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Reconstitution of the decay of the rpsT mRNA, encoding ribosomal protein S20, with purified enzymes Coburn, Glen Andrew 1998

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R E C O N S T I T U T I O N OF T H E D E C A Y OF T H E rpsT mRNA, E N C O D I N G R I B O S O M A L P R O T E I N S 2 0 , W I T H PURIFIED E N Z Y M E S . by G L E N A N D R E W C O B U R N Hon. B .Sc , The University of Western Ontario, 1992. A THESIS SUBMITTED IN PARTIAL F U L F I L L M E N T OF THE REQUIREMENTS FOR THE D E G R E E OF DOCTOR OF PHILOSOPHY in THE F A C U L T Y OF G R A D U A T E STUDIES Department of Biochemistry and Molecular Biology We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH C O L U M B I A September, 1998 © Glen Andrew Coburn, 1998 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of The University of British Columbia Vancouver, Canada Date <§6PT. 23 , m f t DE-6 (2/88) ABSTRACT Metabolic instability, a hallmark property of all mRNAs, can impart a number of important consequences on the regulation of gene expression. To further study the process of mRNA degradation in Escherichia coli, we have developed an in vitro system to determine the requirements for the complete decay of the rpsTmKNA, encoding ribosomal protein S20. The data show that both purified RNase II and polynucleotide phosphorylase can catalyze the degradation of the 5'-two-thirds of the rpsT mRNA and that prior oligoadenylation of the 3'-termini of truncated rpsT mRNA substrates can stimulate significantly the initiation of degradation by the 3'-exonucleases. The intact rpsT mRNA, however, is insensitive to attack by RNase II or polynucleotide phosphorylase. Furthermore, the single addition of a poly (A) tail to the 3'-end of the rpsT mRNA cannot overcome its resistance to either 3'-exonuclease in vitro. Although previous work has implicated the product of the pcnB gene in the decay of a number of RNAs from Escherichia coli, poly(A) polymerase I does not promote the initiation of the decay of the rpsT mRNA in vivo. It does, however, facilitate the degradation of highly folded degradative intermediates by polynucleotide phosphorylase. As expected, purified degradosomes generate an authentic 147-residue RNase E cleavage product from the rpsT mRNA in vitro. However, degradosomes are unable to degrade the 147-residue fragment in the presence of ATP even when it is oligoadenylated. Rather, both continuous cycles of polyadenylation and polynucleotide phosphorylase activity are necessary and sufficient for the complete decay of the 147-residue fragment in a process which can be antagonized by the action of RNase II. Moreover, both A T P and a non-hydrolyzable analog, ATPyS, support the poly (A) polymerase I and polynucleotide phosphorylase-dependent degradation of the 147-residue intermediate implying that ATPase activity, such as that which may reside in RhlB, a putative R N A helicase, is not necessarily required. Alternatively, the rpsT mRNA can be degraded in vitro by a second 3'-decay pathway which is dependent on poly(A) polymerase I, polynucleotide phosphorylase and ATP alone. Complete degradation of a fragment of the malE-malF mRNA in vitro exhibits additional requirements. Degradation of this R N A is dependent on the degradosome, A T P and poly(A) polymerase I. Unlike the situation for the rpsT mRNA, the non-hydrolyzable analog, ATPyS, cannot substitute for ATP suggesting that ATP hydrolysis is required for decay of structured portions of the malE-malF mRNA. Our results demonstrate that a hierarchy of R N A secondary structures controls access to exonucleolytic attack on 3'-termini. Moreover, decay of a model mRNA can be reconstituted in vitro by a small number of purified components in a process which is more dynamic and ATP-dependent than previously imagined. The implications of these findings are discussed in a number of models of mRNA decay. -Hi-TABLE OF CONTENTS Page Abstract ii List of Figures viii List of Tables x List of Abbreviations xi Acknowledgements xiii Chapter 1-mRNA decay in Escherichia coli 1 1.1 Introduction 1 1.2 The ribonucleases 2 1.2.1 The endonucleases 3 1.2.1.1 RNase E 3 1.2.1.2 RNase III 12 1.2.1.3 RNase P 17 1.2.2 Theexonucleases 19 1.2.2.1 Polynucleotide phosphorylase (PNPase) 19 1.2.2.2 RNase II 25 1.2.2.3 Oligoribonuclease 30 1.2.3 Regulation of ribonuclease expression 31 1.3 Other enzymes of mRNA decay 32 1.3.1 Poly(A) polymerases 32 1.3.2 RNAhelicases 34 1.4 Other protein factors 36 1.5 The role of translation 38 1.6 Models of mRNA decay 39 1.7 Experimental rationale 42 -iv-Chapter 2-Cloning, overexpression and purification of the enzymes of mRNA decay 44 2.1 Introduction 44 2.2 Materials and methods 44 2.2.1 Enzymes and chemicals 44 2.2.2 Bacterial strains and plasmids 45 2.2.3 Molecular biological methods 45 2.2.1.1 Isolation of genomic D N A from Escherichia coli 46 2.2.1.2 Oligonucleotide primers for PCR 46 2.2.1.3 Polymerase chain reaction 48 2.2.1.4 Ligations 48 2.2.1.5 Transformations 49 2.2.1.6 Analysis of clones 49 2.2.1.7 Oligonucleotide primers for D N A sequencing 50 2.2.4 Analysis of proteins 52 2.2.4.1 Quantitation 52 2.2.4.2 SDS-polyacrylamide gel electrophoresis (SDS-PAGE) . . 52 2.2.4.3 Enzyme assays 52 2.2.5 Protein purification 53 2.2.5.1 Media 53 2.2.5.2 Induction 53 2.2.5.3 Harvest and lysis 54 2.2.5.4 Crude fractionation of the RnbCA-38 protein 54 2.2.5.5 Purification of the RnbCA-10 protein from strain G O O . 54 2.2.5.6 Purification of recombinant RNase II from strain GC100 56 2.2.5.7 Overexpression and purification of PNPase from strain GC400 57 2.2.5.8 Purification of degradosomes from strain CF881 58 2.2.5.9 Purification of the recombinant RhlB protein from strain GC300 60 2.3 Results 62 2.3.1 Construction of a strain overexpressing the rnb gene 62 2.3.2 Purification of recombinant RNase II 65 2.3.3 Purification of C-terminally truncated derivatives of RNase II . . 6 9 2.3.3.1 TheRnbCA-10 protein 69 2.3.3.2 TheRnbCA-38 protein 70 2.3.4 Construction of a strain overexpressing the pnp gene 72 2.3.5 Purification of recombinant PNPase 72 2.3.6 Construction of a strain overexpressing the rhlB 77 2.3.7 Purification of recombinant RhlB protein 77 2.3.8 Purification of the degradosome 80 -v-2.3.9 Sensitivity of 9S R N A to degradosomes 82 2.3.10 RNA-binding properties of the degradosome 84 2.4 Discussion 86 er 3-The action of the 3-exonucleases against a synthetic R N A substrate 91 3.1 Introduction 91 3.2 Materials and methods 92 3.2.1 Enzymes and chemicals 92 3.2.2 Bacterial strains and plasmids 93 3.2.3 Preparation of crude extracts 93 3.2.4 In vitro synthesis of R N A transcripts 94 3.2.5 Assay for RNase II activity 95 3.2.6 Assays for PNPase activity 96 3.2.6.1 Method A 96 3.2.6.2 Method B 96 3.2.7 U V photochemical crosslinking 97 3.3 Results 97 3.3.1 Exonucleolytic activity in crude extracts from E. coli 97 3.3.2 Properties of recombinant RNase II 101 3.3.2.1 Activity against t40B 101 3.3.2.2 Size of products 103 3.3.2.3 Thermolability of RNase II 105 3.3.2.4 Binding of the Rnb protein to substrate 108 3.3.3 The activity of the RnbCA-38 and RnbCA-10 proteins against thet40B substrate 110 3.3.4 Properties of recombinant PNPase 112 3.4 Discussion 119 3.4.1 The mechanism of action of RNase II on a novel substrate . . . . 119 3.4.2 The deletions RnbCA-38 and RnbCA-10 lie within the putative SI RNA-binding domain of RNase II 122 3.4.3 The mechanism of action of PNPase 123 3.4.4 A model for control of mRNA degradation at the 3'-end 125 3.4.5 Utility of t40B as a substrate for 3'-exonucleolytic decay 126 er 4-The action of the degradosome and the 3'-exonucleases on natural R N A substrates 128 4.1 Introduction 128 4.2 Materials and methods 129 4.2.1 Enzymes and chemicals 129 4.2.2 Bacterial strains and plasmids 130 -vi-4.2.3 Preparation of R N A substrates 130 4.2.3.1 Transcription 130 4.2.3.2 Polyadenylation 131 4.2.4 Ribonuclease assays 132 4.2.5 R N A extraction and analysis 133 4.3 Results 134 4.3.1 Sensitivity of the rpsTmRN As to 3 '-exonucleolytic digestion .134 4.3.2 Polyadenylation stimulates 3'-exonucleolytic digestion 146 4.3.3 Northern analysis of the rpsT mRNAs 153 4.3.4 Sensitivity of the rpsT/365 R N A and derived products to degradosomes 155 4.3.5 Continuous polyadenylation and 3'-exonucleolytic decay 159 4.3.6 Continuous polyadenylation and degradation of the malE-malF REP sequence 169 4.4 Discussion 177 4.4.1 Reconstitution of mRNA degradation with purified components 177 4.4.2 Is poly(A) polymerase I required to initiate mRNA decay ? . . . 181 4.4.3 Hierarchies of R N A secondary structure, poly(A) polymerase I, and 3'-exonucleolytic decay 182 4.4.4 malE REP decay revisited 187 Chapter 5-Conclusions and perspectives 189 5.1 Summary 189 5.2 Models for investigating mRNA decay 189 5.2.1 The 5'-tethering model 190 5.2.2 The 3'-latent decay model 200 5.2.3 The 3'-tethering model 204 5.2.4 5'-decay vs. 3-decayby the degradosome 206 5.3 Paradigms for the selective decay of mRNA 210 5.3.1 RNA1 210 5.3.2 The rpsT mRNAs 212 5.3.3 The rpsO mRNA 213 5.4 Perspectives 216 References 219 -vii-L I S T O F F I G U R E S F I G U R E S Figure Description Page 1.1 Schematic representation of the Rne/Ams protein 8 1.2 Role of R N A secondary structure on endonucleolytic decay by RNase E 11 1.3 RNase III consensus processing signals 14 1.4 Possible recognition elements for RNase P projected onto t R N A p h e 18 1.5 Processive versus non-processive R N A degradation by PNPase 22 1.6 Localization of RNA-binding domains in three ribonucleases from E. coli 24 1.7 The effect of R N A secondary structure on 3-exonucleolytic degradation 27 1.8 RNase II as a repressor of mRNA decay 28 1.9 Conventional models of mRNA degradation 40 1.10 Model for the structure of the rpsTmKNA, encoding ribosomal protein S20 43 2.1 Strategy for cloning and overexpression of rnb 64 2.2 Purity of recombinant RNase II 67 2.3 Overexpression of the recombinant RnbCA-38 protein and purity of the RnbCA-10 protein 71 2.4 Strategy for cloning and overexpression ofpnp 73 2.5 Purity of recombinant PNPase 74 2.6 Strategy for cloning and overexpression oirhlB 78 2.7 Purity of recombinant RhlB 79 2.8 Purity of the degradosome from strain CF881 81 2.9 The action of the degradosome against 9S R N A 83 2.10 The RNA-binding properties of the degradosome and RhlB 85 3.1 3-exonucleolytic degradation of a partial duplex R N A substrate 98 3.2 Degradation of the partially duplexed R N A by PNPase activity in crude extracts . . . . 100 3.3 Degradation of the partial duplex R N A substrate by purified recombinant RNase II . . 102 3.4 Competition between partially digested t40B and complete t40B 104 3.5 Analysis of the products generated by RNase II digestion of t40B 107 3.6 U V crosslinking of t40B to purified recombinant Rnb protein and proteins in the S-150 fraction prepared from strain CF881 109 3.7 Action of the RnbCA-10 protein against t40B R N A I l l 3.8 Degradation of the partially duplexed t40B R N A substrate by purified recombinant PNPase 113 3.9 The action of PNPase in a low ionic strength buffer 115 3.10 The action of PNPase in a moderate ionic strength buffer 117 3.11 Competition between rpsT/365 R N A and #ps77209-poly(A) R N A 118 3.12 The mechanism of action RNase II on t40B R N A 121 -vi i i -Figure Description Page 4.1 Model for the structure of the rpsT mRNA 135 4.2 The rpsT R N A substrates 137 4.3 Digestion of the /psJmRNA by exonucleases 138 4.4 Digestion of the rpsT/3 3 3 Aterm R N A by exonucleases 141 4.5 Digestion of rpsT/221 R N A by exonucleases 142 4.6 Mapping of intermediates generated by RNase II digestion 144 4.7 Degradation intermediates formed by exonucleolytic digestion of the rpsTKNAs . . . . 145 4.8 Digestion of ?ps77395-poly(A) R N A by exonucleases 147 4.9 Digestion of r/?s773 63Aterm-poly(A) by exonucleases 149 4.10 Digestion of rpsT/251 -poly(A) by exonucleases 151 4.11 Competition between rpsTIll 1RNA and rpsT/251 -poly(A) R N A for exonucleases . . . 152 4.12 Effect ofpcnB on the steady-state levels of the rpsT mRNAs 154 4.13 Digestion of the rpsT mRNA by degradosomes 156 4.14 Effect of a single round of polyadenylation on decay of the 3'-end of the rpsT mRNA 158 4.15 Effect of continuous polyadenylation on decay of the 3'-end of the rpsT mRNA 160 4.16 Effect of continuous polyadenylation on decay of the rpsT mRNA 162 4.17 Effect of continuous polyadenylation on 3'-decay of the rpsTmRNA 164 4.18 Effect of RNase II on the decay of the 3'-end of the rpsTmRNA 166 4.19 Effect of continuous polyadenylation on decay of the rpsT mRNA in the presence of RNase II 168 4.20 The REP stem-loop structure 169 4.21 Digestion of the malE REP R N A by the degradosome 172 4.22 Digestion of the malE REP R N A by PNPase 174 4.23 Effect of continuous polyadenylation on the decay of the malE REP R N A by the degradosome 176 4.24 Degradation of the rpsT mRNA 179 4.25 Alternative mechanisms of poly(A)-mediated decay of highly folded RNAs 185 5.1 The 5'-tethering model 192 5.2 Effect of R N A secondary structure on mRNA decay 194 5.3 Effect of translation on mRNA decay 196 5.4 Interrupted 5'-tethering model 198 5.5 The 3'-latent decay model 201 5.6 The 3'-tethering model 205 5.7 RNase E control of mRNA decay 208 5.8 Summary of the decay of RNA1 211 5.9 Summary of the decay of the rpsO mRNA 215 ix-LIST OF TABLES Table Description Page Number 2.1 Oligonucleotide primers for amplification of the rnb gene by the polymerase chain reaction (PCR) 47 2.2 Oligonucleotide primers for amplification of the pnp gene by PCR 47 2.3 Oligonucleotide primers for amplification of the rhlB gene by PCR 47 2.4 Oligonucleotide primers for D N A sequencing of the rnb gene 51 2.5 Oligonucleotide primers for D N A sequencing of the pnp gene 51 2.6 Purification of recombinant RNase II 68 2.7 Purification of recombinant PNPase 71 3.1 Plasmid-encoded transcripts 95 4.1 Summary of plasmid-encoded rpsT transcripts 131 -x-LIST OF ABBREVIATIONS ATP adenosine 5-triphosphate ATPyS adenosine 5'-0-(3thiotriphosphate) bp base pair B S A bovine serum albumin °C degree Celsius cm centimeter C-terminal carboxyl-terminal CTP cytidine 5'-triphosphate dATP deoxyadenosine 5'-triphosphate D E A D (aspartate-glutamate-alanine-aspartate) DEPC diethyl pyrocarbonate D N A deoxyribonucleic acid DNase deoxyribonuclease dNTP deoxynucleotide triphosphate DTT 1,4-dithiothreitol E D T A ethylenediamine tetraacetate FPLC fast protein liquid chromatography g gravity gDNA genomic D N A GTP guanosine 5'-triphosphate h hour HEPES N-2-hydroxy ethyl piperazine-N'-2-ethanesulfonic acid His histidine IPTG isopropyl-P-D-thiogalactopyranoside kbp kilobase pair kDa kiloDalton L B Luria-Bertani broth M molar mg milligram uCi microCurie Hg microgram ul microliter min minutes ml millilter mM milliMolar mRNA messenger ribonucleic acid M W molecular weight NDP nucleoside diphosphate ng nanogram N M P nucleoside monophosphate N M R nuclear magnetic resonance NP-40 nonidet P-40 nt nucleotide N-terminal amino-terminal P A G E polyacrylamide gel electrophoresis PAP poly(A) polymerase P/C/I phenol/chloroform/isoamyl alcohol PCR polymerase chain reaction pmol picomole PMSF phenylmethylsulfonylfluoride Pnp the pnp gene product PNPase polynucleotide phosphorylase poly(A) polyadenylate (polyadenylic acid) R B D R N A binding domain REP repetitive extragenic pallindrome RhlB the rhlB gene product RNase ribonuclease Rnb the rnb gene product Rne/Ams the rne/ams gene product rRNA ribosomal ribonucleic acid S30 supernatant of 30,000 x g centrifugation S150 supernatant of 150,000 x g centrifugation S200 supernatant of 200,000 x g centrifugation SD Shine-Dalgarno sequence SDS sodium dodecyl sulfate SSC standard saline citrate T A E Tris-acetate-Na E D T A TB Terrific Broth T B E Tris-borate-Na E D T A Tris tris(hydroxymethyl) aminomethane tRNA transfer R N A U units UTP uridine 5-triphosphate U V ultraviolet V volt v/v volume/volume w/v weight/volume -xii-ACKNOWLEDGEMENTS Six years, 4,500 cups of coffee, 2,000 miles and two stolen computers = one thesis. A very special thanks to Dr. George Mackie for his outstanding guidance and supervision. None of this work would have been possible without his patience, hard work and dedication. Thanks for everything George. You are a true mentor. I would also like to thank the many members of the Mackie lab for all of their support and friendship throughout the years: Robbie Cormack, Michele Rouleau, Julie Genereaux, Doug Briant, Kristian Baker, Annie Prud'homme Genereux, Stephanie Masterman (the Rose), Catherine Spickler and the many fourth year and summer students. A special thanks goes to Xin Miao (X-man), Ken Niguma (Kenny G) and Anand Rampersaud (Mookie Rotorsaud). I can't think of three better (clowns) guys to move across the country with. Thanks go out to all my friends (overeducated bums) who have made my stay in Vancouver so enjoyable: John Rohde, John Brunstein, Angus McQuibban, Chris Loewen, Manish Joshi, Cameron Mackereth, Deb Sauve, Neena Kuriakose and especially to Pat and Renee Gowdy. A very special thanks to my longtime friends Trevor Birmingham, Jennie Landry and Martin Milne. You guys were never far away from my thoughts during this production. Finally, I would like to thank my family and my beautiful wife Kimberley for their constant love and support throughout this endeavor. This would not have been possible without your patience and understanding. Thank you so much. "Yes mom. The proteins are alright! " - x i i i -C H A P T E R 1 m R N A decay in Escherichia coli 1.1 Introduction The level of expression of a gene is determined by the rates of transcription, translation and decay of its messenger RNA. While much is known about the former two processes, equally detailed information about the factors controlling mRNA decay is lacking. In the bacterium Escherichia coli, each mRNA exhibits a unique half-life which can vary from less than 40 seconds to more than 20 minutes (Belasco and Higgins, 1988; Higgins et al, 1992). In higher eukaryotes, the half-lives of mRNAs vary over several orders of magnitude ranging from 20 minutes to over 24 hours (reviewed in Ross, 1995), while in yeast the most unstable mRNAs decay in 5 minutes while the most stable mRNAs decay with rates greater than 60 minutes (reviewed in Caponigro and Parker, 1996). Although much progress has been made in understanding mRNA decay in eukaryotes, the preferred system for studying mRNA degradation is still E. coli. The inherent instability of prokaryotic mRNA exerts a number of important consequences on the cell. First, mRNA stability affects not only the kinetics of induction but overall steady-state levels of gene expression (reviewed in Nierlich and Murakawa, 1996). Put differently, the more stable of two mRNAs expressed from the same promoter will accumulate faster and maintain a higher concentration than its unstable counterpart. Second, along with translational efficiency, mRNA stability contributes to differential regulation of polycistronic mRNAs (Belasco et al, 1985; Newbury et al, 1987). Third, mRNA decay serves to amplify the effects of repressing gene expression (Singer and Nomura, 1985; Cole and Nomura, 1986). Fourth, degradation of mRNA serves to maintain the pool of ribonucleotides lowering the energy demands on the cell (Danchin, 1996). Finally, R N A stability plays an important role in antisense control both natural and therapeutic (Inouye, 1988). 1.2 The ribonucleases Of the twenty ribonuclease activities identified in Escherichia coli (reviewed in Deutscher, 1993), only RNase E, RNase III, polynucleotide phosphorylase (PNPase) and RNase II are known to directly contribute to mRNA decay. The majority of the other enzymes are mainly involved in tRNA maturation. RNase P, another endonuclease, is involved in the 5'-maturation of pre-tRNA while several exonucleases including RNase T, RNase PH, RNase B N , and RNase D are involved in maturing the 3'-end of pre-tRNAs (Deutscher, 1993) and several small stable RNAs including M l R N A and the lOSa R N A (Li et al, 1998). The finding that RNase T is required for the maturation of pre-5S rRNA (Li and Deutscher, 1995) and the finding that an RNase P cleavage confers stability to the his operon mRNA from Salmonella typhimurium (Alifano et al, 1994) has alerted us to the possibility that known activities not previously associated with mRNA decay may play additional functions in R N A metabolism. Except for RNase I (Neu and Heppel, 1964), a periplasmic ribonuclease, most other activities are poorly characterized and/or appear to be proteolytic fragments of "known" ribonucleases (reviewed in Coburn and Mackie, 1998). The following introduction will review the major players involved in mRNA decay and will attempt to give an overview of the structure, function and mechanism of action of each enzyme. The role of R N A secondary structure in directing the action of each ribonuclease will also be examined. 1.2.1 The endonucleases 1.2.1.1 RNase E RNase E was originally discovered as an activity required for processing of 9S R N A to the immediate precursor to 5S ribosomal R N A (rRNA) (Misra and Apirion, 1979). In addition, it was also found to cleave RNA1 encoded by colEl-type plasmid replicons (Tomcsanyi and Apirion, 1985). A temperature-sensitive mutation, rne-3071, was discovered by Apirion and coworkers which rendered RNase E processing of the 9S precursor thermolabile (Ghora and Apirion, 1978). About the same time, Ono and Kuwano uncovered an apparently independent temperature-sensitive mutation, termed ams-1 (altered j n R N A stability), which stabilized significantly the half-life of bulk mRNA (Ono and Kuwano, 1979). Some years later, a number of laboratories studying mRNA decay simultaneously reported that ams was allelic to me (Mudd et al., 1990; Babitzke and Kushner, 1991; Taraseviciene etal., 1991; Melefors and von Gabain, 1991). The observation that partially degraded mRNA fragments accumulate in rne/ams mutants strongly suggested that RNase E plays a central role in the decay of most, if not all, mRNAs in E. coli. In the following discussion it should be noted that RNase E refers to the endonucleolytic activity while Rne/Ams refers directly to the product of the rne/ams gene. The reason for this distinction lies in a number of misconceptions pertaining to the size and number of subunits which compose the activity known as RNase E. Further efforts to characterize the Rne/Ams protein and its gene have been severely hampered by two major complications. First, RNase E is extremely sensitive to proteolysis during purification (Roy and Apirion, 1983). Furthermore, many proteolytic fragments derived from the Rne/Ams protein maintain at least partial enzymatic activity. In one case, a proteolytic fragment of RNase E was incorrectly described as a "new" activity (RNase K) required for specific cleavage of the ompA mRNA (Lundberg etal, 1990; Lundberg etal, 1995). A second complication emanated from initial reports which described the cloning and sequencing the rne gene. Of four published reports claiming to identify the rne or ams genes by complementation, not one obtained the complete gene (Ray and Apirion, 1980; Chandaetal, 1985;Dallmannetal, 1987; Claverie-Martin etal, 1989) and only two contained a portion (Ray and Apirion, 1980; Claverie-Martin etal, 1989). Partial D N A sequence was obtained from two clones, however, they contained several small errors (Chauhan et al, 1991; Claverie-Martin etal, 1991). Later, the "full-length" gene was cloned fortuitously as a myosin-like protein capable of cross-reacting with yeast anti-myosin heavy chain monoclonal antibodies (Casaregola et al, 1992). A few minor errors in the D N A sequence, which resulted in an underestimate of the size of the open reading frame, were finally corrected by George Mackie to establish the correct size of the Rne/Ams protein at 1061 amino acids (Cormack et al, 1993, GenBank accession #L23942; Casaregola etal, 1994). In addition to proteolysis, RNase E tends to aggregate upon overexpression and generally behaves poorly during purification. Our laboratory has overcome a number of these problems by renaturing RNase E activity from SDS-polyacrylamide gels (Cormack et al, 1993). The gel-purified and renatured protein retains authentic RNase E activity demonstrating, unequivocally, that the 1061 amino acid Rne/Ams protein, encoded by the rne gene, is the only component in RNase E required for its endonucleolytic activity (Cormack et al, 1993). Although this purification method can generate highly purified RNase E for some functional studies, only a fraction of the renatured Rne/Ams protein is fully active. To date, methods for generating homogeneous preparations of RNase E with high yields and activity are still lacking. Improvements in the preparation of RNase E, namely the inclusion of several protease inhibitors and rapid purification procedures at 4 °C, have led to a potentially important finding; RNase E copurifies with a number of proteins as part of a high molecular weight complex referred to as the degradosome (Carpousis et al, 1994). These results were confirmed by immunoprecipitation experiments and by the independent identification of a complex which binds to the REP (repetitive extragenic palindrome) stem-loop of the malE-malF mRNA (Carpousis et al., 1994; Py et al., 1994). The degradosome consists of apparently stoichiometric amounts of the following proteins: Rne/Ams, the major endonuclease \nE. coli; PNPase, a 3'-exonuclease involved in the terminal stages of mRNA decay; RhlB, a putative DEAD-box containing R N A helicase and enolase, a glycolytic enzyme of unknown significance in mRNA decay (Carpousis et al., 1994; Py et al., 1994; Py et al., 1996; Miczak et al., 1996). At present, it is unknown whether or not RNase E, PNPase, or RhlB cycles between free and complexed forms within the cell. However, only 5-10% of the cellular enolase appears to copurify with the degradosome (Py et al., 1996). Sub-stoichiometric quantities of DnaK and GroEL were also found to copurify with the complex in some immunoprecipitation experiments with FLAG-epitope tagged Rne/Ams (Miczak et al., 1996). However, the function of DnaK and GroEL in the degradosome remains unclear since degradosomes which lack these proteins are fully functional in RNase E assays in vitro. Furthermore, there is no evidence that the heat shock response, which induces DnaK and GroEL, is involved in mRNA decay (Henry et al., 1992). For these reasons it is likely that these rather abundant proteins are spurious contaminants. An additional minor component of the degradosome was found to be polyphosphate kinase (Blum etal, 1997), an enzyme ubiquitous to prokaryotes which reversibly catalyzes the synthesis of polyphosphates from the y-phosphate of ATP (reviewed in Kornberg, 1995). Although the function -5-of this protein in mRNA decay remains unclear, Blum