identify a potential substrate on the basis of sequence alone. RNase III can cut the stems of folded single-stranded RNA molecules; although RNAI has three stemloops these are rather shorter than the stems which have principally been described as RNase III substrates. However, Matsunaga et al. (1996)
have reported that a much reduced stem in rncO, shorter than the RNAI stems, remains a substrate, although a less efficient one, for RNase III cutting.
A suggested pathway for RNAI decay which includes RNase III
Based on the observations reported here and elsewhere we propose that these are the major reactions leading to RNAI decay (Fig. 7).
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(2) RNAI108An is not converted to RNAI103An by RNase E, but is instead broken down by reactions dependent on RNase III. Fig. 1 shows a possible structure for RNAI108An, based on the assumption that some of the 3'As can pair with 5'Us. (It is interesting to note that the longest forms of RNAI we observe, ~117 nt, are equivalent to 1 A per unpaired 5' nucleotide.) RNAI108An is not readily removed by exonucleases; in the absence of RNase III it remains stable. This is particularly surprising in view of the fact that PNPase is increased 10-fold in rnc strains.
(3) RNAI103 is converted to RNAI103An by PAP I. That this reaction can be carried out independently of RNase E cutting was shown by inducing PAP I in a pcnB strain. Accumulated RNAI103 then disappeared rapidly (data not shown).
(4) PNPase promotes RNAI103An decay. In the absence of PNPase, RNAI103An accumulates (Xu & Cohen, 1995 ; and Fig. 1
). That PNPase can work independently of RNase E is shown by the fact that pre-existing RNAI103 is rapidly degraded after PAP I induction.
Thus we suggest that RNAI decay can proceed by one of two principal routes, both of which require that it be adenylated by PAP I. The primary pathway of decay starts with RNase E cleavage, and is followed by PAP-I-mediated adenylation and PNPase-mediated exonucleolytic decay. The secondary route begins with adenylation of RNAI108. Since RNase E does not appear to cleave adenylated RNAI108, RNase III is required. In a pcnB rne rnc mutant, from which RNase E, RNase III and PAP I are absent, RNAI108 none the less continues to disappear. There thus must be other enzymes which are able to initiate its decay.
Implications for mRNA decay
Several of the observations we have made here may have implications for the way in which RNAs other than RNAI turn over. Firstly, each step in RNAI decay seems to require a specific enzyme which does not appear to be replaceable by another enzyme with similar activity. Thus we see that although either RNase II or PNPase is required for cell viability (Donovan & Kushner, 1986 ), suggesting that they can substitute for one another, they do not appear to be able to do so here. Presumably the strong 3' stemloop prevents RNase II action, possibly by denying it an anchor (Cannistraro & Kennell, 1994
), while added A residues offer a handle for PNPase attack (Xu & Cohen, 1995
). Secondly, 3' adenylation is not necessarily sufficient to allow PNPase attack: adenylated RNAI108 appears very resistant. This could in part be because the 5'ppp interferes with PNPase activity, as has been reported for pppRNAI103 (Xu & Cohen, 1995
) and RNase E unprocessable RNAI108 derivatives (Bouvet & Belasco, 1992
), but the degree of resistance suggests another cause. One possibility is that, although this would not be a strong interaction, the 5' A-tract base-pairs with the 3' single-stranded region to sequester both the RNase E cutting site and the single-stranded 3' end (Fig. 1
). Although a mRNA, because it is much longer than RNAI, is likely to be susceptible to several alternative decay routes (Haugel-Nielsen et al., 1996
; Coburn & Mackie, 1996
), any given subsegment may be as restricted as is RNAI in the way in which it can be degraded.
Several associations of enzymes which may expedite RNA decay have been described. The degradosome is a copurifying group of enzymes, which include RNase E and PNPase, with RNA-degrading activity. It has been suggested that the absence of RNase E or PNPase may strongly reduce the rate at which the other can act on its RNAI substrate (Xu & Cohen, 1995 ). We do not find evidence of that here. Although the half-life of RNAI is increased in pnp mutants, the stable material has been processed by RNase E; there is no accumulation of RNAI108, unprocessed by RNase E, in pnp pcnB mutants. Furthermore, digestion of RNAI103An by PNPase can be separated in time from RNaseE action (by inducing PcnB production in a pcnB background). Thus although endo- and exonucleases may well be associated in the cell, effective in vivo activity does not require coordinate action.
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ACKNOWLEDGEMENTS |
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Received 13 April 1999;
revised 15 July 1999;
accepted 21 July 1999.
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