et al. have suggested that it may provide the correct "microenvironment" for the degradosome by removing ADP and replacing ATP (Blum et al, 1997). The presence of at least two enzymes of mRNA decay in a high molecular weight complex is provocative and may explain the apparent ("all or none") concerted nature of mRNA degradation. Further work, however, is required to confirm the presence and function of the minor components in the degradosome and to determine the physiological significance, if any, of the degradosome complex itself. Since RNase E homologs are inferred to exist in a number of prokaryotic species by immunological analyses (Taraseviciene et al, 1994), it will be interesting to see if the organization of the enzymes of mRNA decay into "degradosomes" is conserved amongst all bacterial species. Degradosomes have also been isolated from the photosynthetic bacterium Rhodobacter capsulatus. The complex contains three major proteins with the following molecular masses: 60 kDa, 50 kDa and 48 kDa. N-terminal protein sequencing confirmed the identity of the latter two proteins as an R N A helicase-like protein and enolase respectively. Surprisingly, the complex did not contain a homologue toE. coli PNPase. The chloroplast of higher plants also utilizes a high molecular weight complex of proteins for mRNA 3'-end processing and decay (Hayes et al, 1996). The complex contains, among others, a protein of 67 kDa which cross-reacts with anti-RNase E antibodies and a protein called 100 RNP which shares striking homology to E. coli PNPase (43% identical and 63% similar amino acid sequence) (Hayes et al., 1996). Purified p67 exhibits endonucleolytic activity whereas purified 100 RNP was shown to be a phosphorolytic 3-exonuclease in vitro (Hayes et al., 1996). Two reports have predicted that RNase E-like activities exist in mammalian cells by complementation of the temperature-sensitive rne mutation with a portion of a human gene termed -6-ard-1 (Wang and Cohen, 1994) and by immunological analysis and activity as assayed against 9S R N A (Wennborg et al, 1995). One should be cautious, however, when defining homologous activities in eukaryotic cells since even relatively non-specific nucleases can attack single-stranded regions of R N A resulting in a pattern that is indistinguishable from RNase E cleavage. Structure-function investigations by our laboratory and others (Taraseviciene et al., 1995; McDowall and Cohen, 1996) have demonstrated that the Rne/Ams protein can be divided into roughly three domains: the amino-terminal domain; residue 1-500, the central domain; residues 500-850 and the carboxyl-terminal domain; residues 850-1061 (Figure 1) (Taraseviciene et al., 1995; McDowall and Cohen, 1996). The N-terminal domain of the protein is thought to house the catalytic site for RNase E. This region of the protein contains the two temperature-sensitive alleles, ams-1 (G66S) and rne-3071 (L68F), which map to codons 66 and 68, respectively (Figure 1.1) (McDowall et al, 1993). It is unclear whether these mutations directly affect the active site; however, they lie within an SI RNA-binding domain (residues 35-125) (Figure 1.1) (Bycroft et al, 1997). This circumstantial evidence is supported by the finding that truncated versions of the Rne/Ams protein, as small as the first 498 N-terminal amino acids, are enzymatically active (McDowall and Cohen, 1996). The SI RNA-binding fold, named after a repeating motif in its archetypical member, ribosomal protein SI, appears to be an ancient polynucleotide binding motif which is conserved amongst a large group of proteins including RNase E, RNase II and PNPase (Bycroft et al, 1996). Recently, the solution structure of the SI RNA-binding domain from PNPase was solved by N M R spectroscopy and was found to be a five-stranded antiparallel P-sheet/P-barrel with the strands arranged in a "Greek key" topology (Bycroft et al, 1996). The RNA contact residues could not be clearly assigned -7-N-terminal Domain Central Domain C-terminal Tail < ; • > ams-1 rne-3071 608 RRKPRQNNRRDRNER 622 N 1061 ] c 35 125 600 750 S1-RBD R-RBD Catalytic Domain (1-498) •< • RNA-binding Domain PNPase Interaction Domain (608-1061) Figure 1.1 Schematic representation of the Rne/Ams protein. A schematic representation of the linear amino acid sequence of RNase E is shown in this panel. The 1061 amino acid Rne/Ams protein can be divided into three functional domains. The N-terminal domain, the central domain and the C-terminal tail shown above. Two putative RNA-binding domains: SI- RNA-binding domain (Sl-RBD) (Bycroft etal, 1997) and the arginine rich RNA-binding domain (R-RBD) are marked by the black boxes. The two classical temperature sensitive mutations ams-1 and rne-3071 within the Sl -RBD and the putative 15 amino acid R-RBD are denoted above. The proposed function of each domain is shown below. -8-in the SI domain from PNPase; however, by analogy to the single-stranded DNA-binding replication protein A (Bochkarev etal, 1997), the RNA-binding site is likely a channel on the surface of the P-barrel. Thus, the SI domain of RNase E would contact three nucleotide residues in the R N A adjacent to the site of cleavage. This is consistent with the finding that cleavage sites lie within single-stranded regions of the RNA. The central domain of the protein (residues 500-850 approximately) is rich in basic amino acids, containing 45 Arg/Lys residues. A second RNA-binding domain, referred to as the R-RBD, is located within this central portion of the protein and has been mapped to residues 600-750 by deletion analysis of the gene (X. Miao and G. Mackie, unpublished data; McDowall and Cohen, 1996; Taraseviciene etal, 1995) and by partial proteolysis (R. Cormack and G. Mackie, unpublished data) (Figure 1.1). A more refined set of deletions has placed the R-RBD between residues 608-622 (X. Miao and G. Mackie, unpublished). The R-RBD is a functional RNA-binding domain in vitro (Cormack et al, 1993) and resembles known arginine-rich RBDs (> 40% arginine), such as those found in the FflV-1 proteins Tat and Rev (Burd and Dreyfuss, 1994). The role of this RNA-binding domain in vivo remains unsettled. Recent data suggests that the R-RBD may play a role in product retention and/or enzyme processivity (S. Masterman, X . Miao and G. Mackie, unpublished data; Taraseviciene etal, 1995; Kaberdin etal, 1996). Others have suggested that in addition to substrate binding, the R-RBD may also destabilize R N A secondary structure to reveal otherwise cryptic RNase E sites (Kaberdin et al, 1996). The acidic C-terminal tail (residues 850-1061) of the Rne/Ams protein is not required for catalytic function or RNA-binding (Figure 1.1). Two reports , however, suggest that the C-terminal 200 amino acids are essential for interactions with PNPase (Kido et al, 1996; K . Niguma, X . Miao -9-and G. Mackie, unpublished). RNase E activity requires Mg 2 * or similar divalent metal ions. Ca 2 + can substitute for RNA-binding, however, it does not support enzymatic cleavage (Cormack et al., 1993). RNase E is specific for single-stranded R N A and tends to favour A U rich sequences, presumably because these sequences are more likely to be in a single-stranded conformation. Cleavage of single-stranded R N A releases products which contain 5'-monophosphates and 3-hydroxyl termini. The following loose consensus sequence for RNase E cleavage has been proposed: (A/G)l AUU(A/U) (Mackie, 1991; Ehretsmann etal, 1992). It should be noted, however, that not all cleavage sites conform to the consensus and that not all single-stranded A U sequences are cleaved by RNase E. These observations suggest that cleavage site selection is much more complex than the recognition of a specific sequence of nucleotide residues. Site selection may, in fact, involve secondary and possibly higher-order R N A structure. The role of R N A secondary structure in modulating endonucleolytic cleavage by RNase E has also been extensively studied both in vivo and in vitro. R N A stem-loop structures can inhibit cleavages by RNase E in one of two ways: occlusion of potential cleavage sites by base-pairing (Figure 1.2B) or by steric hindrance (Figure 1.2A). Interestingly, "lO-'mer" oligoribonucleotides derived from RNA1 are cleaved by RNase E approximately 20-fold faster than RNA1 (McDowall et al, 1995) and led to the hypothesis that R N A structure can inhibit cleavage by RNase E. In contrast, investigations into the decay of the rpsTmRNA and the processing of 9S R N A demonstrate that removal of some stem-loop structures can actually decrease the rate of cleavage by RNase E at adjacent sites (Cormack and Mackie, 1992; Mackie and Genereaux, 1993). This suggested that R N A structure may play a role in anchoring cleavage sites in a single-stranded conformation (Cormack and -10-D 5' Figure 1.2 Role of RNA secondary structure on endonucleolytic decay by RNase E. Stem-loop structures have been shown to inhibit cleavages by RNase E either sterically (panel A) or by occluding potential sites into double-stranded conformations (panel B). Additionally, panel C demonstrates that the position of a stable stem-loop structure at the immediate 5'-end can decrease significantly the rate of cleavage at internal single-stranded sites. In contrast, the schematic in panel D shows that R N A structure can anchor potential RNase E cleavage sites into single-stranded conformations. Arrows indicate potential RNase E cleavage sites. The Rne/Ams protein is denoted by the black circle. Bent arrows mark blocked cleavage events. -11-Mackie, 1992) (Figure 1.2D). Thus, the position rather than the presence of a stem-loop can have a significant effect on cleavage rates by RNase E. In this regard, several authors have demonstrated that positioning of R N A secondary structure at the immediate 5'-terminus greatly alters the rate of endonucleolytic attack at internal single-stranded cleavage sites by RNase E (Emory and Belasco, 1990; Emory etal, 1992; Bouvet and Belasco, 1992; Mackie et al, 1997) (Figure 1.2C). Addition of as little as three unpaired nucleotides to stem-loop modified 5-ends restores the RNase E cleavage at internal sites. Moreover, covalent circularization of R N A substrates (9S R N A and rpsT mRNA) has demonstrated an absolute requirement for free 5-ends by RNase E (Mackie, manuscript submitted). Taken together, these results show that the role of R N A structure in directing RNase E cleavage is much more complex than previously imagined. 1.2.1.2 RNase III RNase III, from Escherichia coli, was the first enzyme to be discovered that was capable of cleaving double-stranded RNA. Its primary role in the cell is to process the large precursor 3OS R N A into the immediate precursors to 23 S and 16S rRNAs, however, it also performs a number of other important functions (reviewed in Court, 1993 and Nicholson, 1996). RNase III plays an important role in processing many bacteriophage RNAs including: phage T7, phage T4 and lambda (reviewed in Court, 1993 and Nicholson, 1996). In some cases, processing may enhance the rate of translation of some bacterial and bacteriophage mRNAs (e.g. T7 gene 0.3) by cleaving stem-loop structures which otherwise impede the binding of ribosomes to the Shine-Dalgarno sequence (reviewed in Court, 1993). In addition, RNase III cleavage plays an important role in executing antisense control of plasmid D N A replication (Tomizawa, 1993) and IS70 transposition (Case et al, 1990) by cleaving -12-sense-antisense R N A hybrids. Finally, RNase III can initiate the decay of a number of polycistronic mRNAs including: rnc-era-recO, rpsO-pnp, and the metY-nusA-infBoperons (reviewed in Regnier and Grunberg-Manago, 1990). Nevertheless, two lines of evidence suggest that RNase III does not play a general role in mRNA turnover. First, null mutations in the rne gene (rncr.Kari) are viable and display only a mild phenotype (Babitzke et al, 1993). Second, most mRNAs lack the complex double-stranded stem-loop structure required for RNase III cleavage or contain inhibitory "anti-determinants" (reviewed in Nicholson, 1996; Zhang and Nicholson, 1997). RNase III activity requires a divalent cation, preferably M g 2 + , as an essential cofactor for its phosphodiesterase activity. Natural substrates consist of 20 bp of double-stranded RNA, representing approximately 2 turns of an A-form double helix (Figure 1.3 A) (Krinke and Wulff, 1990). RNase I l l -mediated catalysis usually proceeds through a coordinated double cleavage releasing products containing 5'-monophosphates and 3'-hydroxyl termini, with a two nucleotide 3' overhang (Figure 1.3A and 3B) (Court, 1993; Nicholson, 1996). Structural features, such as internal loops may determine the reactivity of double-stranded RNA. For example, an asymmetric loop can restrict RNase Ill-mediated cleavage to only one side of the helix (Figure 1.3B) (Chelladurai etal, 1993). Indeed, many natural RNase III substrates contain deviations from the typical A-form helices, such as non-canonical base pairs, near the site of cleavage (Schweisguth et al, 1993). The role of base pair sequence in determining RNase III cleavage sites has been somewhat controversial. The finding that RNase III can digest homopolymeric double-stranded R N A (releasing products which range from 12-15 bp) demonstrates that it is not a base-specific enzyme (Robertson, 1982). However, a number of substrates exhibit a short, conserved base pair element (CUU/GAA) proximal to the cleavage site (Krinke and Wulff, 1990). Sequence alignments of several RNase III -13-A 5 ' -WN AGWGNNCWU (NNNN)AWGNNCWCUNW -3' 3'- w n UCwC n n GwA (n nn n) UwC n n GwGA n w -5' f B C A G A C G U A U G A U C G U G G C G C A U A_y_ A U y A A U C G y A A U A U C G A A G C G C G U A U G C A U U U . C A G A C G U A U G A U C G U G G C G C A U A U C u U C 3' A G V A A A U A U C G A A G C G C G U A U G C A U 5 . . . U U . . ..3" Figure 1.3 RNase III consensus processing signals. In panel A, the rather weak consensus sequence for an RNase UJ processing signal is shown. The overall length is approximately 22 base pairs or two turns of the A-form helix. The letters N/n represent any base pair while W/w represents A / U or U / A base pairs. The bracketed region can contain any nucleotide, may be of variable length and does not require any Watson-Crick base pairs. Arrows denote the cleavage sites. An RNase III processing signal from bacteriophage T7 R l . l is shown on the right side of panel B . Introduction of mutations which form new Watson-Crick base-pairs abolishes the internal loop and restores the double cleavage. This figure was adapted from Nicholson (1996); see also Krinke and Wulff (1990). -14-processing signal sequences suggests a loose consensus motif containing a hyphenated dyadic symmetry shown in Figure 1.3 A (Krinke and Wulff, 1990). Substitutions within this element do not block accurate cleavage, although they may reduce the reactivity of double-stranded R N A by weakening the RNA-RNase III interaction (Chelladurai et al, 1991; Chelladurai et al, 1993). In addition, sequence alignments also revealed the exclusion of a number of specific Watson-Crick base pair sequences at defined positions relative to the cleavage site (Zhang and Nicholson, 1997). These sequences, referred to as "antideterminants", have been shown to inhibit cleavage in model substrates in vitro by perturbing RNase III binding (Zhang and Nicholson, 1997). The presence of "antideterminants", together with local tertiary structure, are thought to confer site specificity and may serve to protect double-helical elements in other R N A molecules with essential functions (Zhang and Nicholson, 1997). Structurally, RNase III is a homodimer consisting of two subunits of 25.6 kDa (226 amino acids) (Dunn, 1976; Nashimoto and Uchida, 1985). The rnc gene has been cloned (Nashimoto and Uchida, 1985) and the protein overexpressed and purified to homogeneity by double-stranded R N A chromatography (reviewed in Nicholson, 1996). Structure-function analyses demonstrate that the RNase III protein can be divided into three domains: the catalytic domain, the dimerization domain and the double-stranded RNA-binding domain (Nicholson, 1996). The double-stranded RNA-binding domain consists of 65-70 amino acids located in the C-terminal half of the protein (St. Johnston et al, 1992). Approximately 36 amino acid residues are conserved amongst a group of at least nine different proteins which includes, among others, the mammalian double-stranded R N A activated protein kinase (St. Johnston etal, 1992). The structure of the double-stranded RNA-binding domain of RNase III was recently solved by N M R techniques and was found to have an aPPP<x structure -15-(Kharrat etal, 1995). The domain resembles a tightly folded ellipsoid, with the two helices located on one side of an antiparallel P-sheet. Mutagenesis of the double-stranded RNA-binding motif from the Drosophila staufen protein suggests that the a2 helix contains residues which specifically contact the double-stranded R N A (Bycroft et al, 1995). The search for RNase UI homologs in yeast has uncovered at least three activities capable of degrading double-stranded R N A in vitro (Nicholson, 1996). The pacl gene from Schizosaccharomycespombe encodes a 41 kDa double-stranded RNase which shares some similarity (approximately 25% similar amino acid sequence) with the C-terminal portion of E. coli RNase III (lino et al, 1991). Although at least one study suggests that Pacl may be involved in yeast R N A metabolism (Rotondo et al, 1995), the finding that antibodies to RNase III do not react with Pacl and the finding that pacl and rnc fail to complement each other in reciprocal experiments suggests that these two proteins are not close functional homologues (lino et al, 1991). The rnt-1 gene of Saccharomyces cerevisiae shows 20% identity with the full-length rnc gene and 23% identity with the C-terminal half of pacl. An additional conserved 11 amino acid motif was also identified within the proposed catalytic domain of all three proteins (lino et al, 1991; Elela et al, 1996). Moreover, the rnt-1 gene product is a double-stranded RNase in vitro and is required for U3 snoRNP-dependent cleavage of 28 S rRNA in vivo (Elela et al, 1996). Thus, RNT1 appears to be a functional eukaryotic RNase UI homologue. Double-stranded R N A cleaving activities also exist in mammalian cells, but it is not clear whether these activities are functionally or mechanistically similar to RNase III (reviewed in Nicholson, 1996). -16-1.2.1.3 RNase P RNase P was the first ribonuclease shown to be an R N A processing enzyme (Robertson et al, 1972). RNase P catalyzes the endonucleolytic removal of 5-leader sequences from precursor tRNAs to generate the mature 5' ends of tRNAs. As all tRNAs must undergo this processing step it is not surprising that RNase P is ubiquitous amongst prokaryotes and eukaryotes (Pace and Brown, 1995). Interest in this enzyme as a contributing factor to mRNA stability was recently stimulated by the finding that a specific RNase P cleavage stabilizes the his operon mRNA from Salmonella typhimurium (Alifano etal, 1994). RNase P is a ribozyme composed of an R N A subunit ( M l RNA, 377 nucleotides) and a protein subunit (C5 protein, 119 amino acid residues) in a 1:1 complex (Talbot and Altman, 1994). The M l R N A , encoded by the rnpB gene, is the catalytic subunit and is sufficient for accurate processing activity in vitro (Guerrier-Takada et al, 1983) while the 14 kDa C5 protein, encoded by the rnpA gene, is required for in vivo activity but can be substituted by high salt in vitro (Guerrier-Takada et al, 1983). The C5 protein is thought to shield charge repulsion between the phosphate backbones of the M l R N A and its substrate and/or stabilizes the M l R N A into its active 3-dimensional conformation (reviewed in Altman, 1990; Cech, 1993; Pace and Brown, 1995). RNase P appears to recognize tertiary structure in the mature domain of pre-tRNAs. The M l R N A establishes contacts with a helix consisting of the coaxially stacked tRNA acceptor stem and the common arm which contains the T Y C loop (shaded portion of Figure 1.4) (Altman, 1990; Pace and Smith, 1990; Pace and Brown, 1995). This portion of the tRNA is sufficient for recognition since "mini-substrates" consisting of the common arm, the 5'-leader and the C C A acceptor are efficiently processed (Altman, 1990; Pace and Smith, 1990). Moreover, there appears to be no strong evidence -17-RNase P Cleavage Figure 1.4 Possible recognition elements for RNase P projected onto tRNAphe. The shaded area highlights the coaxial stack formed by the acceptor stem and the common arm. This is the minimal substrate which has been reported for RNase P. Presumably mRNA molecules which are cleaved by RNase P (e.g. the his operon mRNA) must mimic the 3-dimensional features of these stacked helices. The site of RNase P action is denoted by the arrow. This figure was adapted from a review of RNase P structure and function by S. Altman (Altman, 1990). -18-for Watson-Crick base pairing between the M l R N A and its substrate during processing. Presumably, mRNAs which mimic the correct three-dimensional conformation may also be substrates for RNase P (Alifano etal, 1994). Mechanistically, RNase P is a phosphodiesterase which hydrolyzes R N A to release products which exhibit 5'-monophosphate and 3-hydroxyl termini. Biochemical studies demonstrate that the enzyme may require three M g 2 + ions for catalysis (Smith and Pace, 1993). As in the case for RNase E (Cormack et al, 1993), Ca 2 + supports substrate binding but will not support the processing activity (Smith et al, 1992). It is hypothesized that the basis for catalysis may lie in the coordination of the M g 2 + ions in the correct orientation by the M l RNA. (Smith et al, 1992; Smith and Pace, 1993) 1.2.2 The 3 '-exonucleases 1.2.2.1 Polynucleotide phosphorylase (PNPase) Polynucleotide phosphorylase was the first enzyme discovered which was capable of synthesizing oligoribonucleotides with a 3', 5-phosphodiester bond. In the forward reaction polyribonucleotides are synthesized from nucleoside diphosphates with the release of inorganic phosphate (reviewed in Littauer and Soreq, 1982; Littauer and Grunberg-Manago, 1998). This property of the enzyme made it an instrumental tool for the synthesis of model nucleic acids (heteropolymers and triplet nucleotides) which led to the establishment of the genetic code (reviewed in Littauer and Soreq, 1982; Littauer and Grunberg-Manago, 1998). In the reverse reaction, PNPase is a processive 3-exonuclease which phosphorylyzes single-stranded R N A to release nucleoside diphosphates (reviewed in Littauer and Soreq, 1982; Littauer and Grunberg-Manago, 1998). Under physiological conditions, the reverse reaction is favoured. Hence, in vivo, PNPase functions as a 3--19-exonuclease (Littauer and Soreq, 1982; Littauer and Grunberg-Manago, 1998). The primary function of PNPase is in the exonucleolytic removal of R N A fragments during the terminal stages of mRNA decay (see section below) (Donovan and Kushner, 1987). Several observations, however, suggest that this enzyme may have additional roles in the cell. First, E. coli strains which harbour a Tn5 insertion within the pnp gene are more susceptible to tetracycline, suggesting that PNPase may play a role in regulating some aspect of the bacterial membrane or cell surface (McMurray and Levy, 1987). Second, strains deficient in both PNPase and a second phosphorolytic enzyme, RNase PH, display a cold-sensitive phenotype (Zhou and Deutscher, 1997). At low temperatures (31 °C) ribosome structure and function are severely impaired suggesting that these enzymes play an important role in ribosome biogenesis (Zhou and Deutscher, 1997). Finally, repair of the 3'-terminal - C C A sequence of tRNA usually requires the action of tRNA-nucleotidyl transferase. However, poly(A) polymerase I and PNPase can participate in the process of 3'-end-repair and are required to maintain functional tRNA levels in strains deficient for tRNA-nucleotidyl transferase (Reuven etal, 1997). The pnp gene has been cloned and sequenced (Regnier et al, 1987). The open reading frame commences with a non-conventional U U G start codon and encodes the 711 amino acid a-subunit of PNPase (Regnier etal, 1987). PNPase has been successfully purified from several different sources by a number of chromatographic techniques including immobilized dye chromatography and R N A affinity chromatography (Barbehenn etal, 1982; Soreq and Littauer, 1982; reviewed in Littauer and Soreq, 1982). The 77.1 kDa monomer is highly acidic and runs anomolously as an 85 kDa protein through SDS-polyacrylamide gels (Soreq and Littauer, 1978). Functionally, PNPase is an a3 homotrimer with a molecular mass of approximately 231 kDa (Portier etal, 1973; Portier, 1975). -20-Additional high molecular weight forms of the enzyme have been detected during purification by gel filtration and by sucrose gradient centrifugation (Portier et al, 1973; Portier, 1975). These complexes are composed of three a-subunits and two P-subunits (a3p2) with a molecular mass of approximately 323 kDa. The P-subunit of PNPase was later identified as enolase, an essential glycolytic enzyme which catalyzes the conversion of 2-phosphoglycerate to phosphoenol pyruvate (Portier, 1975). As described above (see section), a significant fraction of cellular PNPase also exists in the degradosome complex (Carpousis etal, 1994, Yyetal, 1994; 1996; Miczak et al, 1996). Some PNPase is often detected in conventionally purified GroEL, a chaperone involved in protein folding and assembly (Ybarra and Horowitz, 1996). The role, if any, in PNPase or degradosome assembly/ activity remains unknown. PNPase requires M g 2 + or a closely related divalent cation as an essential cofactor for its phosphorolytic activity (reviewed in Littauer and Soreq, 1982). The enzyme is able to degrade short oligoribonucleotides (< 11 residues) by a non-processive mechanism, however, upon binding to single-stranded polyribonucleotides £ 12 residues in length, PNPase undergoes a transition from a non-processive (distributive) state to processive phosphorolysis (non-distributive) associated with tight binding (reviewed in Littauer and Soreq, 1982). Interestingly, a 62 kDa amino-terminal proteolytic fragment of PNPase is not able to bind oligonucleotides longer than 11 residues and displays reduced processivity without altering the quaternary structure of the enzyme (Thang et al, 1971; Guissani and Portier, 1976). These results suggested that PNPase contains two separate RNA-binding domains, one at the active site and a second domain involved in processivity. A model to describe the behaviour of PNPase is shown in Figure 1.5. Two subunits combine to form the processive RNA-binding pocket (lines in Figure 1.5) while the third subunit is involved in catalysis -21-<11-nts Figure 1.5 Processive versus non-processive RNA degradation by PNPase. In panel A, a potential model for the processive mode of phosphorolysis is shown. The circles represent the three a-subunits of PNPase. The short lines represent a high affinity RNA-binding domain, while the triangles represent a lower affinity RNA-binding domain which is part of the active site. When single-stranded polyribonucleotides are greater than 12 nucleotides, the high affinity sites of two subunits form an extended RNA-binding pocket. The third subunit is orientated in such a way that only the active site is engaged in RNA-binding. In panel B, a potential model for non-processive phosphorolysis is depicted. When polyribonucleotides become shorter than 11 nucleotides, only the low affinity RNA-binding domain of the active site is contacting the substrate. The enzyme dissociates from its substrate after each cleavage. The shaded circles indicate PNPase subunits which are not engaged in RNA-binding or catalysis. -22-via a second lower affinity RNA-binding site (triangles in Figure 1.5). When the single-stranded polyribonucleotide becomes shorter than 12 unpaired nucleotides the high affinity RNA-binding sites are no longer filled and the enzyme dissociates after each round of catalysis. This model is supported by the finding that the a-monomer blotted to nylon membranes cannot bind R N A during "northwestern" RNA-binding experiments (Py et al, 1996). The a-subunit of PNPase contains at least two recognizable RNA-binding motifs: a K H R N A -binding domain (residues 557-591) (Musco etal, 1996) and an SI RNA-binding domain (residues 619-691) (Bycroft et al, 1997) (see Figure 1.6). The SI RNA-binding domain has been discussed (see section 1.2.1.1). The K H RNA-binding motif, named after the mammalian hnRNP K protein (KH-K-Homology), is composed of approximately 50 amino acids (Mattaj, 1993). The coordinates of the PNPase K H domain deposited in the SWISS-PROT data base are slightly smaller than the predicted 50 amino acids. Presumably, the primary sequence alignments only detected the most conserved amino acids in the K H motif. This motif is also found in a large family of proteins which includes ribosomal protein S3, the NusA transcription factor and MER1 , a yeast splicing regulator (Mattaj, 1993). Its distinguishing feature is the placement of hydrophobic amino acid residues (I, L , V) at regular intervals and a central I G X X G peptide motif (Mattaj, 1993). The K H domain folds into a stable Pacq3pa conformation consisting of a three-stranded antiparallel P-sheet separated by two helical regions (Musco et al, 1996). There is also evidence that, in addition to mediating RNA-protein interactions, K H domains may mediate protein-protein interactions (Chen et al, 1997). Thus, the K H domain of PNPase may be a possible self-association site or RNase E binding site. Alternatively, the K H domain may contribute to the active site and/or processive RNA-binding site of PNPase. -23-RNase E N 3 5 1 2 5 S1-RBD 1061 6 0 0 7 5 0 R-RBD 1 PNPase N C RNase II N L 711 5 5 7 591 6 1 9 691 KH-RBD S1-RBD 644 S1-RBD Figure 1.6 Localization of RNA-binding domains in three ribonucleases from E. coli. A schematic representation of the linear amino acid sequence of RNase E, PNPase and RNase II. The putative Arg-rich RNA-binding domain of RNase E is denoted by the open box while the SI RNA-binding domains in all three proteins are denoted by the black boxes (Bycroft et al, 1997). The KH-RNA-binding domain (denoted by the open box) of PNPase is also shown. Amino acid coordinates are shown below except in the case of RNase II, where the SI domain was positioned based on the//, influenzae alignment (residues 571-659) (Bycroft etal, 1997). -24-1.2.2.2 RNase II RNase II is a potent 3-exonuclease which hydrolyzes single-stranded R N A processively in the 3' to 5' direction to release mononucleotides (reviewed in Shen and Schlessinger, 1982). It is responsible for 2:90% of all exonucleolytic activity in crude extracts from E. coli while PNPase accounts for the remaining ~ 10% (Deutscher and Reuven, 1991). Interestingly, in the soil bacterium, Bacillus subtilis, the majority of the exonucleolytic activity in crude extracts is phosphorolytic rather than hydrolytic (Deutscher and Reuven, 1991). Phosphorolytic mRNA decay maintains a high-energy phosphate bond in the released nucleotide product (NDP versus NMP) and, therefore, may play an important role in fulfilling cellular energy demands under sub-optimal growth conditions. In E. coli, strains containing single rnb (RNase II) or pnp (PNPase) mutant alleles are viable; however, strains containing both mutations are inviable (Donovan and Kushner, 1987). Moreover, in strains deficient for both RNase II and PNPase, discrete mRNA fragments are stabilized significantly at non-permissive temperatures (Arraiano et al, 1988). Although both enzymes can play a limited role in processing the 3-ends of pre-tRNA (Li and Deutscher, 1994; L i and Deutscher, 1996), their main role is in mRNA metabolism. The rnb gene, which encodes RNase II, has been cloned and sequenced (Zilhao et al, 1993). As in the case for the me gene, several minor errors, due to compressions in the D N A sequence, resulted in an underestimation of the size of the expected protein (598 amino acid residues) (Zilhao etal, 1993).The corrected sequence encodes a 644 amino acid protein (molecular mass of 72.5 kDa) which contains an SI RNA-binding domain located at its immediate C-terminus (Bycroft et al, 1997). Complementation analyses with truncated versions of the rnb gene suggested that this region is necessary, although, not sufficient for enzymatic activity (Zilhao et al, 1993). Unlike PNPase, -25-RNase II functions as a monomer and does not copurify with other proteins. This, however, does not occlude the possibility that weak or transient interactions take place between RNase II and the degradosome and/or other factors in vivo. RNase II requires a divalent cation, preferably Mg 2*, for its hydrolytic activity and is activated by the presence of monovalent K + ions (reviewed in Shen and Schlessinger, 1982). Exhaustive studies of the mechanism of RNase U on homopolymeric substrates show that poly(A) and poly(U) substrates are degraded much more quickly than poly(C), consistent with RNase II being a single-strand specific enzyme (Cannistraro and Kennell, 1994). Interestingly, under conditions which promote folded conformations of poly(C) RNA, RNase II processivity breaks down and the enzyme dissociates every 12 nucleotides. These results suggest that R N A secondary structure may disrupt the processive nature of RNase II. Moreover, RNase II dissociates from all homopolymers when the substrate becomes less than 10 to 15 nucleotides (Cannistraro and Kennell, 1994). RNase II can hydrolyze these smaller oligoribonucleotides non-processively to release a 2 to 4 nucleotide core which is resistant to further attack (Spahr, 1964; Singer and Tolbert, 1964; 1965). As in the case for PNPase, RNase II is predicted to have two separate RNA-binding sites (Cannistraro and Kennell, 1994). One at the catalytic site and a separate RNA-binding domain involved in processivity. The S1 RNA-binding domain located at the immediate C-terminus of the protein may be involved in one or both of these functions. The processive activities of both RNase II and, to a lesser extent, PNPase are blocked by stem-loop structures resulting in the stabilization of the upstream R N A sequence (Mott et al, 1985; McLaren et al, 1991; reviewed in Higgins et al., 1992) (Figure 1.7). While genetic evidence suggests that RNase II and PNPase are functionally redundant (Donovan and Kushner, 1987), the two -26-Exonucleolytic Digestion «• — * 0 r « 0 RNase II / PNPase ^ ^ RNase II / PNPase 1. Stalling 1. Stalling 2. Dissociation 2. Dissociation or (_J CJj Localized Melting F i g u r e 1.7 TTie e#ecf o/"secondary structure on 3'-exonucleolytic degradation.(Panel A) Single-stranded R N A is rapidly degraded by either RNase II or PNPase (denoted by the "pacman" in step 1). The processive action of both RNase II and, to a lesser extent, PNPase breaks down 3' to the base of stable R N A secondary structure. At this point the stalled 3'- exonuclease (circles) may dissociate from its substrate. Alternatively, the 3'-exonucleases remain bound to their substrates until localized melting of the stem-loop structure allows processive degradation to re-initiate. -27-A B 5' Figure 1.8 RNase II as a repressor of mRNA decay. Two alternative models, are depicted in panels A and B. In panel B, RNase II stalled at the 3' base of the stem-loop structure sequesters the 3'-end, thus preventing further exonucleolytic attack. In the second model, modification of single-stranded 3'-ends (see Figure 1.7) by RNase II generates "new" RNAs which contain 3'-terminal stem-loops. These substrates are no longer susceptible to further attack by PNPase and possibly other nucleases. RNase II is denoted by the shaded circle while other 3-exonucleases are shown as open circles. -28-enzymes exhibit differential sensitivities to R N A secondary structure (McLaren et al, 1991; Guarneros and Portier, 1991). That is, PNPase, rather than RNase II appears to be required for the efficient degradation of highly structured RNAs (Figure 1.7). Observations of the decay of several RNAs, includingRNA1 (Xu etal, 1993; Xu and Cohen, 1995), the antisense regulator of colEl-type plasmid D N A replication, RNA-OUT (Pepe et al, 1994), the antisense regulator of Tn/0/7570 transposition, and the rpsO (Braun et al, 1996) and rpsT (Mackie, 1989) mRNAs, encoding ribosomal protein S15 and S20 respectively, support this hypothesis. Surprisingly, in strains deficient for RNase II, both RNA-OUT and the rpsO mRNA are less stable than in otherwise isogenic strains (Pepe er al, 1994; Hajnsdorf et al, 1994). These observations lead to the hypothesis that RNase II acts as a repressor of R N A decay by forming a stable RNA-protein complex upon stalling 3' to the base of a stable stem-loop structure (Pepe et al, 1994; Hajnsdorf et al, 1994). The complex sequesters the 3'-end and prevents additional attack by PNPase and possibly other ribonucleases (Figure 1.8A). An alternative explanation suggests that modification of 3'-ends by removal of unpaired nucleotides results in the formation of an R N A molecule which is resistant to further exonucleolytic attack (Figure 1.8B). This interesting paradox provides additional evidence that subtle differences exist in the substrate specificity of RNase II and PNPase. An exciting related discovery is the recent isolation of the "exosome", a conserved complex of five essential proteins in eukaryotes (Mitchell et al, 1997). Amazingly four of the conserved proteins are homologous to known bacterial 3'-exonucleases. One protein Rrp44p (Dis3p) is homologous to RNase II and displays processive 3'-exonucleolytic activity in vitro (Mitchell etal, 1997). Al l components of the "exosome" are involved in the processing of 5.8 S rRNA (Mitchell et -29-al, 1997) and three members of the complex (related to RNase PH) are involved in the 3-5'-dependent pathway of mRNA decay in eukaryotes (Jacobs et al, 1998). RNase II also displays approximately 60% similarity to VacB, a protein that is involved in some aspect of virulence, from Shigella flexneri (Dmochowska et al, 1995). Recently, the E. coli homologue of VacB was cloned and was found to encode RNase R, a poorly characterized 3-exonuclease (Cheng et al, 1998). Although the role of a ribonuclease in bacterial pathogenesis remains unclear, the finding that a pnp rnr double mutant is inviable suggests that RNase R has additional roles in R N A metabolism (Cheng et al, 1998). RNase II also exhibits some similarity to DSS-1, a protein that is involved in yeast mitochondrial biogenesis (Dmochowska et al, 1995). The significance of the similarity is not yet clear. 1.2.2.3 Oligoribonuclease Oligoribonuclease is a relatively uncharacterized 3-exonucleolytic activity which shows a preference for short R N A oligonucleotides. This enzyme may play a potentially important role in eliminating short oligonucleotides such as limit products (< 5 nucleotides) formed by digestion of R N A fragments by RNase E, RNase II or PNPase. Oligoribonuclease is a unique enzymatic activity, separate from all known 3'-exonucleases in E. coli (Yu and Deutscher, 1995). Recently, oligoribonuclease has been purified as an al dimer of 40 kDa (Zhang etal, 1998). Amino-terminal sequencing of the purified protein identified the gene encoding oligoribonuclease as yjeR (pm) (Zhang etal, 1998). Database searches suggest that oligoribonucleases are ubiquitous in nature. The degree of identity between the£ . coli and some eukaryotic homologues approaches 50% suggesting that oligoribonuclease function is highly conserved (Zhang et al, 1998). Moreover, attempts to disrupt -30-the yjeR (prri) gene by kanamycin insertion have proven unsuccessful suggesting that the gene is essential (Zhang et al, 1998). Although an earlier report claimed that oligoribonuclease copurifies with PNPase (Yu and Deutscher, 1995), there is no indication that this enzyme is associated with the degradosome. 1.2.3 Regulation of ribonuclease expression The organization of the genes encoding the ribonucleases is quite varied and the regulation of their expression remains poorly understood. Two enzymes of mRNA decay, RNase E and RNase II (Claverie-Martin etal, 1991; Zilhao et al, 1993) are monocistronic while RNase III and PNPase are transcribed as part of polycistronic mRNAs (Takiff et al, 1989; Evans and Dennis, 1985). Another level of complexity arises from the finding that RNase II can be transcribed from one of two promoters (Zilhao et al, 1996b). Additionally, PNPase is transcribed as either a polycistronic message, an RNase III processed mRNA, or from a second weak internal promoter (Evans and Dennis, 1985). It is not fully appreciated why these ribonucleases would require multiple promoters or what might be the significance of co-transcribing a ribosomal protein (SI5) with PNPase. Presumably, this organization serves to link PNPase expression to overall growth rates. Moreover, genetic evidence has implicated PNPase in ribosome assembly (Zhou and Deutscher, 1997). It is likely that maintaining specific ratios of the various activities is required for correct processing and balanced decay of R N A molecules within the cell. In order to maintain the correct relationship, the expression of one ribonuclease will likely impinge upon the expression of the other enzymes. A classic example of inter-regulation is the 10-fold overexpression of PNPase observed in strains deficient for RNase III (Takata et al, 1987; Portier et al, 1987). An interesting facet of this -31-autoregulatory mechanism is the finding that the PNPase inhibits translation of its own message and induces the degradation of its RNase Ill-processed message (Robert-Le Meur and Portier, 1994). Not surprisingly, PNPase and RNase II also appear to be inter-regulated by a reciprocal mechanism (Zilhao et al, 1996a). Strains deficient for PNPase were found to have a 2-2.5-fold increase in RNase II activity while inactivation of RNase II resulted in an increase in PNPase activity, albeit to a lesser degree (Zilhao etal, 1996a). Due to the prominent role of RNase E in initiating mRNA decay mE. coli, it is likely that RNase E contributes to the decay of all ribonucleases involved in mRNA decay, as shown for RNase II (Zilhao et al, 1995). Additionally, since RNase E cleavage is required for the maturation of M l R N A from its primary transcript (Lundberg and Altman, 1995), RNase E may also have important effects on the activity RNase P. Finally, in addition to initiating the decay of other mRNAs, both RNase E (Mudd and Higgins, 1993; Jain and Belasco 1995) and RNase III (Portier et al, 1992) are able to autoregulate their own expression upon cleaving their respective messages. The long leader sequence of the rne message is particularly susceptible to cleavage by RNase E (Jain and Belasco, 1995). Presumably, the multi-layered control of ribonuclease expression allows the bacteria a wide degree of flexibility in their response to environmental stimuli. 1.3 Other enzymes of mRNA decay 1.3.1 Poly(A) polymerases Poly(A) polymerases catalyze the template-independent addition of adenylate residues from ATP to the 3'-hydroxyl termini of R N A molecules with the release of PPi. The enzyme requires M g 2 + ions for activity but no other cofactor. Although poly(A) polymerase was purified from E. coli in the early 1960's (August etal, 1962) and several papers reporting the presence of polyadenylated RNAs -32-appeared shortly thereafter (Nakazato et al, 1975; Srinivasan et al, 1975), the concept that prokaryotic mRNAs are polyadenylated was ignored for almost 25 years (reviewed in Sarkar, 1997). This reasoning was most likely due to the established role for polyadenylation in eukaryotic mRNA metabolism and the lack of any apparent physiological parallel in prokaryotes. Moreover, difficulties in characterizing rather unstable bacterial mRNAs relegated the phenomenon of bacterial polyadenylation to mere curiosity. Renewed interest was generated, many years later, by two different groups who simultaneously determined that a mutation named pcnB which resulted in a decrease in plasmid copy number (Lopilato etal, 1985), affected the stability of RNA1, the antisense regulator of colEl-type replicons (He et al, 1993; Xu et al, 1993). The pcnB gene from E. coli was cloned (March et al, 1989; Liu and Parkinson, 1989) and subsequently overexpressed and found to encode the major poly(A) polymerase in E. coli (PAP I) (Cao and Sakar, 1992a). More recently, PAP I activity has been shown to destabilize the rpsO mRNA, encoding ribosomal protein S15 (Hajnsdorf et al, 1995), and mRNA in general (O'Hara et al, 1995). While the role of polyadenylation in mRNA degradation is not completely understood, the addition of 3'-single-stranded extensions catalyzed by PAP I may provide a platform for the 3'-exonucleases (Cohen, 1995) and possibly the degradosome (Ingle and Kushner, 1996) to attack otherwise resistant RNAs. The function of bacterial poly(A) tails in mRNA degradation clearly distinguishes their role from eukaryotic poly(A) tails which promote mRNA stability, R N A transport and translation. In addition to physiological function, several other differences are observed between the process of prokaryotic and eukaryotic polyadenylation. Poly(A) tails in bacteria are much shorter (6-60 nts versus 80-200 nts) and are only associated with 2-50% of the molecules of a given mRNA -33-species (reviewed in Sarkar, 1997). Moreover, polyadenylation in bacteria is a relatively indiscriminate process that can occur at all 3'-ends and, therefore, does not require a consensus sequence as in eukaryotes (Cao and Sarkar, 1992b; Haugel-Nielsen et al, 1996; reviewed in Sarkar, 1997). ThepcnB gene has been sequenced (Liu and Parkinson, 1989) and shows no homology with eukaryotic poly(A) polymerase. It does, however, share significant homology and some functional overlap with tRNA-nucleotidyl transferase (Masters et al, 1990; Reuven et al, 1997). The 52 kDa protein has been overexpressed and purified as either a His-tagged or GST (glutathione-S-transferase) fusion protein. The deduced translation start site appears to be 17 amino acids upstream of the amino-terminal lysine residue suggesting that the protein may be either processed or susceptible to proteolysis during purification (Cao and Sarkar, 1992a). Interestingly, the coding region of the pcnB gene also starts with a rare U U G start codon as does the pnp gene encoding PNPase (Cao and Sarkar, 1992a) The finding that strains deficient for PAP I maintain significant quantities of poly(A) polymerase activity suggests that their may be additional poly(A) polymerases in E. coli (Cao et al., 1996). Recently, a second, poly(A) polymerase, PAP II, from E. coli was partially purified and characterized (Kalapos etal, 1994). PAP II is a hydrophobic protein with a molecular mass of about 35 kDa. The finding that PAP II-mediated addition of adenylate residues shows a preference for poly(A) tails (Kalapos et al, 1994) suggests that PAP II is involved in poly(A) extension or maintenance. Nonetheless, the role of PAP II, if any, in mRNA metabolism remains unsettled. -34-1.3.2 RNA helicases E. coli encodes at least five proteins (Dead, SrmB, DbpA, RhlB and RhlE) (Kalman etal, 1991) that belong to a superfamily of putative R N A helicases containing the signature DEAD-box motif (Under etal, 1989; Schmid and Linder, 1992). The DEAD-box (Asp-Glu-Ala-Asp) is thought to be a special form of the Walker B motif (Walker et al, 1982) and, therefore, may be involved in ATP binding or hydrolysis (Linder etal, 1989; Schmid and Linder, 1992) . The prototypical member of the DEAD-box superfamily is the eukaryotic translation initiation factor eIF-4A which is an RNA-dependent ATPase (Rozen etal, 1990). Moreover, eIF-4A is one of only a handful of R N A helicases that are mechanistically similar to D N A helicases. The DEAD-box proteins are, therefore, preferentially referred to as putative R N A helicases or "unwindases" until true helicase functions are assigned. The recent finding that the putative R N A helicase, RhlB, copurifies with the degradosome complex (Py et al, 1996; Miczak et al, 1996) is particularly exciting since RNase E, PNPase and RNase II are specific for single-stranded RNA. Two models have been proposed to rationalize the role of RhlB in the degradosome. One model suggests that RhlB unwinds R N A secondary structure to permit access by RNase E to cleavage sites which are normally occluded by R N A secondary and possibly higher-ordered structure (Miczak et al, 1996). An alternative model suggests that RhlB alleviates structural impediments to the 3'-exonucleases (Py et al, 1996). In the former model, the helicase would translocate 3'-5' ahead of the 3'-exonucleases while the latter model would be better served by a 5'-3' translocation mechanism. Py etal. have shown a requirement for ATP and RhlB in the degradation of REP sequences by PNPase (Py et al, 1996). Moreover, antibodies raised against the RhlB protein inhibit 3'-5' decay -35-through the stem-loop structure (Py et al, 1996). Further experimentation is required to determine whether RhlB functions as an R N A helicase and to determine the directionality of the helicase activity. Unfortunately, our understanding of this potentially important protein is complicated by the lack of a definitive assay system and by the fact that rhlB mutants are inviable. Of additional interest, is the finding that 3-5' degradation of yeast mRNA by the exosome also requires the Ski2 protein, a member of the DEVH-box family of putative R N A helicases (Jacobs et al, 1998). A second R N A helicase, DBP2, has been implicated in the nonsense-mediated mRNA decay pathway in yeast (He and Jacobson, 1995). Thus, it would appear that a role for R N A helicase proteins in mediating mRNA decay is relatively conserved between bacteria and yeast. In contrast to RhlB, the other E. coli DEAD-box proteins appear to function in ribosome assembly and translation. Surprisingly, overexpression of SrmB and DeaD lead to a 25-30-fold stabilization of the lacZ mRNA (lost and Dreyfus, 1994). It is unclear whether or not the observed stabilization was due to R N A helicase activity, enhanced translation or an indirect consequence of overexpression. It is more likely that overexpression of RNA-binding proteins hinders the degradative machinery in a relatively non-specific manner. 1.4 Other protein factors Several ancillary factors have been proposed which contribute to the stability of specific target mRNAs. One novel system involves the CsrA (carbon storage regulator) protein which downregulates enzymes involved in glycogen biosynthesis. The CsrA regulator binds specifically to the glgCAP mRNA and facilitates its decay by a yet unknown mechanism (Liu and Romeo, 1997). In one model, the CsrA-RNA complex is recognized directly by an endonuclease, presumably RNase -36-E, or may alter R N A secondary structure, making the R N A more susceptible to attack. Alternatively, CsrA may decrease the frequency of translational initiation exposing the mRNA to the degradative machinery. Interestingly, the CsrA protein is controlled by a novel mechanism which involves an R N A molecule, CsrB (Liu etal, 1997). CsrB R N A is thought to antagonize the effects of CsrA on gene expression by sequestering the CsrA protein into an RNA-protein complex (Liu et al, 1997). Recently, Kushner and his colleagues have isolated a new gene, termed mrsC (for mRNA Stability), in a genetic screen for temperature-sensitive alleles of pnp (Granger et al, 1998). Inactivation of mrsC led to the cessation of cell growth and net R N A synthesis (Granger et al, 1998). In addition, at the non-permissive temperature the half-lives of both total pulse-labeled R N A and several specific mRNAs were dramatically increased (Granger et al, 1998). The mrsC gene was later found to be allelic to hflB/ftsH which encodes a membrane-associated ATPase and protease (Wang et al, 1998). Although it has been suggested that HflB/MrsC may proteolytically activate a component of the degradative apparatus (e.g., a ribonuclease), the function of this protein in mRNA decay remains unclear (Granger et al, 1998). To my knowledge, no such processing event has ever been demonstrated for a cytoplasmic protein in E. coli. In contrast to destabilizing factors, there may be factors which stabilize mRNAs against endonucleolytic or exonucleolytic degradation. The EIF (exonuclease inhibitory factor) is a poorly characterized protein (apparent molecular mass = 50 kDa) that was isolated as a member of a large complex of proteins bound to the malE-malFKEP sequence (Causton etal, 1994). It is hypothesized that the EIF stabilizes REP stem-loop structures, thus, preventing exonucleolytic degradation of the upstream mRNA (Causton et al, 1994). Whether or not this protein has an authentic role in mRNA stability remains to be demonstrated. Another factor which may stabilize mRNA is the transcription -37-termination factor Rho. Strains which contain a mutant Rho factor, named rho(nusD), show a significant stabilization in the half-life of bulk cellular mRNA (Sozhamannan and Stitt, 1997). Unexpectedly, expression of the pBR322 encoded Rop(Rom) protein restores nearly normal mRNA decay in the rho(nusD) mutant (Sozhamannan and Stitt, 1997). Although the mechanism of Rho stabilization is unknown, these results suggest that the RNA-binding properties of Rho rather than its transcription termination function are required for R N A stability (Sozhamannan and Stitt, 1997). Whether or not this interaction is specific remains to be seen, however, overexpression of two other RNA-binding proteins, SrmB and DeaD, were unable to prolong mRNA decay in the rho(nusD) mutant (Sozhamannan and Stitt, 1997). 1.5 The role of translation The relationship between translation and mRNA stability is very complex and poorly understood. Early experiments revealed that translational inhibitors, such as chloramphenicol which blocks polypeptide chain growth by inhibiting peptidyl transferase, stabilize bulk mRNA half-lives (reviewed in Petersen, 1993). In contrast, antibiotics such as puromycin which promote chain release destabilize mRNAs (Petersen, 1993). Taken together, these results have been interpreted to mean that ribosomes protect mRNAs from decay by blocking both endonucleolytic cleavage and exonucleolytic digestion (Belasco and Higgins, 1988; Higgins etal, 1992). However, it should be noted that certain mRNAs are relatively stable even in the absence of translation because they are intrinsically resistant to the degradative machinery (Russel et al, 1976; von Gabain et al, 1983). More refined experiments have investigated the effects of limiting the frequency of translational initiation by translational repressors or by the introduction of mutations into the Shine--38-Dalgarno sequence (Cole and Nomura, 1986; Comer et al, 1996; Jain and Belasco, 1993; Rapaport and Mackie, 1994; Yarchuk et al, 1991;Yarchuk et al, 1992). As a general rule, mRNAs which are efficiently translated are more stable than mRNAs which are poorly translated. Similarly, uncoupling transcription from translation (e.g., T7 R N A polymerase) can result in destabilization of mRNAs by generating naked regions of R N A behind the R N A polymerase (lost and Dreyfus, 1995). Although there is a good correlation between an mRNA's translatability and its stability, the mechanism of protection remains a mystery. Some clues to this mechanism have come from studies which examined the effect of 5'-terminal secondary structure on mRNA stability. A large stem-loop located at the 5'-terminus of the ompA mRNA can stabilize significantly a number of mRNAs to which it has been fused (Emory and Belasco, 1990; Emory et al, 1992; Hansen et al, 1994). Moreover, in vitro evidence demonstrates that 5'-terminal secondary structures can inhibit cleavage of the rpsT mRNA by RNase E (Mackie et al, 1997). These results suggest that RNase E initially interacts with the 5'-end and scans downstream in the 5' to 3' direction to the first cleavage site. Thus, it is likely that the ribosome provides more than a passive protective cover for the mRNA. Rather, the initiating ribosome competes with the degradative apparatus for the primary inactivation event, either binding at the 5' end and/or endonucleolytic cleavage by RNase E or RNase III. 1.6 Models of mRNA decay For small RNAs and a few unstructured mRNAs, degradation may be initiated directly by 3'-exonucleolytic attack (Figure 1.9A). Nevertheless, for the vast majority of messages, the 3'-end is protected from exonucleolytic attack by R N A secondary structure such as a Rho-independent -39-0 0 0 a 6 8 6 % 5' 3' 51 I *1 5' 3' 5' 3' | 2A | 2B 5' 3 5' 3 Figure 1.9 Conventional models of mRNA degradation. In panel A, a simple model of mRNA decay is shown. For some unstructured RNAs, decay may be initiated directly by the 3'-exonucleases. For the vast majority of mRNAs, shown in panel B, degradation is initiated by endonucleolytic cleavage events (denoted by scissors) followed by exonucleolytic removal of the remaining oligonucleotides to mononucleotides by the 3'-exonucleases (denoted by the "pacmen"). Translating ribosomes, which are thought to compete with endonucleolytic cleavages, are depicted by shaded ellipses. In this model endonucleolytic cleavages allow the 3'-exonucleases to by-pass stem-loops (step 2A). Alternatively, RNase III cleavage may attack the stem-loop directly (step 2B).This model was adapted from Higgins etal. (1992) and Higgins etal. (1993). -40-transcription terminator (Figure 1.9B; top). Thus, most mRNAs must enter a more complex pathway of mRNA decay involving both endonucleases and exonucleases (Figure 1.9B). The consensus model proposes that mRNA decay is initiated by one or more endonucleolytic cleavages catalyzed by RNase E and sometimes RNase III (Babitzke and Kushner^ 1991; Melefors and von Gabain, 1991; Mudd etal, 1990; Portier etal, 1987; Regnier and Grunberg-Manago, 1990). The internal cleavage events provide new free 3'-ends which permit RNase II and PNPase to by-pass structural impediments (Figure 1.9B; step 2A) (Belasco and Higgins, 1988; Higgins et al, 1992). This results in the rapid clearance of the remaining R N A fragments to mononucleotides. In some rare cases, direct attack of a stem-loop structure by RNase III removes a structural impediment to 3'-exonucleolytic decay (Figure 1.9B; step 2B) (Guarneros etal, 1988; Guarneros and Portier, 1991). This model has been invaluable for rationalizing the roles of the endonucleases, exonucleases, R N A secondary structure and to a lesser extent translation in the decay process. Nonetheless, simple models of mRNA decay are incomplete: they are unable to accurately predict initiating events in mRNA decay, they are unable to explain the relative stabilities of different mRNAs and they are unable to account for the fact that decay is usually a concerted "all or none" process. In addition, previous models of mRNA decay do not accurately reflect the role of each ribonuclease or anticipate the requirement for novel mechanisms to degrade highly-folded RNAs (Figure 1.9B; step 2A). The finding that at least two enzymes of mRNA decay (RNase E and PNPase) are associated in a high molecular weight complex (Carpousis et al, 1994; Py et al, 1994) and the finding that poly(A) tails function in mRNA degradation have rendered previous models somewhat obsolete (Hajnsdorf et al, 1995; O'Hara et al, 1995; Haugel-Nielsen et al, 1996). These arguments demonstrate the need to create models which have better predictive power and which incorporate the new information which -41-has emerged in the past 5-10 years (see Coburn and Mackie, 1998). 1.7 Experimental rationale The mRNAs encoding ribosomal protein S20 (356- and 447-nucleotides respectively) are transcribed from two tandem promoters in the rpsTgene. The rpsTmRNA is an ideal system to study mRNA decay in E. coli for several reasons. First, the rpsT mRNAs decay with half-lives typical of most messages from is. coli (PI: 90 seconds; P2: 118 seconds) (Mackie, 1989). Second, the rpsT mRNA is a monocistronic mRNA for which there is a well tested R N A secondary structure model (Mackie, 1992) (see Figure 1.10).Third, the most prominent RNase E cleavage sites have been mapped both in vivo and in vitro (Mackie, 1991; Mackie, 1992) (arrows in Figure 1.10). Finally, the decay of the rpsTmRNA, which depends on RNase E and PNPase, has been well studied in vivo and in crude extracts from E. coli (Mackie, 1992; Mackie and Genereaux, 1993; Rapaport and Mackie, 1994; Coburn and Mackie, 1996b; Mackie et al, 1997). While the ribonucleases have been well characterized, their relative roles in the degradation of model mRNAs remain poorly defined. Furthermore, the role of R N A secondary structure in directing the action of the ribonucleases and the mechanism of degradation of highly structured R N A fragments remains to be elucidated. In order to further study the interactions of the ribonucleases (either free or complexed in the degradosome), and poly(A) polymerase I with natural mRNA substrates, our objective was to reconstitute the decay of the rpsTmRNA in vitro. To accomplish this goal, we have purified several of the key components of the degradative machinery and examined their action on synthetic RNAs derived from the rpsT mRNA. -42-•230 Ilia II c o x X X c-o o - x u - x u - x C - 0 o B - X X a x o "•0-240 C -0 o - c a - x x a a x a c o 1Mb V Via c D X X X o-x 1 4 0 - X - O X c a c 170 C - a O-X U - X X X X 0 -X-1S0 C-0 O-C a o o-x C - 8 „ C-0 a - c c o o - c 1S0 C O X o c o x a '„ o o , \ o - c a - c x c X C - 0 X 320 -IV 2 0 0 . a-c 270 X, a 130 » I x - a c - a o - x 160-c-o w a - c C - 0 0 0 " C O c x x x x a c x c x x a c x c x a x x 190 2 8 0 . X C x a O X o-x 2 9 0 o-x X - 0 c - a a - c X X O X C X a x o a c - a a u a - c X - D a o c - o - 3 3 0 I > c x-o 310 ° - c a - c o-x Y o - c » c VI x c c a x o i 300 a a o o o x c x c x x a x o x o x c c o a o o x x a o o o c c 120 n'o too 5 1 - g a m u a c a c g g a a u a C O 3 6 0 - a x X X X c o c X - 0 c - a O - X . 3 7 0 a - c c I X - 0 3 5 0 - C - 0 340 O - C X I , o - x x x x x c x x x a c c X o a o o o c a a o o x x O-X 4 2 0 X - 0 . o - x 4 1 0 - a - c o - x o - x o - x C X o - x o - c o a o - x 4 4 0 - O - C o - x a - c a - x o - c x c C O - 4 3 0 X - 0 a - c c - a - 3 9 0 x a 4 0 0 - c - a x c a - c c o •330 Vll o o X X Vlb Figure 1.10 Modelfor the structure of the rpsT mRNA, encoding ribosomal protein S20. A model of the secondary structure of the rpsT mRNA is shown above. This model was generated by a combination of single-strand specific enzymatic and chemical probing of the synthetic rpsT mRNA in solution (Mackie, 1992). The major sites of RNase E cleavage are shown with arrows. -43-CHAPTER 2 Cloning, overexpression and purification of the enzymes of mRNA decay 2.1 Introduction In order to determine the relative roles of the ribonucleases and potential ancillary factors in the degradation of mRNA we have constructed strains which overexpress many of the key enzymes of the degradative machinery including the 3'-exonucleases (RNase II and PNPase) and the putative DEAD-box R N A helicase (RhlB). The overexpressed proteins have been purified to near homogeneity by a number of chromatographic techniques. The degradosome, a multi-protein complex containing, RNase E, PNPase, RhlB, and enolase has also been purified by a modified version of previously published methods (Py et al, 1996; Carpousis et al, 1994). Most of these methods have been published elsewhere (Coburn and Mackie, 1996a; Coburn and Mackie, 1998). 2.2 Materials and methods 2.2.1 Enzymes and chemicals All restriction enzymes were purchased from Amersham-Pharmacia or Promega. Modifying enzymes were purchased from Amersham-Pharmacia, Promega, or Gibco-BRL. Taq D N A polymerase was purchased from Promega or New England Biolabs (NEB), while Sequenase™ was purchased from United States Biochemicals (USB). Enzymes were used according to the manufacturers' specifications. Radioactively labeled [cc35S]-dATP (1000 Ci/mmol), for D N A sequencing and poly[8-3H]adenylic acid (400-600 mCi/mmol), for exonuclease assays, were -44-purchased from Amersham-Pharmacia. Growth media were purchased from Difco Laboratories while chromatographic resins were purchased from Bio-Rad or Amersham-Pharmacia. Al l other chemicals were of reagent grade and were purchased from B D H , Bio-Rad, Fisher or Sigma. 2.2.2 Bacterial strains and plasmids TheE. coli strain CF881 (F~ Alac argA trp recB\009 A(xthA-pnc) Arna) was obtained from Dr. M . Cashel (NIFL Bethesda), while strain 18-11 (rna~, rnb', met, rbn, rnf) was obtained from Dr. M.P. Deutscher (University of Connecticut Health Center, Farmington). Strain MV1190 ((Alac-pro), thi, supE, A(sr/-recA)306::Tnl0 (tef) [F' tra D36, pro A B , /aeTqZAM15]) was purchased from Bio-Rad. The vector pET-11 and its host stain BL21(DE3) F" ompT hsdSB (rB"mB") gal dem (DE3) (Studier et al, 1990), were obtained from Novagen. Two plasmids, pRZA17 and pRZA18, containing a 740-bp Aatll-Actll fragment of the rnb gene cloned in opposite orientations (in pT7T318U) were obtained from Dr. C M . Arraiano (I.T.Q.B. -University of Lisbon, Portugal). The D N A sequence contained within these plasmids is located immediately downstream of that for the previously published rnb gene (Zilhao etal, 1993). 2.2.3 Molecular biological methods 2.2.3.1 Isolation of genomic DNA from Escherichia coli Genomic D N A (gDNA) from Escherichia coli was prepared by a method compiled by Dr. Anne Johnson, in this laboratory, which was based on modifications of previously published methods (Marmur, 1961). Cell cultures were grown in Luria-Bertani broth (LB) (Sambrook etal, 1989) at 37°C. Approximately 200-300 mg of cells (750 pi) were harvested by centrifugation from 2.5 ml -45-overnight cultures of strain MV1190. The cell pellet was washed once with 1.0 ml of buffered-saline-E D T A (10 mM Tris-HCl, pH 8.0, 150 mM NaCI, 5 mM EDTA). Cells were collected by centrifugation (3,000 x g for 2 minutes) and resuspended in 0.5 ml of lysis buffer (10 mM Tris-HCl, pH 8.0, 150 m M NaCI, 5 mM EDTA, 15 mM sodium-citrate). Cells were lysed by addition of lysostaphin (Sigma), at a final concentration of 100 ug/ml, and/or lysozyme (Sigma), at a final concentration 200 pg/ml. RNase A was added to a final concentration of 50 pg/ml and the suspension was incubated at 37°C for 60 minutes with occasional mixing (by inversion). The suspension was then treated with 100 pg of proteinase K (Sigma) in the presence of 0.1-0.5% SDS at 56°C for 60 minutes. The clarified cell lysate was then extracted 3 times with equal volumes of phenol/chloroform/isoamyl alcohol (P/C/T) [25:24:1] and precipitated with 2.5 volumes of 95% ethanol. The remaining nucleic acids were pelleted by centrifugation (14,000 x g for 10 minutes) at 4°C and washed once with 1 ml of 80% ethanol. The pellet was resuspended in 100 pi of dH 2 0 and the nucleic acids were quantitated by U V absorbance at 260 nm. 2.2.3.2 Oligonucleotide primers for PCR Oligonucleotide primers for PCR (shown in Table 2.1) were synthesized based on the previously published sequences of the rnb (Zilhao et al, 1993), pnp (Regnier et al, 1987), and rhlB (Kalman etal, 1991) genes Escherichia coli. Nucleotides depicted in bold represent a 4-5 base-pair clamp which promotes the efficient cleavage of the polymerase chain reaction (PCR) products by the restriction endonucleaseBamHl. The recognition sequence for BamHl, 5-GGATCC-3' , is underlined. The Shine-Dalgarno (ribosome binding site) sequence is shown in italics. Nucleotide coordinates are shown in the 5-3' orientation of the primer. -46-Table 2.1 Oligonucleotide primers for amplification of the rnb gene by the polymerase chain reaction (PCR). Name Nucleotide Sequence Coordinates fPl rPl rP2 rP3 5'-GCGAGGATCC4 GGA GGTGAC AATTATGTTTC AGGACAAC-3' 5'-GCCGTGGATCCTTTCCTGGCTGCAAAC-3' 5'-GCGAGGATCCTTTCCATGCGGACTTCGGCATTA-3' 5'- GCGAGGATCCATCGACGGTCAGACTCATCATCA-3' 273-294 2098-2083 2182-2160 2684-2662 Table 2.2 Oligonucleotide primers for amplification of the pnp gene by PCR. Name Nucleotide Sequence Coordinates fP2 rP4 5'-GGCGCGGATCG4GG4GG7TATTACAATGCTTAATCCGATCGTT-3' 5'-CGGGGCGGATCCTTACTCGCCCTGTTCAGCAGC-3' 657-681 2799-2779 Table 2.3 Oligonucleotide primers for amplification of the rhlB gene by PCR. Name Nucleotide Sequence Coordinates fP3 rP5 5'-GGCGCGGATCG4GGvtGGrCCACACTATGAGCAAAACACAT-3' 5'-GGCGCGGATCCTTAACCTGAACGACGACGATT-3' 126-147 1398-1378 -47-2.2.3.3 Polymerase chain reaction The predicted coding sequences of the rnb (RNase II), pnp (PNPase) and rhlB (RhlB) genes of Escherichia coli were amplified from the genomic D N A of strain MV1190 by the polymerase chain reaction (PCR) in aHypercell Biologicals PTC-100 thermocycler. PCR reactions were assembled in a 100 pi volume containing 1 mM in each of the four deoxyribonucleotide triphosphates, 0.5 pg of template (gDNA), 100 pmoles each of forward primer (fP) and reverse primer (rP), and 2.5 U Taq D N A polymerase in a buffer containing 10 mM Tris-HCl, pH 9.0, 0.5-5 m M MgCl 2 , 0.01% (v/v) gelatin, 0.1% (v/v) Triton X-100 according to the manufacturer's instructions. The amplification was carried out for 30 cycles as described (Sambrook et al, 1989). Briefly, each cycle consisted of the following three steps: (1) denaturation at 95 °C for 1 minute, (2) annealing at 55 °C for 2 minutes, (3) extension at 72 °C for 4 minutes. The amplified D N A products were purified electrophoretically through horizontal 0.8% (w/v) agarose gels containing I X T A E (40 mM Tris-HCl, pH 8.0, 20 mM sodium-acetate, 1 m M E D T A ) at 75 V. The D N A was stained with 0.5 pg/ml ethidium bromide and visualized under U V light. The band containing the desired product was excised and the amplified product was purified with a Qiagen gel extraction kit according to the manufacturer's instructions. 2.2.3.4 Ligations Gel purified PCR products were digested with BamHL at 3-5 U/ug D N A in the buffer suggested by the manufacturer at 37 °C for no less than 3 hours. The cleaved D N A products were separated by electrophoresis, extracted as described in section 2.2.3.3 and were precipitated by the addition of 0.25 MNa-acetate and 2.5 volumes of ethanol at -20 °C overnight. The nucleic acids -48-were pelleted by centrifugation at 14,000 r.p.m. in a microcentrifuge, washed once with 500 pi 80% ethanol and subsequently dried for 5 minutes in a SpeedVac concentrator (Savant). Plasmid D N A (pET-11) was linearized with BarniU as above and dephosphorylated in a 50 pi reaction volume with 0.2 U of calf intestinal alkaline phosphatase in a buffer containing 10 mM Tris-HCl, pH 7.9, 10 mM NaCI, 10 mM MgCl 2 , 1 mM DTT at 37 °C for 45 minutes. Linearized, dephosphorylated plasmid D N A (60-100 ng) was added to the purified, digested target D N A and the mixture was incubated with 1 U of T4 D N A ligase in a buffer containing 30 mM Tris-HCl, pH 7.8, 10 mM MgCl 2 , 10 mM DTT, 1 mM ATP according to the manufacturer's instructions. Ligation reactions (10 pi) were incubated at 15 °C for 15-18 hours. 2.2.3.5 Transformations Competent cells were prepared by the calcium chloride method (Sambrook et al., 1989). Transformations were performed by incubating competent cells (300 pi) with the ligation reaction on ice for 60 minutes. The transformation was then heat shocked for 3 minutes at 42 °C and then chilled on ice for 5 minutes. One ml of L B media was added and the suspension was incubated for 60 minutes at 37 °C. Cells were plated on LB/agar plates containing 50-100 pg/ml ampicillin and incubated overnight at 37 °C. 2.2.3.6 A nalysis of clones Plasmid D N A was prepared by alkaline analysis lysis as previously described (Sambrook et al, 1989). Plasmids to be sequenced were purified further by precipitation with polyethylene glycol by the addition of 30 pi 20% (w/v) P E G 8000 in 2.5 M NaCI to 50 pi plasmid preparations. -49-Alternatively, plasmid DNAs for sequencing were prepared by the Promega Wizard-Plus™ plasmid isolation kit according to the manufacturer's instructions. The orientation of the fragments in the recombinant plasmids was verified by mapping with a unique restriction enzyme which cleaves within the D N A insert and Hindlll which cleaves the plasmid vector pET-11 at nucleotide 29 (~500-bp from the BamHL cloning site). The digestion products were separated electrophoretically through 0.8-1.0% (w/v) agarose gels in I X T A E and the sizes of the D N A fragments were estimated based on their mobility as compared to Hindlll digested lambda D N A markers. The plasmid inserts were subsequently sequenced by Sanger's dideoxy method in the presence of [35S]-dATP (1000 Ci/mmol) following the instructions in the Pharmacia sequencing kit. Samples were separated on thin 5-6% in acrylamide (19:1 acrylamide:bis-acrylamide), containing 8 M urea at 1600 V for 3-5 hours. The gels were fixed in a solution containing 5% ethanol and 5% acetic acid, rinsed in H 2 0 for 5 minutes and dried onto Whatman paper. The dried gels were exposed to Kodak XAR-2 film for 12-48 hours. After verifying the cloned insert, recombinant plasmids were transferred from the propagation strain MV1190 to the overexpression strain BL21(DE3) which contains a chromosomal copy of the T7 R N A polymerase as a lambda lysogen. 2.2.3.7 Oligonucleotide primers for DNA sequencing The following oligonucleotide primers for D N A sequencing were synthesized based on the previously published sequences of the rnb (Zilhao et al., 1993), and pnp (Regnier et al, 1987) genes from Escherichia coli. The coordinates of the primers are shown in the 5' to 3' orientation of the primer. -50-Table 2.4 Oligonucleotide primers for D N A sequencing of the rnb gene. Name Nucleotide Sequence Coordinates rnb-1 5 '-C AGC AAATGACTGGA-3' 1705-1691 rnb-2 5 '-TGGATTCGATTGTGG-3' 1201-1187 rnb-3 5'-CACCACGGTACAAAG-3' 764-750 rnb-4 5 '-TCC AGTC ATTTGCTG-3' 1691-1705 rnb-5 5'-CCACAATCGAATCCA-3' 1187-1201 rnb-6 5'-CTTTGTACCGTGGTG-3' 750-764 rnb-7 5 '-TCTGCCTGGATATAACG-3' 2684-2662 rnb-8 5'-AGACGCCATTCCTTG-3' 600-614 rnb-9 5'-GGCATCTATAACGTG-3' 1510-1524 Table 2.5 Oligonucleotide primers for D N A sequencing of the pnp gene. Name Nucleotide Sequence Coordinates pnp-1 5'-AAGCGAAGGCGAAAC-3' 915-929 pnp-2 5'-TACTGAACCCGACTC-3' 1148-1162 pnp-3 5 '-GC ACTGGCTGAAGCT-3' 1405-1419 pnp-4 5'-TCTGGATGTGCGTAC-3' 1641-1655 pnp-5 5'-ATGCCGGATATGGAC-3' 1906-1920 pnp-6 5'-ACGGTATCTCTGCAC-3' 2162-2176 pnp-7 5'-GCACCACCATCGAAA-3' 2414-2428 pnp-8 5'-CTTCGCTCAGCAGTT-3' 1258-1244 pnp-9 5'-GCCTGTTCCATTACG-3' 2282-2268 -51-2.2.4 Analysis of proteins 2.2.4.1 Quantitation Protein concentrations were determined by the Bradford dye-binding method (Bradford, 1976) using the Bio-Rad protein assay reagent. A range of concentrations of B S A in dH 2 0 (2-10 ug/ml) were used as the protein standard. 2.2.4.2 SDS-polyacrylamide gel electrophoresis (SDS-PAGE) Protein samples were boiled for 5 minutes in an equivalent volume of 2X SDS-sample buffer (120 mM Tris-HCl, pH 6.8, 10% glycerol, 3% sodium-dodecylsulfate (SDS), 50 mM DTT and 0.1% bromophenol blue) prior to electrophoresis through 10-13% polyacrylamide gels (36:1 acrylamide:bis-acrylamide) containing 0.1% SDS in Laemmli's running buffer (25 m M Tris, 192 mM glycine, and 0.1% SDS) (Laemmli, 1970). The proteins were visualized by staining with a Coomassie Brilliant-blue solution containing 0.05% (w/v) Coomassie Brilliant-blue R-250, 45% methanol and 10% acetic acid. Gels were destained with a solution of 5% ethanol and 5% acetic acid. 2.2.4.3 Enzyme assays Exonucleolytic activity was measured by the method of Donovan and Kushner (1983). Reactions (300 pi) contained 12 mM Tris-HCl, pH 8.0, 100 mM KC1, 1 m M MgCl 2 , and 20 nmoles of polyadenosine (150 nCi of poly[8 3H]adenylic acid, 40-140 nucleotide residues in length). Protein was added last and the reactions were incubated at 37 °C for 25 minutes. Assays for PNPase activity were supplemented with 10 mM Na-phosphate. At various times (in minutes) 50 ul aliquots were removed and quenched with an equivalent volume of yeast R N A (5mg/ml). Twenty per cent -52-trichloroacetic acid was added to a final concentration of 6.7% and the samples were left on ice for at least 30 minutes. Acid insoluble nucleic acids were precipitated by centrifugation at 14,000 x g for 10 minutes and 100 pi of supernatant was counted in 3 ml of ScintiVerse scintillation fluid (Fisher) in a Beckman LS6000 IC scintillation counter. One unit (U) is defined as the activity required to solubilize 1 pmol of A M P per hour. 2.2.5 Protein purification 2.2.5.1 Media Cell growth was carried out in one of Luria-Bertani broth (LB), Terrific Broth (TB) or M9ZB. L B medium (1% (w/v) bactotryptone, 0.5% (w/v) yeast extract, and 86 mM NaCI); M9ZB medium (M9: 18 mMNFLCl, 22 mM K H 2 P 0 4 , and 42 mM Na,P0 4; ZB: 1% (w/v) N-Z-amine A, and 86 mM NaCI); TB (TB: 1.2% (w/v) bactotryptone, 2.4% (w/v) yeast extract, 0.4% (v/v) glycerol, 17 mM K H 2 P 0 4 , 72 mM K 2 P 0 4 ) (Sambrook et al, 1989). Growth media was supplemented with 0.2%) (v/v) glucose, 1 mM M g S 0 4 and 50-100 pg/ml of ampicillin where required. 2.2.5.2 Induction Cell cultures (0.5-10) were grown at 30-37 °C with good aeration to early logarithmic phase. At an ^4^—0.5, expression of the recombinant proteins was induced by the addition of isopropyl p-D-thiogalactopyranoside (IPTG) to a final concentration of 0.4 mM and the cultures were grown for an additional 3-5 hours. -53-2.2.5.3 Harvest and cell lysis The cultures were chilled on a slurry of wet ice and the cells were harvested by centrifugation at 4,000 x g for 10 minutes. The cell pellets were resuspended once in 10 volumes of wash buffer containing 10 mM Tris-HCl, pH 8.0, 0.8 mM NH 4C1, 0.2 mM EDTA, 5% glycerol and 5 mM 0-mercaptoethanol. The washed cells were collected by centrifugation, quick frozen in an ethanol/dry ice bath and stored at -70 °C until use. Frozen cell pellets were thawed at 4 °C and resuspended in 3-5 volumes of lysis buffer. Unless otherwise stated the resuspended cells were ruptured by 3 passages through an Aminco French pressure cell at 15,000 p.s.i. 2.2.5.4 Crude fractionation of the RnbCA-38 protein Cultures of strain GC1 were grown in 1 { of M9ZB medium as described in section 2.2.5.1 Induction of the recombinant RnbCA-38 protein and cell harvest were as described in section 2.2.5.2 and 2.2.5.3. The cell pellet was resuspended in lysis buffer containing 10 mM Tris-HCl, pH 8.0, 0.8 mMNH 4 Cl , 0.2 mM EDTA, 0.1 mMDTT and 5% glycerol and the cells were ruptured as described in section 2.2.5.3. The crude cell lysate was centrifuged at 10,000 x g for 30 minutes in a Beckman JA-20 rotor to generate the soluble (S-10) and insoluble fractions (P-10). The P-10 fraction was resuspended in an equivalent volume of lysis buffer and the proteins were analyzed by SDS-PAGE as described in section 2.2.4.2. 2.2.5.5 Purification of the RnbCA-10 protein from strain GC10 Cultures of GC10 (formerly GC101 in Coburn and Mackie, 1996a) were grown in L B media as described in section 2.2.5.1. Induction of the recombinant RnbCA-10 protein and cell harvest were -54-as described in section 2.2.5.2 and 2.2.5.3. The harvested cells were made hypotonic by resuspension in 10 ml of 20 mM Tris-HCl, pH 7.5, 20% sucrose, 1 mM E D T A and lysed in ice cold dH 2 0. The resultant spheroplasts were harvested by centrifugation and resuspended in 10 ml of buffer D (20 mM Tris-HCl, pH 7.5, 5 mM EDTA, 0.1 mM PMSF, 0.8 pg/ml leupeptin and 2 pg/ml aprotinin) and were ruptured as described in section 2.2.5.3. The overexpressed RnbCA-10 protein (formerly Rnb* in Coburn and Mackie, 1996a) remained with the insoluble fraction (Fraction II) after low speed centrifugation at 4,000 x g, presumably in inclusion bodies. The latter were subjected to several washes with buffer D containing 25% sucrose, and 1% Triton X-100 to solubilize membranes and associated proteins. Washed inclusion bodies were then solubilized in 3 mL of 50 mM Tris-HCl, pH 8.0, 5 mM E D T A , 5 M guanidine-HCl, 100 mM DTT on ice for 1 hour prior to centrifugation at 12,000 x g. The supernatant containing the solubilized RnbCA-10 protein was diluted 10-fold with 50 mM Tris-HCl, pH 8.0, 0.2% Tween-20, 1 mM DTT, 20% glycerol, 0.5 mM MgCl 2 , 75 mM KCI to a final concentration of 0.1 mg/ml. RNase U activity was renatured by dialysis for 12 hours against 20 volumes of 10 mM Tris-HCl, pH 8.0, 5% glycerol, 1 mM DTT, 0.8 mM NH 4C1, 0.02 mM E D T A and concentrated to 0.5 mg/ml with an Amicon YM-10 concentrator to generate Fraction III. This fraction was subjected to chromatography (FPLC-Fast Protein Liquid Chromatography) on a Resource Q column (Amersham-Pharmacia) equilibrated with 3-5 column volumes of buffer E (10 mM Tris-HCl pH 8.0, 0.8 mM NH 4C1, 0.2 mM EDTA, 1 mM DTT, 150 mM NaCI). The renatured RnbCA-10 protein was eluted from the column with a 50 mL gradient of NaCI (150 mM-350 mM) over a large concentration range of NaCI (200 mM-350 mM). The presence of the RnbCA-10 protein in various fractions was measured qualitatively by SDS-PAGE and quantitatively by enzyme assay as described in section 2.2.4. Fractions containing the active exonuclease activity were pooled to -55-generate Fraction IV and stored at -70°C. 2.2.5.6 Purification of recombinant RNase II from strain GC100 Cultures of GC100 were grown in L B media as described in section 2.2.5.1. Induction of the recombinant Rnb protein and cell harvest were as described in section 2.2.5.2 and 2.2.5.3. Cell pellets were resuspended in 20-25 ml of buffer B containing 50 mM HEPES-NaOH, pH 7.5, 500 mM NaCl, 1 mM M g C l 2 , 0.1 mM EDTA, 5 mM DTT, 0.1 mM PMSF, 0.8 ug/ml leupeptin and 2 ug/ml aprotinin. The cells were disrupted as described in section 2.2.5.3 and the lysate was centrifuged at 30,000 x g for 60 minutes in a Beckman JA-20 rotor to pellet unbroken cells and insoluble material (Fraction I). Approximately 60 mg of this fraction was loaded onto a column of Affi-Gel blue (Biorad) (1.25 cm x 21.5 cm) previously equilibrated with 3 column volumes of buffer C (25 mM HEPES-NaOH, pH 7.5, 5% glycerol, 2 mM DTT , 1 m M M g C l 2 , 0.1 m M EDTA) containing 500 mM NaCl. The column was washed with 3-5 column volumes of this buffer at a flow rate of 8.3 ml/hr (6.75 cm/hr) driven by a PI peristaltic pump (Amersham-Pharmacia). The Rnb protein was eluted with 5 column volumes of buffer C containing 3 M NaCl to generate Fraction II. The eluant was pumped directly onto a column of hydroxylapatite (Bio-Rad) (0.75 cm x 8.5 cm) at a flow rate of 6.7 ml/hr (15 cm/hr). After washing with 5 column volumes of buffer C containing 1 mM Na-phosphate, pH 7.5, the Rnb protein was eluted with a 50 ml gradient of Na-phosphate, pH 7.5, (1 mM-250 mM) in buffer C at a concentration of 75 mM Na-phosphate. Fractions containing the Rnb protein were divided into pool A (Fraction III) or pool B based on the contaminants present in the fractions. A portion of pool A (Fraction III), was loaded onto a column of Affi-Gel heparin (Bio-Rad) (0.75 cm x 8.0 cm). The column was washed with 3-5 column volumes of buffer C at a flow rate of 7.2 ml/hr -56-(16 cm/hr). The Rnb protein was eluted from the column with a 50 ml gradient of NaCI (0 mM-400 mM) in buffer C at a concentration of 130-140 mM NaCI (Fraction V). Alternatively, chromatography (FPLC) of pool A on a Resource Q column (Amersham-Pharmacia) was substituted for the Affi-Gel heparin step. After loading the sample and washing it with 5 column volumes of buffer C containing 150 mM NaCI, the Rnb protein was eluted from this resin with a 50 ml gradient of NaCI (100 mM-400 mM) in buffer C at a concentration of 220 mM NaCI (Fraction IV). The presence of the 72.5 kDa protein in various fractions was monitored qualitatively by SDS-PAGE and quantitatively by enzyme assay as described in section 2.2.4. 2.2.5.7 Overexpression and purification of PNPase from strain GC400 Cultures of GC400 were grown in 500 ml of L B media as described in section 2.2.5.1. Induction of the recombinant Pnp protein and cell harvest were as described in section 2.2.5.2 and 2.2.5.3. Cell pellets were resuspended in 5 ml of lysis buffer (50 mM HEPES-NaOH, pH 7.5, 0.1 mM EDTA, 2 mM DTT and 0.1 mM phenylmethylsulfonyl fluoride). The cells were ruptured as described in section 2.2.5.3 and the cell lysate was centrifuged to pellet unbroken cells and insoluble material (Fraction I) as described in section 2.2.5.6. Approximately 20 mg of the S-30 (Fraction I) was loaded onto a column of Affi-Gel heparin (0.75 x 10 cm) (Bio-Rad) which had been previously been equilibrated with 3-5 column volumes of Buffer B (25 mM HEPES-NaOH, pH 7.5, 5% glycerol, 1 mM DTT, and 0.1 mM EDTA). After washing with 5 column volumes of Buffer B at a flow rate of 9 ml/hr, the bound proteins were eluted with a 30 ml gradient of NaCI (0 M - l M). The protein content of each fraction was monitored by SDS-polyacrylamide gel electrophoresis. Fractions containing the Pnp protein (based on the size of the a-subunit, approximately 85 kDa) which eluted -57-at a concentration of 160 mMNaCl were pooled (Fraction II) and subjected to chromatography on a column of Affi-Gel blue (1.0 x 10 cm) (Bio-Rad). The column was washed with 5 column volumes of Buffer B containing 1 M N a C l at a flow rate of 9 ml/h and the Pnp protein was eluted with a single step of Buffer B containing 3 M KC1. Fractions containing Pnp were concentrated with an Ultrafree centrifugal filter device (Millipore). Approximately 2.5 mg of the blue-agarose pool (Fraction III) was then subjected to gel filtration on a Bio Gel A 0.5 M column (1 x 48 cm) (Bio-Rad) previously equilibrated with 3-5 column volumes of Buffer B containing 100 mM NaCl. Fractions containing the Pnp protein, which eluted with a volume of 17-27 ml, were pooled and concentrated with an Ultrafree centrifugal filter device (Millipore) to generate the final fraction, Fraction IV. 2.2.5.8 Purification of degradosomes from strain CF881 Cultures of CF881 were grown in 1L of Terrific Broth at 30°C to an A600 of 22.3 in a fermenter. Cells (30 g) were harvested as described in section 2.2.5.3, washed twice in buffer containing 10 mM Tris-HCl, pH 7.5, 10 mM Mg-acetate, and 10 mM KC1 as described previously (Misra and Apirion, 1979) and stored frozen at -70 °C until use. Purified degradosomes were prepared as described (Carpousis et al., 1994) with some modification. Cell pellets were thawed at 4 °C and resuspended in 30 ml (1 ml/g of cells) of lysozyme-EDTA buffer (50 mM Tris-HCl, pH 7.5, 100 mM NaCl, 5% glycerol, 3 mMEDTA, 1 mMDTT, 1.5 mg/ml lysozyme, 1 m M P M S F , 2 ug/ml aprotinin, 0.8 ug/ml leupeptin, 0.8 ug/ml pepstatin A) at room temperature. The suspension was subjected to the following alternating rounds of mixing and incubation on ice: stirring for 3 minutes, ice for 10 minutes, stirring for 1 minute, ice for 20 minutes, stirring for 1 minute, ice for an additional 40 minutes. After the 40 minute incubation, 15 ml (0.5 ml/g of cells) of DNase I-Triton buffer (50 -58-mM Tris-HCl, pH 7.5, 100 mM NaCI, 5% glycerol, 1 mM DTT, 30 mM Mg-acetate, 3% Triton X -100, 20 pg/ml DNase I, 1 mM PMSF, 2 pg/ml aprotinin, 0.8 pg/ml leupeptin, 0.8 pg/ml pepstatin A) at room temperature was added and the cell suspension was mixed for 1 minute and then placed on ice for 30 minutes. Ammonium-chloride (5 M) was added slowly with stirring to a final concentration of 1 M and the lysate was incubated with stirring on ice for an additional 30 minutes. The cell lysate was centrifuged at 30,000 x g in a Beckman JA-20 rotor for 60 minutes to remove unbroken cells and debris to generate an S-30 fraction. The S-30 (Fraction I) was then subjected to second centrifugation at 200,000 x g for 2 hours with a Beckman Ti75 ultracentrifuge rotor. The S-200 (Fraction II) (60 ml) was fractionated by the addition of 26% (w/v) (NH 4 ) 2 S 0 4 with stirring at 4 °C. The precipitated proteins were collected by centrifugation at 10,000 x g for 60 minutes in a Beckman JA-20 rotor. Interestingly, due to the presence of detergent some of the protein pellet floated on top of the supernatant. The ammonium-sulfate pellet and the floating pellet were resuspended in 40 ml of buffer A (10 mM Tris-HCl, pH 7.5, 5% glycerol, 0.5% Genapol X-080, 1 mMEDTA, 0.1 mMDTT, 0.1 m M P M S F , l mM PMSF, 2 pg/ml aprotinin, 0.8 pg/ml leupeptin, 0.8 pg/ml pepstatin A) containing 50 mMNaCl to generate Fraction m. This material was loaded directly onto a column of SP-sepharose FF (1.0 x 20 cm) (Amersham-Pharmacia) which had been previously equilibrated with 3 column volumes of buffer A. After the column had been washed with 3 volumes of buffer A and 5 volumes of buffer A containing 300 mM NaCI at a flow rate of 20 ml/h, the bound proteins were were eluted from the column in a single step with 50 ml of buffer A containing 1% Genapol X-080 and 1 M NaCI. Fractions containing the degradosome complex were pooled (Fraction IV). This pooled fraction was subjected to chromatography on a column of Affi-Gel blue (Bio-Rad) which was previously equilibrated with 5 column volumes of buffer A containing 50 m M KCI. After -59-the column had been washed with 5 volumes of buffer A containing 50 mM KC1 at a flow rate of 9 ml/h, the bound proteins were eluted with a 30 ml gradient of KC1 (0.3 M - 3 M). Fractions containing the degradosome were pooled (Fraction V) and precipitated with 26% (w/v) ammonium-sulfate as before. The ammonium-sulfate pellet was resuspended in buffer A containing 50 mM NaCl and subjected to gel filtration on a Bio Gel A 1.5 M column (1.5 x 70 cm) (Bio-Rad) which had been equilibrated with buffer A containing 50 mM NaCl. Fractions containing the degradosome complex, which eluted with a volume of43-56 ml, were pooled and concentrated using an Ultrafree centrifugal device (Millipore) to generate the final fraction, Fraction V. The presence of the degradosome in various fractions was monitored qualitatively by SDS-PAGE gels using the distinctive size of the Rne protein as a visual marker. 2.2.5.9 Purification of the recombinant RhlB protein from strain GC300 Cultures of GC300 were grown in 500 ml of L B media as described in section 2.2.5.1. Induction of the recombinant RhlB protein and cell harvest were as described in section 2.2.5.2 and 2.2.5.3. Cells were resuspended in 5 ml of buffer A (50 mM Tris-HCl, pH 8.5, 100 m M NaCl, 0.1 mMEDTA, 2 mMDTT). The cells were ruptured as described in section 2.2.5.3 and the cell lysate was clarified as described in section 2.2.5.6 to generate the S-30 fraction (Fraction I). Since the overexpressed RhlB protein was insoluble at pH 7.5, presumably due to isoelectric precipitation (calculated pi =7.5), purification of the recombinant protein was performed at pH>8.5. Approximately 60 mg of Fraction I was loaded onto a column of Affi-Blue (0.75 x 10 cm) (Bio-Rad) which had been previously equilibrated with 3-5 column volumes of Buffer B (25 m M Tris-HCl, pH 8.5, 5% glycerol, 0.1 mMEDTA, 1 mMDTT) containing 100 mM NaCl. After the column had been -60-washed with 5 column volumes of Buffer B at a flow rate of 20 cm/h, the RhlB protein was eluted from the column with a 30 ml gradient of KCI (0-3 M) at a concentration of 1.7 M KCI. Fractions containing the RhlB protein were pooled and concentrated in an Ultra-free centrifugal filtration device (Millipore) to generate Fraction U. A portion of this pool was subjected to gel filtration on a column of Bio Gel A 0.5 M (1 x 48 cm) (Bio-Rad) previously equilibrated with 3-5 volumes of Buffer B containing 50 mMNaCl. Fractions containing the RhlB protein were pooled and concentrated in an Ultra-free centrifugal device (Millipore) to generate Fraction III. A significant fraction of Fraction III was loaded onto a column of Affi-Gel heparin (0.75 x 10 cm) which had been previously equilibrated with 3-5 column volumes of buffer B containing 50 mM NaCI. After the column had been washed with 5 volumes of starting buffer at a flow rate of 20 cm/h, the RhlB protein was eluted with a 30 ml gradient (0 M - l M) NaCI at a concentration of 500 mM NaCI. Fractions containing the RhlB were pooled, dialyzed against 1 L of Buffer B containing 50 m M N a C l and concentrated as before to generate Fraction IV. This fraction was loaded onto a 1 ml Resource Q column (Amersham-Pharmacia) which had been previously equilibrated with several volumes of Buffer B containing 50 mMNaCl. After the column had been washed with 5 volumes of starting buffer at 1 ml/min, the RhlB protein was eluted with a 30 ml gradient of NaCI (50 mM-500 mM) at a concentration of 120 mM-150 mMNaCl. Fractions containing the RhlB protein (Fraction V) were pooled and stored at -70°C. The presence of RhlB in various fractions was monitored qualitatively by SDS-PAGE gels as described in section 2.2.4. -61-2.3 Results 2.3.1 Construction of a strain overexpressing the rnb gene Several predicted coding sequences spanning nucleotides 280-2214 of the rnb gene of Escherichia coli (X67913; Zilhao etal, 1993; Figure 1, panel A) were amplified by the polymerase chain reaction (PCR) from the genomic D N A of strain MV1190 as described in section 2.2.3.3. The amplified products were cleaved with BamHl and ligated into the unique BamHl site of pET-11 to generate plasmids p G C l , from primers fPl - rPl (see Table 2.1) (Figure 1, panel B), pGCIO from primers fPl-rP2 (Figure 1, panel C) and pGClOO from primers fPl-rP3 (Figure 1, panel D). The forward primer, fPl , contains a Shine-Dalgarno sequence 5' to the rnb start codon (italicized in Table 2.1) such that the amplified rnb gene could be subsequently overexpressed using the T7 R N A polymerase encoded by BL21(DE3) (Studier et al, 1990). The orientation of the 1.9-kbp BamHl fragments in the recombinant plasmids, pGCl and pGCIO, was verified by restriction mapping with Nsil and Hindlll. The D N A sequence of the rnb gene, in all three recombinant plasmids was confirmed by D N A sequencing as described in section 2.2.3.6. The resulting plasmids, p G C l , pGCIO and pGClOO, were used to transform BL21(DE3) (Studier et al, 1990) to generate strains GC1, GC10, and GC100, respectively. The partial structures of plasmids containing all or part of the rnb gene are depicted in Figure 1. Plasmid p G C l (Figure 1, panel D) contains all of the previously published coding sequence (280-2078) of the rnb gene (Zilhao et al, 1993). However, sequencing of this construct revealed two errors in the published rnb sequence: an additional G residue at position 2002 and an additional C residue at position 2039. Inclusion of these residues resulted in an extension of the open reading frame past the predicted U G A stop codon at position 2078 (Zilhao et al, 1993) repositioning it to -62-Figure 2.1 Strategy for cloning and overexpression of rnb. Panel A illustrates a linear representation of the rnb gene of E. coli and its flanking sequences. Plasmids pGClOO (panel B), pGCIO (panel C) and pGCl (panel D) were constructed by PCR as described in section 2.2.3.2. The solid black boxes represents the cloned sequence and the line represents sequences derived from the vector pET-11. Coordinates of the rnb sequence and the predicted start and stop codons are shown in panel A. The Shine-Dalgarno sequence is represented by an enclosed "SD". Restriction sites are denoted as follows: B (BamYS), N (NsiT), H (HindUT). The open box represents regions 3' to the rnb gene also cloned into plasmid pGClOO (panel B). Plasmid pGCIO (panel C) and plasmid p G C l (panel D) lack 33 and 117 nucleotide residues respectively from the 3' end of the rnb coding sequence. This deletion has extended the open reading frames 66 nucleotide residues into the T7 terminator region of pET-11 shown by the shaded boxes. The "TGA" stop codon at position 2074 (actually position 2078), predicted by the original rnb sequence (Zilhao etal, 1993), is not in frame in the corrected sequence. In panel A, the D N A nucleotide sequence is numbered by the following convention: the start and stop codons are indicated according to the 5' position of the nucleotide triplet, whereas restriction sites are denoted by the position of the nucleotide 5' to the cleavage site. -63-ATG 280 442 A/s/l rnb "TGA" 2078 _ J TAA 2214 I 2062 Dra III 2442 A/s/l T7 pronv/acO B ATG ATG T7 prom-/acO N N N I pGC100 pGC10 pGC1 N B TAA i H i B TAG T7-fe/m H • T7-temi B TAG H I T7-term -64-position 2123 (UGA). As a result of this error, it was necessary to repeat the entire amplification and subsequent ligation into pET-11 using primer fPl and the reverse primer rP2 (see Figure 1 and Table 2.1). Sequencing of the resultant plasmid, pGCIO (Figure 1, panel C), revealed the omission of two additional A residues at positions 2112 and 2117. Addition of the four new residues to the rnb sequence (Zilhao etal, 1993) extended the reading frame, passing all sequences appearing in the data base. We, therefore, obtained two plasmids, pRZA17 and pRZA18 from Dr. Cecilia Arraiano which contained sequences distal to the 3'-end of the published portion of the rnb gene (Zilhao et al, 1993). Once an authentic rnb sequence was obtained from these two plasmids, it was possible to repeat the amplification of the chromosomal rnb gene using primers fPl and the reverse primer rP3. The amplified rnb fragment generated from fP 1 and the reverse primer rP3 was cleaved with BamHL and ligated into pET-11 to generate pGClOO (Figure 1, panel B). This plasmid contains the entire predicted 1935 nucleotide residue open reading frame, the 3' untranslated region including the putative Rho-independent terminator, and approximately 400 nucleotide residues of intercistronic spacer under the control of the 11-lac promoter-operator region in pET-11 (Studier et al, 1990). D N A sequencing of this construct revealed a C - G transition, resulting in a single amino acid change Ser572 - Gly 5 7 2 , that had inadvertently been incorporated into pGClOO. Otherwise, the entire sequence agreed with the revised D N A sequence of the rnb gene (Zilhao et al, 1993) and the sequence obtained from the E. coli genome (Blattner et al, 1997). 2.3.2 Purification of recombinant RNase II After induction of GC100 with TPTG, the Rnb protein (72.5 kDa) was expressed to the extent that it represented the most abundant protein in whole cell extracts and a significant fraction of the -65-total cellular protein (Figure 2, compare lanes 2 and 3). When assayed against poly(A), crude extracts (S-30) from strain GC100 displayed a specific activity of 1,184 U/mg (see Table 2.6), 300-fold higher than that obtained from crude extracts prepared from the haploid strain CF881 (specific activity= 3.9 U/mg). An efficient method of purification was developed in part by exploiting several effective steps from previously published methods (Spahr, 1964; Singer and Tolbert, 1965; Gupta et al, 1977; Cudny and Deutscher, 1980). The initial step relies on chromatography on Cibacron blue-agarose to remove the bulk of the nucleic acids and contaminating proteins while the majority (>90%) of the Rnb protein remains bound to the column (Figure 2, lane 5). Considerable efficiency was gained by loading the 3 M N a C l eluate from this column directly onto a hydroxylapatite column. This proved to be an invaluable step in the purification method since concentration, desalting and significant purification of the Rnb protein could take place in a single step (Figure 2, lane 6). The apparent loss of activity after hydroxylapatite chromatography (Table 2.6) may have been due to the inhibition by Ca 2 + ions leached from the column at high ionic strength as Ca 2 + has been reported to inhibit RNase II activity (Spahr, 1964; Singer and Tolbert, 1965; Gupta et al, 1977). Final purification of Rnb from most contaminants could be achieved by affinity chromatography on heparin-agarose or by ion exchange chromatography (FPLC) (see section 2.2.5.6). A sample of the purified recombinant Rnb protein is shown in Figure 2, lanes 7 and 8. Based upon Coomassie-blue or silver staining of overloaded polyacrylamide gels, the preparation was judged to be about 95% pure with a few faint minor contaminating bands. The specific activity of the Rnb protein purified to the end of the heparin -66-M C E I II III IV V Figure 2.2 Purity of the recombinant RNase II. Cultures of GC100 were induced with IPTG and grown for 5 hours at 30°C prior to harvest, lysis and purification as described in section 2.2.5. The following samples were denatured, separated through a 13% SDS-polyacrylamide gel and stained with Coomassie Brilliant-blue: lane 1, molecular mass standards (Bio-Rad); lane 2, boiled cell extract from a noninduced culture of GC100; lane 3, boiled cell extract from an induced culture of GC100; lane 4, S-30 extract (10 pg) (Fraction I); lane 5, pooled fractions obtained from chromatography of the S-30 fraction on blue-agarose (Fraction II) (2.5 pg); lane 6, pooled fractions from chromatography of Fraction II on hydroxylapatite (Fraction III) (1.75 pg); lane 7, pooled fractions from chromatography of Fraction III on Resource Q (Fraction IV) (1.5 pg); lane 8, pooled fractions from chromatography Fraction IV on heparin-agarose (Fraction V) (1.5 pg). -67-Table 2.6 Purification of recombinant RNase II. Fraction Protein Units Specific Activity mg fxmole AMP/hr units/mg S-30 60 71040 1184 Blue Agarose1 19.4 57317 2955 Hydroxylapatite A 6.3 16265 2603 Hydroxylapatite B 6.1 10290 1695 Heparin Agarose2 2 8195 4098 Resource Q 3 0.7 2533 3725 1. Based on loading 60 mg of the S-30 fraction onto the blue agarose column. 2. Based on loading 2.5 mg of the hydroxylapatite (pool A) fraction onto the heparin agarose column. 3. Based on loading 0.8 mg of the hydroxylapatite (pool A) fraction onto the Resource Q column. -68-agarose step was determined to be 4,100 U/mg which is nearly 2-fold greater than that reported by others for the enzyme purified from a haploid strain (Singer and Tolbert, 1965; Gupta et al, 1977; Ghosh and Deutscher, 1978). However, the specific activity of this preparation is approximately 2.5-fold lower than the best reported purification (Cudny and Deutscher, 1980, 1994). It is quite possible that not all of the overexpressed Rnb protein is properly folded or fully active. Alternatively, substitution of a serine for a glycine residue at posisiton 572 near the SI RNA-binding domain may also explain the reduction in activity. Nonetheless, this method provides a rapid and facile purification of RNase II with good yields and activity. If recoveries from the heparin-agarose chromatography step in Table 2.6 are extrapolated to include all the material from the hydroxylapatite column, the overall yield is 29%. This apparent overall yield is low for three reasons. First, the activity in crude extracts represents the sum of activities of a number of endo and exonucleases and overstates the activity of RNase II. Second, particularly after hydroxylapatite chromatography, fractions were pooled to maximize purity rather than yield. Third, RNase II is readily inactivated at 37 °C and may be sensitive to oxidation during purification (it contains 10 cysteines). 2.3.3 Purification of C-terminally truncated derivatives of RNase II 2.3.3. J The RnbCA-10 protein After induction of GC10, a recombinant Rnb protein lacking 10 C-terminal amino acid residues (RnbCA-10), was expressed to high levels (Figure 3B, compare lanes 2 and 3). Due to errors in the rnb sequence, approximately 22 amino acid residues derived from the vector pET-11, were added to the C-terminus of the truncated protein (Figure 1, panel C). The RnbCA-10 protein -69-remained in the insoluble fraction after a low speed centrifugation (Figure 3B, compare lanes 4 and 5) of the crude cell lysate and was presumed to form inclusion bodies after induction of cultures of GC10 with IPTG. The inclusion bodies were subsequently purified to near homogeneity by differential centrifugation in the presence of detergent. Authentic RNase II activity was recovered following solubilization, reduction and refolding of the truncated RnbCA-10 protein (see section 2.2.5.5). Further purification of the renatured truncated RnbCA-10 protein from most contaminants could be achieved by ion exchange chromatography (FPLC) (Figure 3B, lane 7). The truncated RnbCA-10 protein eluted from the Resource Q column over a broad range of NaCl concentrations likely reflecting the presence of several different populations of misfolded and inactive polypeptides in the preparation. RNase II activity eluted from the column as a sharp peak at a NaCl concentration of220 mM. Although a significant amount of activity could be recovered from the inclusion bodies, the specific activity of this preparation was quite poor, 54 mU/mg, a small fraction of that obtained for the full-length Rnb protein (compare to Table 2.6). 2.3.3.2 The RnbCA-38 protein After induction of GC1 with IPTG a recombinant Rnb protein, lacking 38 C-terminal amino acid residues (RnbCA-38), was expressed to high levels (Figure 3A, lane 2). As in the case of RnbCA-10 overexpression, 22 amino acid residues derived from the vector pET-11 were added to the C-terminus of the truncated RnbCA-38 protein (Figure 1, panel D). In contrast to the RnbCA-10 protein, which was insoluble, the RnbCA-38 protein remained in the soluble fraction after a rigorous centrifugation (150,000 x g for 2 hrs). The RnbCA-38 protein could be partially purified by chromatography on DEAE-sepharose and hydroxylapatite as well as by gel filtration but -70-Figure 2.3 Overexpression of the recombinant RnbCA-38 protein and purity of the RnbCA-10 protein. Cultures of GC1 (panel A) or GC10 (panel B) were induced with IPTG and grown for 5 hours at 30°C prior to harvest, lysis and purification as described in section 2.2.5. In panel A, the following samples were denatured, separated through a 10% SDS-polyacrylamide gel and stained with Coomassie Brilliant-blue: lane 1, molecular mass standards (Bio-Rad); lane 2, S-10 extract (Fraction I); lane 3, P-10 resuspended (Fraction II). In panel B, the following samples were denatured, separated through a 13% SDS-polyacrylamide gel and stained with Coomassie Brilliant-blue: lane 1, molecular mass standards (Bio-Rad); lane 2, boiled cell extract from a noninduced culture of GC10; lane 3, boiled cell extract from an induced culture of GC10; lane 4, S-30 extract (10 pg) (Fraction I); lane 5, P-30 resuspended (1.5 pg) (Fraction II); lane 6, detergent washed, solubilized and renatured inclusion bodies (Fraction III) (2 pg); lane 7, pooled fractions from chromatography of Fraction III on Resource Q (Fraction IV) (1.5 pg). -71-was not studied further since it lacked any detectable RNase II activity (see section 3.3.3). 2.3.4 Construction of a strain overexpressing PNPase The predicted coding sequence (nucleotides 664-2799) of the pnp gene from Escherichia coli (J02638; Regnier et al, 1987) was amplified from the genomic D N A of strain MV1190 by the polymerase chain reaction (Figure 4, panel A) as described in section 2.2.3.3. The amplified products were cleaved with ita/wFfl and ligated into the unique BamHl site of pET-11 to generate the plasmid pGC400 (Figure 4, panel B). The forward primer, fP2, contains a Shine-Dalgarno sequence (italicized in Table 2.2) 5' to the pnp start codon such that the amplified product could be subsequently overexpressed using the T7 R N A polymerase encoded by BL21(DE3) (Studier et al, 1990). In addition, primer fP2 was also designed to mutate the wild-type U U G start codon of the pnp gene to a more efficient A U G codon. The orientation of the 2.1-kbp (fP2-rP4) fragment in the recombinant plasmid was verified by restriction mapping of the resulting plasmid, pGC400, with Smal and Hindlll and by D N A sequencing as described in section 2.2.3.6. Sequence analysis of the recombinant pnp gene revealed a O G transition in pGC400, resulting in a single amino acid change, A r g ^ - G l y 3 5 7 in the recombinant PNPase. This plasmid was used to transform BL21(DE3) (Studier et al, 1990) to yield strain GC400 as described above. No other deviations from the published D N A sequence of the pnp gene (Regnier etal, 1987) were detected. The partial structure of the plasmid containing the pnp gene is depicted in Figure 4B. 2.3.5 Purification of recombinant PNPase After induction of GC400 with IPTG, the recombinant Pnp protein (predicted molecular -72-A TTG TAA 664 pnp 2799 II— I 885 Sma I B B S B TAA H ATG pGC400 1 SD T7 prom-/acO Figure 2.4 Strategy for cloning and overexpression of pnp. Panel A illustrates a linear representation of the pnp gene ofE. coli and its flanking sequences. Plasmid pGC400 (panel B) was constructed by PCR as described in section 2.2.3.3. The solid black box represents the cloned sequence and the line represents sequences derived from the vector pET-11. Coordinates of the pnp sequence and the predicted start and stop codons are shown in panel A. The Shine-Dalgarno sequence is represented by an enclosed SD. Restriction sites are denoted as follows: B (BamHl), S (SmaT), Ff (Hindlll). In panel A, the DNA nucleotide sequence is numbered by the following convention: the start and stop codons are indicated according to the 5' position of the nucleotide triplet, whereas restriction sites are denoted by the position of the nucleotide 5' to the cleavage site. -73-kDa M I II HI IV 200 116 97 66 45 31 85 kDa 1 2 3 4 5 Figure 2.5 Purity of recombinant PNPase. Cell growth, harvest, lysis and enzyme purification are described in section 2.2.5. The following samples were denatured, separated through a 10% SDS-polyacrylamide gel and stained with Coomassie Brilliant-blue: lane 1, molecular mass standards (Bio-Rad); lane 2, S-30 (Fraction I) (6.5 pg); lane 3, pooled fractions obtained from chromatography of the S-30 on heparin-agarose (Fraction II) (2.5 pg); lane 4, pooled fractions from chromatography of Fraction II on blue-agarose (Fraction III) (1.5 pg); lane 5, pooled fractions from chromatography of Fraction III fraction by gel filtration (Fraction IV) (1.5 pg). -74-weight of the a subunit= 77 kDa; 85 kDa by SDS-PAGE) was expressed to the extent that it represented the most abundant protein in whole cell extracts and a significant fraction of the total cellular protein (Figure 5, lane 2). To compare PNPase activity during purification to that of RNase II, a standard poly(A) assay for RNase II activity supplemented with 10 mM sodium-phosphate was utilized (Donovan and Kushner, 1983). When assayed against poly(A), crude extracts (Fraction I) from strain GC400 displayed a specific activity of 6.5 U/mg (see Table 2.6), 200-fold higher than that obtained from crude extracts prepared from the haploid strain 18-11 (RNase II") (specific activity= 0.035 U/mg). In both cases, phosphate-independent activity was subtracted from the phosphate-dependent activity. As in the case for recombinant RNase II, an efficient method of purification was developed in part by exploiting several effective steps from previously published methods (Portier et al, 1973; Soreq and Littauer, 1977; Drocourt etal, 1978). Details are given in section 2.2.5.7. The initial step relied on heparin-agarose chromatography to remove the bulk of the nucleic acids and contaminating proteins (Figure 5, compare lanes 2 and 3). Fractions containing the recombinant Pnp protein were pooled and subjected to chromatography on Cibacron blue-agarose which removed most of the remaining contaminating proteins (Fraction III, Figure 5, lane 4). Finally, since PNPase is known to exist as an oc3 trimer (Portier et al, 1973; Portier, 1975) with a predicted molecular weight of 231 kDa, gel filtration proved to be an excellent step since desalting and significant purification could be accomplished in a single step. Moreover, the gel filtration step was crucial for the removal of contaminating RNase II activity. The final fraction, Fraction IV (gel filtration), was judged to be greater than 95% homogeneous as determined visually on an overloaded Coomassie-blue stained SDS-polyacrylamide gel (Figure 5, lane 5). Based on its activity, the final specific activity of Fraction -75-IV was calculated to be 40 U/mg, a 6-fold enrichment over crude extracts from the overexpressing strain and a 1200-fold enrichment over extracts from the haploid strain. If recoveries from the gel filtration chromatography step in Table 2.7 are extrapolated to include all the material from the blue-agarose column, the overall yield is approximately 50 %. Since the binding capacity of the heparin-agarose support is relatively low (2 mg/ml of BSA), a large proportion of PNPase in the S-30 fraction flowed through the column and was not examined further. Therefore, this calculation assumes that only a maximum of 10 mg of the recombinant PNPase protein was retained on the column. As before, fractions from each chromatography step were pooled to maximize purity rather than activity. Table 2.7 Purification of recombinant PNPase. Fraction Protein Units Specific Activity mg_ fimole AMP/hr units/mg S-30 (I) 108 702 6.5 Heparin Agarose1 (II) <10 nd nd Blue Agarose (III) 5.7 nd nd Gel Filtration2 (IV) 0.4 16 40 1. Although all of the S-30 (Fraction I) was loaded onto the heparin agarose column, only 5-10 mg of protein bound to the column. 2. Based on loading of 2.8 mg of the blue agarose fraction (Fraction III). -76-2.3.6 Construction of a strain overexpressing rhlB The predicted coding sequence (nucleotides 233-1378) of the rhlB (mmrA in Blattner et al, 1997) gene from Escherichia coli (X56310; Kalman et al, 1991) was amplified from the genomic D N A of strain MV1190 by the polymerase chain reaction as described in section 2.2.3.3 (Figure 7, panel A). The amplified products were cleaved with the restriction enzyme BamHl and ligated into the unique BamHl site of pET-11 to generate plasmid pGC300. The forward primer, fP3, contains a Shine-Dalgarno sequence 5' to the rhlB start codon (italicized in Table 2.3) such that the rhlB gene could be subsequently overexpressed using the T7 RNA polymerase encoded by BL21(DE3) (Studier et al, 1990). The structure of pGC300, containing the amplified rhlB gene, was verified by restriction mapping. The presence and size of the rhlB insert was confirmed by excision of 1.3-kbp fragment from pGC300 by BamHl (data not shown). The correct orientation of the rhlB fragment, with respect to the T7 promoter, was confirmed by digestion with Kpnl and Hindlll which generates the expected 1.6-kbp fragment (data not shown). D N A sequence analysis was not undertaken. The partial structure of plasmid pGC300 containing the rhlB gene is depicted in Figure 6B. 2.3.7 Purification of recombinant RhlB After induction of cultures of GC300 with IPTG, the RhlB protein (50 kDa) was expressed to relatively high levels. An effective four step purification, as described in section 2.2.5.7, was developed by exploiting the putative R N A and nucleotide binding properties of the enzyme (Figure 7, lanes 3-6). Chromatography of the S-30 fraction (Fraction I) on blue-agarose removed the majority of the contaminating nucleic acids and proteins. Fractions were pooled based on purity as ascertained by Coomassie Blue-stained SDS-polyacrylamide gels. The pooled blue-agarose fraction, (Fraction -77-Figure 2.6 Strategy for cloning and overexpression of rhlB. Panel A illustrates a linear representation of the rhlB gene of E. coli and its flanking sequences. Plasmid pGC300 (panel B) was constructed by PCR as described in section 2.2.3.3. The solid black box represents the cloned sequence and the line represents sequences derived from the vector pET-11. Coordinates of the rhlB sequence and the predicted start and stop codons are shown in panel A. The Shine-Dalgarno sequence is represented by an enclosed SD. Restriction sites are denoted as follows: B (BamHl), K (KpnT), H (HindllT). In panel A, the D N A nucleotide sequence is numbered by the following convention: the start and stop codons are indicated according to the 5' position of the nucleotide triplet, whereas restriction sites are denoted by the position of the nucleotide 5' to the cleavage site. -78-M I II III IV V kDa ._ 1 2 3 4 5 6 Figure 2.7 Purity of recombinant RhlB. Cell growth, harvest, lysis and enzyme purification are described in section 2.2.5. The following samples were denatured, separated through a 10% SDS-polyacrylamide gel and stained with Coomassie Brilliant-blue: lane 1, molecular mass standards (Bio-Rad); lane 2, S-30 (Fraction I) (7 pg); lane 3, pooled fractions obtained from chromatography of the S-30 on blue-agarose (Fraction II) (3 pg); lane 4, pooled fractions from chromatography of Fraction II by gel filtration (Fraction III) (2 pg); lane 5, pooled fractions from chromatography of Fraction III fraction on heparin-agarose (Fraction IV) (1.75 pg); lane 6, pooled fractions from chromatography of Fraction IV on Resource Q (Fraction V) (1.75 pg). -79-II) , was subjected to gel filtration chromatography which removed some contaminating proteins (above 50 kDa) but mainly served to desalt Fraction II. The pooled gel filtration fraction (Fraction III) was then subjected to chromatography on heparin-agarose which removed the main contaminating protein which migrated as a 48 kDa band on SDS-polyacrylamide gels. Finally, the pooled heparin-agarose fraction (Fraction IV) was then subjected to FPLC-chromatography on a Resource Q column. The final fraction, Fraction V, is about 95% homogeneous, with only some minor contaminating bands, as judged by Coomassie-blue-stained SDS-polyacrylamide gels. Since we have not assayed ATPase activity or R N A unwinding activity, it is not clear whether, or not, the recombinant RhlB protein is active. 2.3.8 Purification of the degradosome The degradosome was originally purified by a combination of ammonium sulfate fractionation, ion-exchange chromatography on SP-sepharose and finally by preparative glycerol gradient centrifugation (Py etal, 1996; Carpousis etal, 1994). We have applied these authors' methods with some modifications as described in section 2.2.5.8. During chromatography of the ammonium-sulfate fraction (Fraction III) on SP-sepharose, several contaminating proteins continued to elute from the column throughout the 0.3 M NaCl wash step and were carried over into the final 1 M NaCl fraction (Fraction IV) (see Figure 8, lane 4). In order to remove these contaminating proteins, Fraction IV was subjected to an additional chromatographic step on blue-agarose. The degradosome eluted from the column as a broad peak from 0.3 M-3 M NaCl. The pooled and concentrated blue-agarose fraction (Fraction V ; not shown) was then subjected to gel filtration chromatography on BioGel A 1.5 M (see section 2.2.5.8) instead of preparative glycerol gradients as previously published -80-M II IV VI K D a 200 116 97 66 4 5 31 Rne (180 kDa) * Pnp (85 kDa) RhlB (50 kDa) Enolase (48 kDa) 3 4 5 Figure 2.8 Purity of the degradosome from strain CF881. Cell growth, harvest, lysis and enzyme purification are described under 2.2.5. The following samples were denatured, separated through a 10% SDS-polyacrylamide gel and stained with Coomassie-blue: lane 1, molecular mass markers (Bio-Rad); lane 2, S-200 (Fraction II) (50 pg); lane 3, resuspended 26% w/v ammonium sulfate pellet (Fraction III) (50 pg), lane 4; pooled fractions from chromatography of Fraction III on SP-sepharose (Fraction IV) (20 pg); lane 5, pooled fractions from chromatography of Fraction V by gel filtration (15 pg) (Fraction VI Cormack et al., 1993). The four major protein bands Rne (180 kDa), Pnp (85 kDa), RhlB (50 kDa), and enolase (48 kDa) are indicated in the margin to the right. The 97 kDa band, denoted by an asterisk, represents a proteolytic fragment of the Rne protein as determined by western blotting (Py et al., 1996; S. K. Masterman and G. A. Mackie unpublished). -81-(Carpousis et al, 1994). The final fraction, Fraction VI, was judged to be comparable to other published preparations (Carpousis et al, 1994; Py et al, 1994; 1996; Miczak et al, 1996; Blum et al, 1997) in both composition and quality by inspection of Coomassie-blue-stained SDS-polyacrylamide gels (see Figure 8, lane 5). Four major protein bands are detected in the degradosome preparation; Rne (180 kDa), Pnp (85 kDa), RhlB (50 kDa) and enolase (48 kDa). The 100 kDa band, previously denoted as * (Carpousis et al, 1994; Py et al, 1996), is a proteolytic fragment of the Rne protein as shown by western blotting (Py et al, 1996; data not shown). The presence of RNase E and PNPase in the degradosome preparation was confirmed by both western blotting and by activity assay (S. K. Masterman and G. A. Mackie unpublished; see section 2.3.9). Two other faint bands, which migrate with apparent molecular weights of 69 kDa and 80 kDa are, presumably, DnaK and polyphosphate kinase respectively. Consistent with previous reports, these proteins are not associated with the degradosome stoichiometrically (Miczak et al, 1996; Blum et al. 1997). Other notable contaminants are the presence of the P and P' subunits (150 kDa, 155 kDa respectively) of R N A polymerase which have been observed in at least one other degradosome preparation (Carpousis et al, 1994). 2.3.9 Sensitivity of 9S RNA to degradosomes Historically, the conversion of 9S R N A to the precursor to 5S rRNA has been the defining in vitro assay for RNase E activity. We have tested the purified degradosomes against 9S R N A to confirm the presence of RNase E activity in the complex isolated from strain CF881. Degradosomes were incubated with labeled 9S R N A at a final concentration of 5 ng/ul for 60 minutes at 30 °C (Figure 9B). During the first 15 minutes of digestion, the 9S R N A substrate is rapidly converted to -82-5" B 246 207 A i B I 0 2.5 5 7.5 10 15 20 30 45 165 126 - * • mm m mm 9 S 4 S 7 S 5S 8S 2 S 9 S 8 S 7 S 5 S 81 — 4 S Figure 2.9 The action of the degradosome against 9 S RNA. Panel A shows a schematic representation of the 9S R N A precursor. The letters A and B represent the two RNase E cleavage sites which liberate the pre-5S rRNA precursor. The various cleavage products 2 S-8 S are indicated in the margin to the right. Panel B shows a typical assay for RNase E activity using degradosomes at a final concentration of 5 ng/pl as described in section 3.2.5. Preparation of substrates and assay conditions have been published (Cormack and Mackie, 1992) and are described in Chapter 3. At various time points (indicated in minutes) aliquots were removed and quenched in 3 volumes of loading buffer containing 90% formamide and tracking dyes as described in section 3.2.5. The products of the reaction were separated electrophoretically through 6% polyacrylamide gels containing 8 M urea and then visualized with a Molecular Dynamics Phosphorlmager system. The 9 S R N A substrate and the various intermediates are indicated in the margin to the right while the sizes of the R N A fragments (in nucleotides) are denoted in the margin to the left. -83-pre-5S rPvNA (Figure 9B). Three transient intermediates, 8S, 7S and 4S R N A also appear during the reaction. While all other RNAs are digested by the degradosome during prolonged incubations, the pre-5S rRNA remains stable for 60 minutes of digestion in the presence of both RNase E and PNPase (data not shown). 2.3.10 RNA-binding properties of the degradosome In order to investigate the RNA-binding activity of the degradosome and to confirm the results of Py et al. (1996), "northwestern blotting" experiments were performed on aliquots of purified degradosome and, for comparison, on purified recombinant RhlB protein. Protein blots were probed with labeled rpsT mRNA (Figure 10, lanes 3 and 4) or with a fragment of the malE-malF mRNA (Figure 10, lanes 1 and 2) for 90 minutes as previously described (Cormack et al, 1993). Consistent with previous reports (Cormack et al, 1993; McDowall and Cohen, 1994; Taraseviciene et al, 1995; Py et al, 1996), the 180 kDa Rne/Ams protein binds strongly to the labeled rpsT and malERNAs. Proteolytic fragments of the Rne/Ams protein (100 kDa and 80 kDa) are also capable of binding to R N A as shown previously (Cormack et al, 1993; McDowall and Cohen, 1994; see Figure 2.10). In the case of the 100 kDa fragment, the observed relative RNA-binding is comparable to that of the intact Rne/Ams protein. In addition, both native and recombinant RhlB (50 kDa) are capable of binding to the rpsT and malERNAs, albeit to a lesser degree than the Rne/Ams protein as described previously (Py et al, 1996). In contrast, RNA-binding by the a-subunit (85 kDa) of PNPase was not detectable by "northwestern blotting" as previously described (Py et al, 1996). Overall, the RNA-binding pattern for our degradosome preparation was not significantly different from that of previous authors (Py et al, 1996). -84-Figure 2.10 The RNA-binding properties of the degradosome and RhlB. Degradosomes (16.8 pg) (lanes 1 and 3) and purified RhlB (1.2 pg) (lanes 2 and 4) were separated by 10% SDS-PAGE, electroblotted onto nitrocellulose and probed with either 32P-labeled rpsT mRNA (lanes 3 and 4) or 3 2P-labeled malE REP R N A (lanes 1 and 2) as described in Cormack et al, 1993. The blots were washed three times and the labeled proteins were visualized with a Molecular Dynamics Phosphorlmager system. -85-2.4 Discussion Although purification of 3'-exonucleases to homogeneity was first accomplished over 20 years ago by Gupta et al. (1977) (RNase II) and Soreq and Littauer (1978) (PNPase), previous purifications from non-overexpressing strains required large quantities of cells (1 kg; Soreq and Littauer, 1978) and often relied on tedious, large scale purification methods. We have, therefore, developed more rapid and efficient purification strategies by utilizing the T7 overexpression system (Studier etal, 1990). The T7 system offers significant advantages since proteins which normally represent a small fraction of the total cellular protein, < 1%, may be expressed to extremely high levels, as high as 20%. Thus, in many cases, only a 5-fold purification of the protein is required to obtain a nearly homogeneous preparation. At first glance, one would predict that overexpression of the 3'-exonucleases in E. coli would result in anomalous cellular R N A decay which would, in turn, lead to attenuated cell growth and perhaps cell death. However, both RNase II and PNPase can be expressed to very high levels for several hours after induction without exhibiting significant defects in cell growth or mRNA decay (K. Baker and G. Mackie, unpublished). These observations are consistent with a secondary role for these enzymes in mRNA decay. In contrast to these results, strains which overexpress the catalytic subunit of RNase E, the Rne/Ams protein, show a significant decrease in growth rate (J. L . Genereaux and G. A. Mackie, unpublished data). The fact that RNase E is toxic at high levels is suggestive of a key role in cellular R N A metabolism. In addition to the use of the T7 system to maximize gene expression, we and others have made significant improvements in the purification of the 3'-exonucleases by exploiting the inherent RNA-binding properties of these enzymes. A combination of two "affinity" steps plus additional -86-chromatographic steps including: ion exchange chromatography, hydroxylapatite chromatography and/or gel filtration led to rapid and facile preparations of the 3'-exonucleases with good yields and activity. One successful "affinity" method relies on immobilized dye chromatography (Cudny and Deutscher, 1980; M . P. Deutscher, personal communication). Cibacron blue (Ciba-Geigy) is a textile dye consisting of a polysulfonated (anthraquinone) chromophore linked to a chlorotriazine group by an aminoether bridge (Stellwagen, 1990) (see below). The blue dye functions as an ionic, hydrophobic, aromatic or sterically active ligand in various applications. Blue-agarose chromatography has been applied successfully in the purification of a number of different proteins from a wide variety of organisms and is of particular value in the purification of kinases, dehydrogenases, and other nucleotide-dependent enzymes (Thompson etal., 1976; Vasquez etal., 1979; McCoy et al, 1979). It has been suggested that enzymes which contain a dinucleotide fold can bind specifically to blue-agarose (Thompson et al, 1976). Most of the E. coli exoribonucleases have been successfully enriched by blue-agarose chromatography (M. P. Deutscher, personal communication), although they are unlikely to contain such a fold. s o , -0 NHj -87-R N A affinity chromatography on poly(A)-, poly(U)- or RNA-sepharose has also proven to be a useful technique for the purification of PNPase (Barbehenn et al, 1982; Soreq and Littauer, 1982). These supports are expensive and are not usually reusable since they can be digested by ribonucleases during purification. One alternative, heparin-agarose, has been previously utilized in the preparation of a number of nucleotide- and nucleic acid-binding proteins (Zhang et al, 1991; Bhikhabhai etal, 1992; Moore and Blobel, 1993). We have successfully substituted heparin-agarose in the purification of the 3'-exonucleases. Presumably, since heparin (see below) mimics the sugar-phosphate backbone of a nucleic acid, it may interact with the 3'-exos specifically via their R N A -binding domains. Another alternative to traditional purification methods involves the use of recombinant D N A techniques to generate fusion proteins which contain 6 or 10 histidines at either the N - or C-terminus of a target protein. Histidine-tagged fusion proteins can be purified rapidly in one step by nickel chelation chromatography (Van Dyke et al, 1992). His-PNPase, containing 7 N-terminal histidine residues, was successfully purified by these methods and was used to generate PNPase-specific antibodies (Py et al, 1994; K. H. Niguma and G. A. Mackie, unpublished data). Interestingly, oscv NHSO; -88-however, when assayed against poly(A), the purified His-PNPase fusion protein lacked any detectable phosphate-dependent exonucleolytic activity (G. A. Coburn and G. A. Mackie unpublished observation). The R N A helicase, RhlB, has also been purified as a His-RhlB fusion protein (Py et al, 1996). Although His-RhlB has proven useful for the production of RhlB-specific antibodies, it does not appear to function as an R N A helicase in the absence of other components of the degradosome (Py etal, 1996). Likewise, N-terminal modifications of the Rne/Ams protein (either His-tagged or GST fusion) result in partial or complete inactivation of RNase E (J. L . Genereaux, L . M . Duncan and G. A. Mackie, unpublished observations). It is possible that addition of a fusion tag may disrupt the quaternary structure of the enzyme and/or protein-protein interactions among components of degradosome. Thus, the use of fusion protein constructs to facilitate purification of the enzymes of mRNA decay has been somewhat disappointing. A determined effort to purify RNase E led to the finding that RNase E copurifies with a number of other proteins including PNPase (Carpousis et al, 1994). Interestingly, the only difference between many steps in the original RNase E preparation (Roy and Apirion, 1983) and that of the degradosome was the inclusion of a cocktail of protease inhibitors and careful work at 4°C. It is unfortunate that more than a decade passed before this important "trick" was uncovered. To date the degradosome has been purified by a variety of methods including immunoprecipitation (Carpousis et al, 1994; Py et al, 1994) and immunoaffinity chromatography (Miczak etal, 1996); however, purification from haploid strains by conventional techniques remains the favoured method. Whether or not the degradosome is the physiological source of RNase E remains to be demonstrated. However, it is clear that our understanding of this important enzyme has been severely limited by the lack of suitable methods to achieve homogeneous preparations of RNase E. Further -89-investigation of the role of this important enzyme will require the development of novel purification strategies followed by reconstitution experiments. We are presently exploring the possibility of utilizing secretion systems in both Pichiapastoris and E. coli to achieve this end. -90-CHAPTER 3 The action of the 3'-exonucleases against a synthetic RNA substrate 3.1 Introduction Unlike ribosomal and transfer RNAs, conventional depictions of mRNA assumed that it was relatively unstructured,. However, it is now clearly accepted that mRNAs are folded and contain important structural determinants within both the 5'- and 3- untranslated regions of the molecule which influence both the functional and chemical half-lives of their respective messages (Belasco et al, 1985; Emory and Belasco, 1990; Emory et al, 1992; Bouvet and Belasco, 1992; Mott et al, 1985; Newbury et al, 1987; McLaren et al, 1991; Mackie, 1992). Internal stem-loop structures, which likely re-fold behind translocating ribosomes, may also contribute to the chemical and, to a lesser extent, functional half-lives of mRNA. The degradation of highly structured R N A presents a unique problem for the cell since the major enzymes of mRNA decay, RNase E, PNPase and RNase II, are single-strand specific (Ehretsmann et al, 1992; Mackie, 1992; Littauer and Grunberg-Manago, 1998; Cannistraro and Kennell, 1994) and the cleavage specificity of RNase III is relatively stringent (Court, 1993; Nicholson, 1996). Early models of mRNA decay predicted that endonucleolytic cleavages would eliminate structural impediments to the 3'-exonucleases by providing new free 3'-ends (Belasco and Higgins, 1988; Higgins etal, 1992; Higgins etal, 1993). However, these relatively simple models failed to explain how the highly-folded R N A remnants might be degraded. One novel mechanism would invoke the use of an R N A helicase to unwind R N A secondary structure. R N A unwinding -91-would, therefore, permit access by RNase E to otherwise buried cleavage sites and/or remove structural impediments to the 3'-exonucleases. As previously discussed in section 1.3.2, E. coli contains at least five known putative DEAD-box R N A helicases (Kalman et al, 1991). It is possible, therefore, that one or more of these putative helicase activities may play a role in R N A stability. We have employed the use of a simple R N A unwinding assay (Rozen et al, 1990) to search for activities in crude extracts from E. coli, which may participate in mRNA decay. Helicase activity is measured by the conversion of a partially duplexed R N A substrate to single-stranded monomers. Instead of detecting activities which could unwind the duplex, we observed the shortening of the substrate by an activity which was consistent with RNase II. The helicase/exonuclease substrate was further utilized to characterize some of the properties of purified recombinant RNase II and PNPase. Our results demonstrate that upon stalling 3' to the base of stable R N A secondary structure RNase II and PNPase dissociate from their substrates and reassociate with a new single-stranded 3'-end. Most of these results have been published elsewhere (Coburn and Mackie, 1996a). 3.2 Materials and methods 3.2.1 Enzymes and chemicals Purified recombinant RNase II (Fraction V, Figure 2.2; lane 8) PNPase (Fraction IV, Figure 2.5; lane 5) were prepared as described in sections 2.2.5.6 and 2.2.5.7 respectively. T7 and SP6 R N A polymerase, RNAguard and DNase I were obtained from Amersham-Pharmacia and Promega. [a 3 2P]CTP (3000 Ci/mmol), for R N A labeling, was obtained from Amersham-Pharmacia. Al l other chemicals are described in section 2.2.1. -92-3.2.2 Bacterial strains and plasmids The E. coli strains CF881 and 18-11 are described in section 2.2.2. The plasmid pRP40 containing the synthetic R N A helicase substrate, R N A I (hereafter called t40B to avoid confusion with the anti-sense regulator of colEl-type plasmids) (Pause et al, 1992), was obtained from Dr. N . Sonenberg (McGill University, Montreal). Plasmid pGM87, contains the P2 leader, coding sequences and 3-untranslated region of the rpsT mRNA encoding ribosomal protein S20 under the control of a T7 promoter in pTZ18U (Mackie and Genereaux, 1993). Plasmid pJG175, encompassing residues 268-447 of the rpsT mRNA under the control of a T7 promoter, was described previously (Mackie etal, 1997). The plasmids, pGM87p(A) and pJG175p(A), containing a 30 bp poly(A)-poly(T) tract were constructed by ligating two annealed oligonucleotides (GC-10: 5'-GATTCC(A)3 0 -3' and GC-11: 5'-CTAGA(T)3 0-3') between the BamHl and Xbal restriction sites immediately adjacent to the Rho-independent terminator within pGM87 and pJG175 respectively (Coburn and Mackie, 1998). 3.2.3 Preparation of crude extracts Cultures (10) of strain CF881 and 18-11 were grown with good aeration at 37 °C to late log phase (A600 of 1.0). Cells were harvested as described in section 2.2.5.3 and the cells were frozen at -70 °C until use. The thawed cells were resuspended in 3 volumes of buffer A (60 mM Tris-HCl, pH 7.5,10 mMMgCl 2 , 60 mMNFLCl, 0.05 mMEDTA, 1 mM DTT) and ruptured in a French pressure cell as described in section 2.2.5.3. The cell lysate was centrifuged at 30,000 x g for 30 minutes in a Beckman JA-20 rotor at 4 °C. The supernatant (S-30) was then centrifuged at 150,000 x g in a Beckman Ti70.1 rotor for 2 hours at 4°C. The supernatants, S-30 and S-150, were the source of crude extracts for subsequent experiments. -93-3.2.4 In vitro synthesis of RNA transcripts Transcription reactions contained 20-50 ng/pl of linearized plasmid (summarized below in Table 3.1), 20 mMDTT, 1 mM ATP, GTP, UTP, 0.1 mM CTP, 30 pCi [a3 2P] CTP (3,000 Ci/mmol) (Amersham-Pharmacia), 1 U/pl RNAguard and either 2.5 U/pl SP6 or T7 R N A polymerase (Promega or Amersham-Pharmacia) in a 100 pi reaction volume. Reactions were buffered with Promega transcription buffer containing 40 mM Tris-HCl, pH 7.5, 6 m M M g C l 2 , 2 mM spermidine and 10 mM NaCI according to the manufacturers' instructions. Synthesis of uniformly labeled R N A was directed from either an SP6 or T7 promoter contained within the plasmid constructs for 90 minutes at 37 °C. Ten units of DNase I were added and the reaction mixture was incubated for an additional 15 minutes at 37 °C. The quantity of R N A synthesized, in pmoles, was determined by calculating the percentage of radiolabeled nucleotide incorporated into RNA. This was accomplished by spotting 2 pi onto a glass filter (Whatman GF-C) (to give total radioactivity in the reaction) and precipitating an additional 2 pi in 5% (w/v) trichloroacetic acid on ice for 30 minutes (to give total radioactivity incorporated into RNA). The precipitate was collected by filtration and the glass filters were counted in a Beckman LS6000 IC scintillation counter for Cerenkov radiation. Ammonium acetate was added to the transcription reaction to a final concentration of 2 M , the mixture was extracted once with an equal volume of phenol/chloroform/isoamyl alcohol (24:25:1) and precipitated with 2.5 volumes of 95% ethanol overnight at -20 °C or for several hours at -70 °C. The R N A was pelleted by centrifugation at 14,000 rpm in a 4°C microcentrifuge, washed once with 1 ml of 80% ethanol and dried under vacuum in a Speed-Vac (Savant). The pelleted R N A was resuspended in 0.25 M sodium acetate and precipitated in 2.5 volumes of 95% ethanol a second time as described above. The in vitro transcripts were then resuspended in 50 pi of DEPC (diethyl pyrocarbonate) treated dH 2 0 and stored at -20 °C -94-until use. 5-end-labeled RNAs were prepared as described above with the following exceptions. The transcription reaction were initiated with a 5'-32P-labeled ApG dinucleotide at a concentration of 50 p M for 5 minutes at 37 °C in the presence of 1 mM each of UTP, ATP, CTP and 50 p M GTP. The concentration of GTP was increased to 1 mM and the reaction was incubated for an additional 45 minutes at 37 °C. Alternatively, 5'-end-labeled RNAs were prepared by transcription of linear D N A templates in the presence of 50 pCi of [y3 2P]-GTP to a final concentration of 50 pM. ATP, CTP and UTP were present at 1 mM in all reactions. The products were purified as before. Table 3.1 Summary of plasmid-encoded transcripts. Plasmid Polymerase Linearization R N A Substrate Coordinates1 pRP40 SP6 BarnHL t40B pGM87 T7 Dral rpsT/365 92-447 pJG175p(A) T7 Dral rpsT/209-po\y(A) 268-447+ poly(A) 1. Coordinates of the rpsTmKNA are described in Mackie (1992). 3.2.5 Assay for RNase II activity Assays for RNase U activity were assembled in a 70 pi reaction volume containing 10 pmoles of labeled t40B in a reaction buffer containing 17 mM HEPES-NaOH, pH 7.5, 0.5 mM Mg-acetate, 100 mM KCI, 2 mM DTT, 5% (v/v) glycerol and 10 pg/ml acetylated B S A (New England Biolabs) as described (Coburn and Mackie, 1996a). Protein was added last to the final concentration specified -95-in the Figure legends and incubations were performed at 37 °C. Samples were withdrawn at various times and quenched in 3 volumes of loading buffer containing 90% deionized formamide, 22 mM Tris, 22 mM boric acid, 0.5 mM EDTA, 0.1% xylene cyanol FF and 0.1% bromophenol blue. The products were resolved by electrophoresis on 10% polyacrylamide gels containing 8 M urea and visualized by autoradiography or with a Molecular Dynamics Phosphorlmager system. Al l assays were performed a minimum of two or three times to ensure reproducibility. Degradation assays expressed as rates of substrate decay or product formation (see text) were calculated as the average of at least two experiments. 3.2.6 Assays for PNPase activity 3.2.6.1 Method A Assays for PNPase activity were assembled in a 70 pi reaction volume containing 10 pmoles of labeled t40B in a reaction buffer containing 20 mM Tris-HCl, pH 7.5, 1.5 m M DTT, 1 mM MgCl 2 , 20 m M KC1 and 10 mM Na-phosphate. Purified recombinant PNPase was added last to a final concentration specified in the Figure legends, and the reactions were performed at 37°C. Samples were withdrawn at various times, quenched with three volumes of loading buffer and processed as previously described for RNase II (see section 3.3.5 above). 3.2.6.2 Method B PNPase assays were also performed under conditions which have been previously published for RNase E activity (Mackie, 1991). Reactions were assembled in a 40 pi reaction volume containing 25 mM HEPES-NaOH, pH 7.5, 5% (v/v) glycerol, 5 mM MgCl 2 , 100 mM NH 4C1, 0.02 m M EDTA, -96-and 60 rnMKCl. The reaction was supplemented with 10 mM Na-phosphate and the incubations were performed at 30 °C. The products of the reaction were analyzed as above (see section 3.2.5). 3.2.7 UV photochemical crosslinking ofproteins to RNA Assay mixtures were prepared as described in section 3.2.5 with 160 frnoles of t40B substrate. After incubation on ice for 2-5 minutes, the sample was subjected to one 2-6 ns pulse (40-50 mJoules) with a 266 nm U V laser (Spectra Physics) as previously described (Ho et al, 1994). The sample was then incubated with 5 pg of RNase A and 5 U of RNase T l at 37 °C for 45 minutes to remove excess RNA. Each digested sample was boiled in an equal volume of SDS sample buffer and separated electrophoretically on a 15% SDS-polyacrylamide gel. The crosslinked proteins were visualized by autoradiography. 3.3 Results 3.3.1 Exonucleolytic activity in crude extracts from E.coli A partially duplexed R N A substrate (Figure 3.1a) was used to assay extracts, generated from various E. coli strains, for putative R N A helicase activities. Instead of detecting an activity which could unwind the duplexed R N A to monomers, we observed the partial degradation of the synthetic substrate in extracts which are wild type for RNase II activity but not in extracts deficient for a number of exonucleases including RNase II (Figure 3.1, compare panels b and c). Complete conversion of the 92-nt substrate to a relatively stable 77-nt degradative intermediate was observed in crude extracts prepared from strain CF881 over a 60 minute time course (Figure 3.1b). The exact size of the product obtained with pure RNase II (see below) was determined on a sequencing gel -97-(a) (b) (c) CF881 (S-150) 18-11 (S-150) RNase II wt RNase II -NP 0 10 20 30 60 NP 0 10 20 30 60 — s — — p Figure 3.1 3'-exonucleolytic degradation of a partial duplex RNA substrate. Panel a presents a schematic diagram of the 3'-exonucleolytic degradation of the 92 nucleotide R N A substrate on the left to the 77-nucleotide product on the right. A time course of digestion of the synthetic R N A transcript t40B (a) as described in section 3.2.4 is shown in panels b and c. Crude extracts (S-150 fraction) prepared from strain CF881 (b) or strain 18-11 (c) were added to a final concentration of 0.5 pg/ml and incubated with substrate at 37°C. Aliquots were removed from the reaction mixture at the times indicated (in minutes) as described in section 3.2.5. The digestion products were analyzed by electrophoresis through a 10% polyacrylamide gel under denaturing conditions. NP denotes a control lane containing substrate incubated in the absence of protein for 60 minutes. The 92 nucleotide substrate (S) and the stable degradative intermediate (P) (77 nucleotides) are indicated with arrows. -98-(shown in Figure 3.5; see below). In contrast, crude extracts prepared from strain 18-11 were unable to digest the substrate (Figure 3.1c). Several additional experiments were undertaken to confirm that the 77-nt degradation product (shown in Figure 3.1a) corresponds to the product of RNase II stalling 10 nucleotides 3' to the double-stranded region of the substrate. First, the denatured 77-nt product retains a 5-end-label (shown in Figure 3.4; see below). Secondly, the partial duplex substrate is resistant to digestion by the purified Rne/Ams protein, the catalytic subunit of RNase E (Cormack et al, 1993), under conditions where authentic substrates would be processed to completion (data not shown). Third, the other major exonucleolytic activity in E. coli, PNPase, is completely dependent on phosphate under the assay conditions and is not active against the synthetic substrate in the absence of phosphate (Figure 3.2a compare lanes 1-4 with lanes 5-8). Moreover, extracts prepared from a strain containing the mutantpnp-7 allele, which largely lacks PNPase activity, also generate the 77-nt product in the presence or absence of phosphate (data not shown). Although contributions from other exonucleases cannot be excluded completely, the observed partial degradation is most consistent with RNase II activity. This was confirmed (see below) using the purified recombinant Rnb protein. Early in vitro studies of the decay of tRNA suggested that PNPase would "power" through secondary and tertiary structure to processively phosphorylyze structured RNAs (Thang et al, 1967). Accordingly, the appearance of product (approximately 70-77-nt) is relatively transient in a prolonged incubation with crude extracts containing PNPase activity compared to the stable digestion product generated by the action of RNase II (compare Figure 3.2a, lanes 6-8 with Figure 3.3a, lanes 3-6). Thus, in crude extracts, PNPase appears to stall briefly at the base of the duplex and then processively phosphorolyzes through the structure as inferred by previous studies (Thang etal, 1967; McLaren -99-(a) (b) - P i + Pi NP 10 20 30 NP 10 20 30 + Pj NP 10 20 30 S — P — S' 1 2 3 4 5 6 7 8 1 2 3 4 Figure 3.2 Degradation of the partially duplexed RNA by PNPase activity in crude extracts, (a) Crude extracts (S-30 fraction) prepared from strain 18-11 at a final concentration of 0.2 mg/ml were incubated with t40B in the absence (lanes 1-4) or presence (lanes 5-8) of 10 mM phosphate at 37°C. (b) Extracts (S-30 fraction) prepared from strain 18-11 at a final concentration of 0.2 mg/ml were incubated in the presence of 10 mM phosphate at 37°C with t40B which had been previously digested to 77-nucleotides (S*) with the purified Rnb protein (see legend to Figure 3.4), phenol/chloroform extracted and precipitated with ethanol. Aliquots were removed from the reaction mixture at the times indicated (in minutes) as described in section 3.2.4. The digestion products were analyzed by electrophoresis through a 10% polyacrylamide gel under denaturing conditions. The substrate (S), partially digested substrate (S*) and the degradative intermediate (P) are indicated with arrows. NP denotes a control lane containing substrate incubated in the absence of protein for 30 minutes. -100-etal, 1991). 3.3.2 Properties of recombinant RNase II 3.3.2.1 Activity against t40B The purified recombinant Rnb protein was active against the partial duplex t40B substrate in a manner similar to the activity originally detected in crude extracts from strain CF881 (Figure 3.3a, lanes 2-6). Under conditions in which enzyme is limiting (molar ratio of substrate to enzyme of 2300:1), the 3'-single-stranded tails are removed from the substrate during a 60 minute incubation at 37°C to generate a degradative intermediate which has been shortened by about 15 nucleotides. The appearance of the degradative product is linear for 30 minutes after which the rate declines gradually (Figure 3.3a, lanes 2-6). Thus each enzyme molecule is turning over more than 30,000 times. Digestion of the t40B transcript is complete after a 60 minute incubation with 2.0 mU of RNase II activity (Figure 3.3a, lane 7). Approximately 20% of the substrate is resistant to degradation by RNase II even after addition of 200 mU of fresh enzyme (data not shown). A fraction of the substrate appears to form concatemers and as a result does not have free 3'-ends accessible to the enzyme. Interestingly, digestion of t40B for 60 minutes at 37°C with 4 U of enzyme resulted in a further shortened (73-nt) but stable degradation intermediate depicted by the arrowhead (Figure 3.3a, lane 8). This experiment suggests that at high concentrations the Rnb protein can remove 3-4 additional unpaired residues prior to the t40B duplex remaining from a previous round of digestion (see below). -101-(a) (b) NP 0 10 20 30 60 60 60 NP 0 5 15 mmmmm s p 1 2 3 4 5 6 7 8 1 2 3 4 Figure 3.3 Degradation of the partial duplex RNA substrate by purified recombinant RNase II (Panel a) lanes 2-6, the t40B transcript was incubated with purified recombinant Rnb protein (1.2 mU) at a concentration of 5 ng/ml for the indicated times (in minutes) at 37°C as described in section 3.2.4. Lanes 7 and 8 show a 60 minute digestion of the t40B transcript incubated with the purified RNase II (2.0 mU and 4.1 TJ) at a concentration of 8.3 ng/ml and 16 pg/ml respectively. The digestion products were subsequently analyzed by gel electrophoresis. (Panel b) Thermal inactivation of the Rnb protein. The Rnb protein (0.5 pg/ml) was incubated in the absence of substrate for the indicated times (in minutes) prior to addition to a complete reaction mixture. Incubation was continued for an additional 15 minutes at a final concentration of 8.3 ng/ml. The digestion products were analyzed by gel electrophoresis. The substrate (S) and the degradative intermediate (P) are indicated by arrows while the shorter degradation product shown in lane 8 is indicated by an arrowhead. NP denotes control lanes containing substrate incubated in the absence of protein for 60 minutes. -102-We have also tested whether the 77-nt product would inhibit the activity of RNase II in subsequent rounds of digestion. In the first experiment, the Rnb protein (3.3 mU) was incubated with a 25 pmoles of unlabeled t40B (2.5-fold molar excess over labeled t40B) for 2.5 minutes at 37°C prior to addition of labeled t40B. The kinetics of digestion of labeled t40B over a 60 minute time course were identical to those in an incubation in which the same amount of enzyme was incubated directly with labeled t40B. This suggests that RNAs which lack free 3'-ends are not efficient substrates for RNase II. Two independent groups have reported that RNase II can protect "upstream" R N A sequences from PNPase attack through the formation of a stable RNA-RNase II complex (Hajnsdorf et al, 1994; Pepe et al, 1994). However, our finding that RNase II stalls and dissociates several nucleotides 3' to the base of a stable duplex suggests an alternative hypothesis. We believe that removal of a 3'-single-stranded overhang by RNase II generates a modified R N A which is, in turn, a poor substrate for PNPase. To further investigate this hypothesis, we have incubated the 77-nt product, produced by the action of RNase II (see above), with crude extracts prepared from strain 18-11 in the presence of 10 mM phosphate. The data demonstrate that the 77-nt product is resistant to digestion by a PNPase-like activity (Figure 3.2b, lanes 1-4), whereas the 70-nt digestion product generated directly by PNPase in crude extracts is relatively unstable and accumulates to sub-stoichiometric levels (Figure 3.2a, lanes 5-8). Thus, PNPase appears to require a single-stranded 3'-end to initiate processive phosphorolysis, not unlike the situation for RNase II. 3.3.2.2 Size of products 5'-end-labeled t40B R N A was digested with purified Rnb protein and the products were -103-8 7 H 1 " 0 5 10 15 20 25 30 Time (min) F i g u r e 3.4 Competition between partially digested t40B and complete t40B. Recombinant Rnb protein (2.0 mU) at a concentration of 8.3 ng/ml was incubated with intact t40J3 in the presence (•) or absence (•) of 10 pmoles of t40B which had been previously digested to yield a 77 nucleotide product with the purified RNase II. The products were resolved by gel electrophoresis and the relative amounts of t40B were quantified with a Phosphorlmager, expressed as pmoles of RNA remaining and plotted as a function of time. -104-separated on a sequencing gel for greater resolution. As shown in Figure 3.5, lane 3, the R N A product retains the 5'-end-label suggesting that it is generated by 3'-exonucleolytic digestion. The most abundant product is 77 nucleotides, although shorter products of 74-75 nucleotides are easily detected. An overdigestion of the t40B substrate is shown in Figure 3.5, lane 4. In this case, the most abundant product is 73 nucleotide residues, however, shorter products of 71-72 nucleotides are also detected. An additional product of 67 nucleotides was also detected at very low levels during the digestion (-1%). Thus, digestion of t40B by the Rnb protein removes single-stranded 3'-ends from the t40B R N A up to 4-7 nucleotides prior to the G-C duplexed stem. These results are in good agreement with those obtained previously for RNase II acting on tRNA precursors both in vitro and in vivo (Li and Deutscher, 1994; L i and Deutscher, 1996). 3.3.2.3 Thermolability of RNase II Several previously published reports have suggested that RNase II, of varying degrees of purity, is readily inactivated by heat (Spahr, 1964; Singer and Tolbert, 1965; Gupta etal, 1977; Ghosh and Deutscher, 1978; Cannistraro and Kennell, 1994). We have tested the purified Rnb protein and found that the recombinant enzyme is also susceptible to thermal inactivation (Figure 3.3b). A comparison of Figure 3.3b lanes 2 and 3, shows that less than 1% of the activity remains after a 5 minute incubation of RNase II in the absence of substrate at 37°C. Interestingly, the enzyme is stabilized in the presence of substrate and can remain active up to 60 minutes at 37 °C (Figure 3.3 a). Activity can also be stabilized by the addition of substrate to RNase II which has been partially inactivated by a brief incubation in buffer at 37°C. Once activity has been lost to thermal inactivation, however, it cannot be regained upon addition of substrate (data not shown). The 77-nt product also -105-Figure 3.5 Analysis of the products generated by RNase II digestion of NOB. (Panel a) 5'-end-labeled t40B RNA, prepared from pRP40 previously linearized by BamHl, was treated as follows: lane 1, partial digestion with 1.2 units of RNase T l in 8 M urea, pH 5.0 for 6 minutes at 50 °C; lane 2, partial alkaline digestion, pH 9.0 for 5 minutes at 100 °C; lane 3, digestion of t40B with 2.0 mU of RNase II activity for 60 minutes at 37 °C; lane 4, digestion of t40B with 4.0 TJ of RNase II activity for 60 minutes at 37 °C. The reactions were quenched by the addition of three volumes of loading buffer and the digestion products were analyzed electrophoretically through a 10% polyacrylamide sequencing gel containing 8 M urea and visualized by autoradiography. Selected G residues are indicated in the left margin. The sizes of the substrate (92) and the digestion products (77) and (73) are denoted in the margin to the right. The 10-bp GC duplexed region is also denoted in the margin to the right. (Panel b) A schemtatic diagram of the t40B substrate is shown. Arrows denote the most abundant products formed by limited RNase II digestion (77) and the overdigestion (73). -106--107-stabilized the enzyme against heating. The Rnb protein was incubated in the presence of 1.5 pmoles of partially digested t40B for 5 minutes at 37°C prior to incubation with full-length t40B transcript. The 77-nt product not only protected the Rnb protein against thermal inactivation but also appeared to stimulate the activity of the enzyme for the full-length substrate by approximately 2-fold (data not shown). The apparent stimulation may be attributable to a decreased rate of thermal inactivation. Taken together, the data demonstrate that the enzyme can be stabilized by both substrate and product. In contrast, both a single-stranded D N A oligonucleotide (33-mer) and double-stranded plasmid D N A inhibited the activity of RNase II but were unable to provide significant protection from heating (data not shown) unlike oligos of deoxy(C)27 which can reduce the rate of thermal inactivation (Cannistraro and Kennell, 1994). 3.3.2.4 Binding of the Rnb protein to substrate Stabilization of the Rnb protein by the digested t40B transcript implies that, in the absence of any free 3'-single-stranded ends, RNase II can bind R N A even if it is not a substrate. In order to test this hypothesis, t40B was incubated briefly with a large excess of the Rnb protein, sufficient to digest it to 73-nt and then subjected to U V photocrosslinking. The data in Figure 3.6, lane 1 shows labeling of a band of 70-72.5 kDa, the size expected for RNase II. In addition, there is label associated with a band of 14 kDa. We believe that the latter represents non-specific association between oligoribonucleotides and RNase A. The recombinant enzyme is, therefore, able to bind its product (Figure 3.6, lane 1) in the absence of any other proteins or cofactors. A 70-72.5 kDa protein, corresponding to the molecular weight of RNase II, was also labeled in crude extracts prepared from strain CF881 (3. 6, lane 3). All bands were sensitive to proteinase K treatment (Figure 3.6, lanes 2 -108-Figure 3.6 UVcrosstinking of t40B to purified recombinant Rnb protein and proteins in the S-150 fraction prepared from strain CF881. Labeled t40B was incubated with purified recombinant Rnb protein (2.1 TJ) at a concentration of 50 ug/ml or with 10 pg of an S-150 fraction prepared from strain CF881 (33 mU) at a concentration of 1 mg/ml, irradiated with U V , digested with ribonucleases, and then separated by SDS/PAGE as described in 3.2.7. A duplicate sample was treated with proteinase K (+) prior to electrophoresis (lanes 2 and 4). Lanes 1 and 2: purified recombinant Rnb protein; lanes 3 and 4, crude extract prepared from strain CF881. -109-and 4). A comparison of Figure 3.6, lanes 1 and 3 also demonstrates that U V crosslinking can provide an important assessment of the purity of the enzyme preparation in light of the affinity chromatography techniques utilized in the purification. Since there are a large number of RNA-binding proteins in crude extracts prepared from E. coli which have a significant affinity for the t40B transcript, the presence of even a small percentage of these contaminants would be readily detected in the purified material (Figure 3.6, compare lane 3 to lane 1). 3.3.3 The activity of the RnbCA-38 and RnbCA-10 proteins against the t40B substrate RNase II activity in crude fractions from strain CF881 (S-30 fraction) was compared to an identical fraction generated from strain GC1 which overexpresses the RnbCA-38 protein, a truncated form of the Rnb protein which lacks 38 C-terminal amino acids. Serial dilutions of the two extracts were prepared and subsequently incubated with t40B R N A as described in section 3.2.5. RNase II activity was measured qualitatively by the relative formation of the 77-nt product of digestion corresponding to the removal of 15 nucleotides from t40B (data not shown). Although the RnbCA-38 protein is expressed to high levels and is soluble, partially purified fractions from the overexpression strain, G O , are not enriched significantly for RNase II activity suggesting that the RnbCA-38 protein is inactive. We have also tested the ability of RnbCA-10, which lacks 10 C-terminal amino acids to digest the t40B substrate. The solubilized, renatured and purified RnbCA-10 protein was incubated with the t40B substrate for 90 minutes at 37 °C at a final concentration of 260 ng/pl. During the digestion, the 3'-single-stranded tails were gradually removed from the substrate to generate a product which has been shortened by 15 nucleotides (Figure 3.7, lanes 2-7). The 77-nt product accumulates linearly -110-NP 0 10 20 30 60 90 92 77 1 2 3 4 5 6 7 Figure 3.7 Action of the RnbCA-10 protein against t40B RNA. The RnbCA-10 protein was overexpressed solubilized, renatured and purified by Resource-Q chromatography as described in section 2.2.8. A 6 pi sample of the Resource-Q fraction was incubated with 10 pmoles of labeled t40B for 90 minutes at 37 °C. Samples were removed at the various times indicated (in minutes) and quenched in three volumes of loading buffer. The products of the digestion were separated electrophoretically through 10% polyacrylamide gels containing 8 M urea and then visualized by autoradiography. NP denotes an incubation of the t40B substrate in buffer alone. The sizes of the substrate (92) and stable intermediate (77) (in nucleotides) are shown in the margin to the right. - I l l -throughout the assay and is relatively stable for 90 minutes demonstrating that the preparation is free of contaminating nucleases (Figure 3.7, lane 7). These results show that the RnbCA-10 protein, lacking 10 C-terminal amino acid residues contains renaturable RNase II activity. The specific activity of the RnbCA-10 protein acting against poly(A) R N A was calculated to be 50 mU/mg, a fraction of that obtained for the full-length recombinant Rnb protein. For this reason, the renatured RnbCA-10 protein was not studied further. 3.3.4 Properties of recombinant PNPase Purified recombinant PNPase (3 mU) was also active against the partial duplex t40B substrate in a manner similar to RNase II (compare Figure 3.3a, lanes 2-6 and Figure 3.8, lanes 1-6). Under conditions in which enzyme is somewhat limiting (molar ratio of substrate to enzyme, 30:1), the 3-single-stranded tails are removed from 30-40% of the substrate molecules during a 60 minute incubation at 37°C (Figure 3.8, lanes 1-6). Interestingly, under the conditions of Method A described in section 3.2.6.1, the kinetics of t40B digestion appear to be biphasic. Tail removal occurs rapidly within the first 10 minutes of incubation and then becomes more gradual over the next 50 minutes of digestion (Figure 3.8 compare lane 2 with lanes 3-6). Although the product was not examined on a sequencing gel, the degradative intermediate appeared to be shortened by an estimated 20-24 nucleotides. More importantly, the appearance of the 70-nt product is completely dependent on the presence of phosphate demonstrating that the preparation is free of contaminating RNase II activity (shown in Figure 3.8, lanes 7 and 8). Previous authors have suggested that PNPase remains bound to its substrate upon stalling 3' to the base of stable R N A secondary structure (Guarneros and Protier, 1990; McLaren etal, 1991). -112-0 10 20 30 60 90 o 90 m m m m m 92 70 1 2 4 5 7 8 Figure 3.8 Degradation of the partially duplexed t40B RNA substrate by purified PNPase. In lanes 1-6, the t40B transcript (10 pmoles) was incubated with purified recombinant PNPase (3 mU) at a concentration of 2.14 ug/ml for the indicated times (in minutes) at 37°C by Method A, as described in section 3.2.6, supplemented with 10 mM phosphate. Lanes 7 and 8 show a 60 minute digestion of the t40B transcript incubated with an equivalent amount of purified PNPase in the absence of 10 mM phosphate. The digestion products were subsequently analyzed by gel electrophoresis as described. The sizes of the substrate (92) and product (70) are indicated (in nucleotides) in the margin to the right. -113-To fUrther examine this property, we have tested the ability of "stalled" PNPase to cycle from one substrate to another. Purified PNPase (2 mU) was incubated with t40B for 30 minutes at 37 °C (Figure 3.9b, lanes 1-5) prior to the addition of a second substrate, /7>s77209-poly(A) RNA, which contains a 30-nt single-stranded 3'-poly(A) tail (a cartoon of which is shown in Figure 3.9, panel a). As before, single-stranded 3-ends were gradually removed from the t40B substrate to generate the 70-nt product. At T=30 minutes, /7?s77209-poly(A) RNA, was added to the mixture and the reaction was incubated for an additional 15 minutes at 37 °C (Figure 3.9b, lanes 5-9), lanes. After 15 minutes of incubation, the /7_«77209-poly(A) R N A substrate remained fully intact as almost none of the 3'-single-stranded poly(A) tails were digested by PNPase to generate the 179-nt intermediate (Figure 3.9b, lane 5). Similar results were obtained from the reciprocal experiment or from competition experiments with additional single-stranded substrates (data not shown). Under the low salt conditions in Method A (see section 3.2.6.1), we calculate that each PNPase trimer turns over approximately 370 times in 30 minutes, corresponding to the digestion of 12 poly(A) tails. Thus under relatively low salt conditions, PNPase does not readily dissociate upon stalling at regions of R N A secondary structure. Contrasting results were obtained when relatively moderate ionic strength conditions of Method B were employed in PNPase assays. Under conditions in which enzyme is limiting (molar ratio of substrate to enzyme, 186:1), the 3'-single-stranded poly(A) tails were rapidly removed from r/?s77209-poly(A) R N A to generate an intermediate (rpsTI\19 RNA) which has been shortened by 30 nucleotides (Figure 3.10, panel a). The formation of this degradative intermediate is linear for 20 minutes after which the substrate becomes limiting. Under these conditions each PNPase trimer turns over approximately 5,600 times or 15-fold more often than under the low salt strength conditions -114-(a) S t e mAV " Stem VII C A r A * * AAAAAAAAAAAAAAA ' I, 2 6 8 4 4 7 268 447 rps77209-poly(A) rpsT/179 (b) 0 2.5 5 10 30 31 32.5 37.5 45 Figure 3.9 The action of PNPase in a low ionic strength buffer. A schematic representation of the r/?s77209-poly(A) R N A substrate is shown in panel a. In panel b, purified recombinant PNPase (2 mU) was incubated with 10 pmoles of labeled t40B for 30 minutes at 37 °C by Method A supplemented with 10 mM phosphate, as described in section 3.2.6. At T=30 minutes an equivalent amount of A7?s77209-poly(A) R N A was added to the mixture and the reaction was incubated for an additional 15 minutes. At the various times indicated (in minutes) aliquots were removed and quenched with three volumes of loading buffer. The products of the reaction were analyzed by gel electrophoresis as described. The size (in nucleotides) of the substrates and products are indicated in the margin to the right. The presence of the 179-nt band in lanes 5-9 is the result of premature transcription termination by the T7 R N A polymerase. -115-of Method A. As shown in Figure 3.11, the shortening of the substrate follows first-order kinetics. These results imply that PNPase dissociates from its substrate and reassociates with a new free 3'-end upon stalling 3' to the base of stable R N A secondary structure. In order to determine whether PNPase shows a preference for free 3'-ends, we have tested the ability of rpsT/365 RNA, which terminates with a Rho-independent terminator stem-loop (similar to the structure of rpsTI\19 shown in the right side of Figure 3.9a; Mackie, 1992), to compete with /•/?s77209-poly(A) RNA. Equimolar concentrations of unlabeled rpsT/365 R N A and labeled rpsT/209-poly(A) R N A were incubated with 0.04 mU of PNPase for 30 minutes at 30 °C. Surprisingly, in the presence of the rpsT/365 RNA, the rate of formation of the rpsTI\19 product decreased approximately 1.7-fold compared to the rate observed for the r/w77209-poly(A) R N A alone (Figure 3.10, compare panels a and b; the data are plotted in Figure 3.11). Interestingly, addition of a 2-5-fold molar excess of unlabeled rpsT/365 RNA, did not further decrease significantly the rate ofrpsT/179 product formation (see Figure 3.11). Although PNPase shows a marked preference for single-stranded 3'-ends, our results suggest that RNAs with structured 3'-ends can compete with single-stranded substrates. -116-0 2.5 5 7.5 10 15 20 25 30 - 209-poly(A) - 1 7 9 (b) 0 2.5 5 7.5 10 15 20 25 30 i l l i i l A i - 209-poly(A) - 179 Figure 3.10 The action of PNPase in a moderate ionic strength buffer. Recombinant PNPase (0.04 mU) at a final concentration of 25 ng/ml was incubated with 0.8 pmoles of labeled Aps77209-poly(A) R N A (20 nM) in the presence (panel b) or absence (panel a) of equimolar concentrations of competitor R N A (20 nM) for 30 minutes at 30 °C. Reactions were performed by Method B , as described in section 3.2.6, supplemented with 10 mM phosphate. Aliquots were removed from the reactions at the various times indicated (in minutes) and quenched with three volumes of loading buffer. The products of the digestion were separated electrophoretically through 6% polyacrylamide gels containing 8 M urea and visualized with a Molecular Dynamics Phosphorlmager system. The size of the substrate (209) and product (179) are indicated (in nucleotides) in the margin to the right. -117-Figure 3.11 Competition between rpsT/365 RNA and rpsT/209-poly(A) RNA. Recombinant PNPase was incubated with 0.8 pmoles of labeled r/w77209-poly(A) R N A (20 nM) (•) in the presence of equimolar (20 nM) (•) or 2-fold molar excess unlabeled rpsT/365 competitor R N A (40 nM) (A) at 30 °C as described in the legend to Figure 10. Samples were removed at the times indicated (in minutes) and the products were resolved by gel electrophoresis. The relative amounts of rpsT/209-poly(A) R N A were quantified with a Molecular Dynamics Phosphorlmager system, expressed as log % R N A remaining and plotted as a function of time. -118-3.4 Discussion 3.4.1 The mechanism of action of RNase II on a novel substrate We envisage that the action of RNase II on t40B can be described by the following sequential steps: (1) binding to a free 3'-end on the 92-nt substrate; (2) processive hydrolysis of 15 phosphodiester bonds; (3) stalling of the enzyme approximately 9 unpaired nucleotides from the 10-bp G-C rich stem; (4) dissociation of the enzyme from the substrate; and (5) thermal inactivation of a fraction of the dissociated enzyme. The duration of each such cycle at steady state can be calculated from the apparent turnover number which we estimate as 9 nts-sec"1 based on a rate of 0.16 pmoles of product formed per minute at 4.3 fmoles of enzyme. This yields a cycle time of 1.67 seconds, the time to remove 15 nucleotides from each 3'-end (15 nts / 9 nts-sec"1). The time actually required for hydrolysis of 15 phosphodiester bonds (step 2 in the cycle) is only 0.21 seconds, however, as the reported turnover number for RNase II acting on poly(A) is 70 nts-sec"1 (Cannistraro and Kennell, 1994). If we assume that this turnover number also applies to the 15 residues removed from t40B and that no enzyme is lost to thermal inactivation (step 5), then steps 1, 3 and 4 account for 1.46 seconds (1.67 seconds - 0.21 seconds) of each cycle. As a consequence, RNase II cannot remain bound to a substrate once processive hydrolysis has ceased any longer than 1.46 seconds. The latter represents a maximum value for step 3 in the proposed cycle as binding (step 1), dissociation (step 4) and thermal inactivation (step 5) are not negligible. The first-order linear kinetics observed for the reaction demonstrate that RNase II stalls at regions of secondary structure, however briefly, but can disengage from the "stalled" substrate and reassociate with a new free 3'-end. This is substantiated by the demonstration that RNase II can cycle from an unlabeled to a labeled substrate. Our finding of dissociation from a substrate with 9 unpaired -119-protruding nucleotides at the 3'-end of the 77-nt product is in good agreement with the 10-15-nt digestion products obtained for RNase II acting processively on homopolymeric R N A substrates (Cannistraro and Kennell, 1994), The 10-15-nt oligoribonucleotide fragments produced by RNase II digestion of homopolymeric R N A can be degraded further by a non-processive mechanism involving successive hydrolysis/dissociation events to produce a limit digest of 2-4 nucleotide residues (Cannistraro and Kennell, 1994). Likewise, digestion of the t40B substrate under conditions of high enyme concentration releases shorter products (71-75 nucleotides) which are likely generated by non-processive "nibbling" at 3'-ends. A model summarizing the action of RNase II on t40B is shown in Figure 3.12. We envisage that RNase II binds to the extended single-stranded 3'-end (24 residues) and acts processively to hydrolyze -15 nucleotide residues (Figure 3.12, step 1). Upon reaching the 3' base of the stable G-C rich duplex, the enzyme stalls and releases an R N A fragment with -10 unpaired nucleotides protruding from the 3'-end (Figure 3.12, steps 2 and 3). Further digestion proceeds by non-processive hydrolysis of successive nucleotide residues to release a final product with 4-7 unpaired 3'-residues (Figure 3.12, steps 4 and 5). In good agreement with our findings, RNase II can participate in the processing of some tRNAs in vitro and in vivo by degrading long trailing sequences, but dissociates from the precursor to generate a tRNA molecule with 4 additional nucleotides (Li and Deutscher, 1994; L i and Deutscher, 1996). Final maturation of the tRNA is accomplished by other processing ribonucleases (Deutscher, 1993; L i and Deutscher, 1994; L i and Deutscher, 1996). Two lines of evidence suggest that RNase II can also reassociate with the 77-nt product of digestion. First, at relatively high concentrations (compare Figure 3.5 lanes 3 and 4) of enzyme -120-73 77 ^ _ 5' GCGCGCGCGCUAUAGUGAGUCGUAUUAGGGGAUC> ) I I I I I I I II I ^ - ^ 3" CGCGCGCGCG 5' 5' GCGCGCGCGCUAUAGUGAGU I II I I I I II I 3' CGCGCGCGCG 5" I 1 | 1 * 5" GCGCGCGCGCUAUAGUGAGU I I I I I I I I I I — W 3" CGCGCGCGCG 5" 5' GCGCGCGCGCUAUAGUGAG I I II I I I I I I 3- CGCGCGCGCG 5' 1 ' * * 5 5" GCGCGCGCGCUAUA I I I I I I I I I I 3' CGCGCGCGCG Figure 3.12 The mechanism of action of RNase II on t40B RNA. The nucleotide sequence (24 residues) of the 3'-end of the t40B substrate is depicted (Pause et al, 1993). RNase II binds to the extended single-stranded 3'-end and processively hydrolyzes approximately 15 unpaired nucleotides (step 1). The enzyme stalls 3' to the base of the stable G-C duplex denoted by the first arrow (step 2) and dissociates from the substrate (step 3). Successive rounds of hydrolysis/dissociation can shorten the 77-nt product further to release shortened products ranging in size from 71-75 nucleotides (steps 4-6) denoted by the second arrow. The processive enzyme is shown by the open "pac-man" cartoon while the non-processive enzyme is denoted by the black "pac-man" cartoon. -121-recombinant RNase II can remove 3-4 additional unpaired residues remaining from a previous round of digestion. Second, RNase II can bind its 73-77-nt product as evidenced by U V crosslinking and protection from thermal inactivation. Although partially digested t40B can bind to RNase II, it does not compete with the full-length substrate indicating that the preferred substrate for the enzyme has an extended free 3'-end. Moreover, the lack of competition by product implies that product binds to a site distinct from that of the substrate. 3.4.2 The deletions RnbCA-38 and RnbCA-10 lie within the putative SI RNA-binding domain of RNase II The solution structure of the S1 RNA-binding domain from PNPase was recently solved by N M R techniques (Bycroft et al., 1997). Data base searches have shown that this RNA-binding motif is conserved amongst a large number of different proteins including both RNase E and RNase II (Bycroft et al, 1997). Moreover, the S l -RBD lies within the catalytic domain of RNase E and encompasses residues which are critical for the activity of the enzyme (McDowall et al, 1993; McDowall and Cohen, 1996; L . M . Duncan and G.A. Mackie, unpublished data). Sequence alignments indicate that the SI RNA-binding domain of RNase II is located at the immediate C-terminus of the Haemophilus influenzae homologue (659 amino acid residues) (Bycroft et al, 1997) which is slightly larger than RNase II protein from E. coli (644 amino acid residues). In our attempts to clone the entire E. coli rnb gene we have inadvertently performed a deletion analysis of this region of the RNase II protein. While the RnbCA-10 deletion lacks 10 C-terminal amino acids and is insoluble, authentic RNase II activity can be renatured from this truncated protein. In contrast, the soluble RnbCA-38 deletion, which lacks 38 amino acids, is inactive. This crude analysis suggests that the -122-catalytic site or a site involved in RNA-binding lies between residues 606-634 which corresponds to the position of the Sl-RBD in H. influenzae (residues 571-659). This deletion likely removes the P-5 strand and possibly the P-4 strand of the S l -RBD which may form one face of the RNA-binding pocket. 3.4.3 The mechanism of action of PNPase As in the case for RNase II, the action of PNPase on t40B or /-/w77209-poly(A) R N A can be described by a similar set of sequential steps: (1) binding to a free 3'-end; (2) processive phosphorolysis of several phosphodiester bonds; (3) stalling of the enzyme at regions of R N A secondary structure; and (4) dissociation of the enzyme from the substrate; or (5) processive digestion through the secondary structure. Since the turnover number for PNPase acting on single-stranded homopolymers has yet to be determined, the duration of stalling could not be calculated. It is likely, however, that PNPase does not remain bound to its substrate for more than 1-2 seconds upon stalling at the 3'-base of stable R N A secondary structures. The first-order linear kinetics observed for the reaction, under moderate salt conditions (Method B), demonstrate that PNPase stalls at regions of secondary structure, however briefly, but can disengage from the "stalled" substrate and reassociate with a new free 3'-end. The finding that PNPase can participate in the dynamic processes of tRNA maturation and end-repair in vivo suggests that PNPase must be able to dissociate from precursor tRNA to allow final maturation by other processing enzymes (Li and Deutscher, 1994; L i and Deutscher, 1996; Reuven et al., 1997). Thus, we believe that the activity of PNPase, as assayed by Method B, accurately reflects the action of the enzyme in vivo. In contrast, under low salt conditions (Method A), PNPase appears to "stick" to the -123-R N A substrate upon stalling at a region of secondary structure, resulting in a slower rate of dissociation (step 4). It is also likely that the rate of binding (step 1) and phosphorolysis (step 2) are also decreased significantly under these relatively unphysiological conditions. In fact, 10-20-fold more enzyme was required to achieve equivalent levels of PNPase activity in Method A compared to that of Method B . A number of authors have demonstrated a requirement for PNPase in the degradation of highly-folded R N A fragments both in vivo and in vitro (Mackie, 1989; McLaren et al, 1991; Xu et al, 1993; Hajnsdorf et al, 1994; Xu and Cohen, 1995; Braun etal, 1996). Our results, however, show that PNPase stalls and dissociates 3' to the base of a stable Rho-independent terminator stem-loop in vitro suggesting that PNPase is more susceptible to inhibition by R N A secondary structure than previously imagined (McLaren etal, 1991). It is possible that PNPase-mediated decay through stable stem-loop structures requires additional factors such as RNase E, RhlB and/or poly(A) polymerase I (PAP I) as shown in Chapter 4. Although PNPase shows a dramatic preference for single-stranded 3'-ends, the enzyme can associate with RNAs which contain 3'-terminal secondary structures such as the rpsT/365 RNA. These observations are reminiscent of the protection of RNase II from thermal inactivation by the 77-nt product. Conceivably, the R N A associates with an RNA-binding site which is distinct from the active site (see Figure 1.5 in Chapter 1). In contrast to the situation for RNase II, however, RNAs which lack extended 3'-ends are able to compete effectively with single-stranded substrates. The significance of this competition is unknown since RNAs which lack single-stranded 3'-ends are not processively phosphorylyzed by PNPase. Previous results by Xu and Cohen (1995), however, have indicated that the 5'-end of an R N A may also influence PNPase-mediated decay of RNA1. In this -124-study, PNPase-mediated decay of RNA1 was enhanced significantly both in vivo and in vitro (Xu et al, 1993; Xu and Cohen, 1995) by the presence of a monophosphorylated 5'-end versus a triphosphate. It is important to note that the rpsT/365 R N A contains a triphosphorylated 5'-end unlike RNase E-cleaved RNA1 utilized in the previous studies. Although interactions with the double-stranded 3'-end of the rpsT/365 R N A cannot be excluded, it is possible that other portions of the rpsT/365 R N A may compete for PNPase binding. 3.4.4 A model for the control of mRNA degradation at the 3 '-end As previously discussed, 3'-stem-loop structures have been shown to protect upstream R N A sequences from digestion by 3'-exonucleases (Mott etal, 1985; McLaren et al, 1991). The observed protection of upstream sequences was originally attributed to the impeding of the processive activities of RNase II or PNPase by R N A structure. Our results, however, demonstrate that recombinant RNase II loses its apparent processivity 10 residues 3' to a region of strong R N A secondary structure, where it leaves the substrate rapidly and reassociates with a new free 3'-end. The data imply that stabilization of the Tn/0/IS70 antisense RNA-OUT (Pepe et al, 1994) and the stabilization of rpsO mRNA (Hajnsdorf et al, 1994) by RNase II are probably due to the removal of the 3'-overhang rather than to the formation of a stable RNA-RNase II complex which blocks access of PNPase to the 3'-end of mRNAs. However, it should be noted that dissociation and/or binding events could be retarded in vivo if free 3'-ends are limiting or if stem-loop binding proteins stabilize an RNase II-product complex (McLaren etal, 1991; Causton etal, 1994). It has been suggested that mRNAs with an immediate 3'-stem-loop structure, analagous to the 77-nt product, are poor substrates for PNPase (O'Hara et al, 1995). Our data clearly confirm this supposition since the 77-nt product, -125-produced by the action of the purified Rnb protein on t40B is not an efficient substrate. Conceivably, extension of the 3'-end by poly(A) polymerase I (PAP I) (O'Hara et al, 1995; X u et al, 1993; Xu and Cohen, 1995; Hajnsdorf et al, 1995) could provide a necessary single-stranded platform for PNPase to overcome the apparent indirect inhibition by RNase II. These observations suggest a possible model for the control of mRNA degradation at the 3-end. As RNase II encounters a region of secondary structure it stalls. If the structure is unstable the enzyme may advance through the stem-loop in the 3' to 5' direction. However, if the structure is a stable REP sequence or a Rho-independent terminator, RNase II will dissociate from the transcript before the duplex opens. We propose that loss of the single-stranded 3'-overhang which reduces the affinity of RNase II may also reduce the ability of the much larger PNPase to bind and degrade such RNAs. Addition of a new 3'-end by PAP I followed by the action of PNPase, which is less susceptible to inhibition by R N A secondary structure than RNase II (Mott et al, 1985; Guarneros and Portier, 1990; McLaren etal, 1991; Pepe etal, 1994), would be required for the degradation of strong REP and terminator sequences. Thus, a competition between removal of a 3'-overhanging sequence by RNase II and extension-degradation by PAP I and PNPase respectively would develop at the 3'-end of extended R N A secondary structures and may account for the heterogeneity in the 3'-ends of oligoadenylated RNA1 (Xu et al, 1993). 3.4.5 Utility of t40B as a substrate for 3'-exonucleolytic activity This partially duplexed R N A is an effective substrate for investigating the properties of RNase II and PNPase and offers at least four significant advantages over assays previously utilized for detecting exonucleolytic activity. First, the t40B transcript resembles natural mRNA substrates more -126-closely than the homopolymeric substrates utilized in traditional assays as it contains both 3' unpaired extensions of essentially random composition and a stable duplex mimicking stem-loop structures found in natural mRNAs. Secondly, the stalling of the enzyme at the duplexed region reflects the known behaviour of RNase II and PNPase on RNAs containing regions of extensive secondary structure (Mott et al, 1985; McLaren et al, 1991). Third, the formation of a stable degradative intermediate provides an internal control which distinguishes 3'-exonucleolytic activity from single-and double-strand specific endonucleases. Finally, the high specific activity of the synthetic transcript increases the sensitivity of the assay and allows for the detection of activity at low substrate concentrations (10"10-10"n M) closer to the physiological range. -127-CHAPTER 4 The action of the degradosome and the 3'-exonucleases on natural RNA substrates 4.1 Introduction As discussed previously in section 3.1, the degradation of highly structured R N A fragments is a particularly challenging problem for Escherichia coli. Until recently, it has been unclear how highly folded R N A fragments are degraded. Two possible solutions to this problem have emerged recently: R N A modification via polyadenylation or R N A unwinding. The product of the pcnB gene, poly(A) polymerase I (PAP I), has been implicated in the decay of RNA1, the anti-sense regulator of colEl-type plasmid D N A replication (Xu etal, 1993; Xu and Cohen, 1995). Addition of a single-stranded 3'-poly(A) tract to the 3'-terminus of RNA1 stimulates its complete degradation by PNPase in vitro (Xu and Cohen, 1995). Polyadenylation has also been reported to destabilize the mRNA encoding ribosomal protein S15 (Hajnsdorf et al, 1995; Haugel-Nielsen et al, 1996) and mRNA in general (O'Hara et al, 1995). These results have been rationalized by the hypothesis that polyadenylation facilitates decay by providing a single-stranded platform for the 3'-exonucleases (Cohen, 1995; Hajnsdorf etal, 1995; Haugel-Nielsen etal, 1996; Coburn and Mackie, 1996b). A second means of overcoming secondary structure is suggested by the finding that at least two enzymes of mRNA decay (RNase E and PNPase) copurify with RhlB, a putative DEAD-box R N A helicase in a high molecular weight complex referred to as the degradosome (Carpousis et al, 1994; Py et al, 1994; Py et al, 1996; Miczak et al, 1996). Moreover, 3-exonucleolytic degradation through a REP (repetitive extragenic palindrome) sequence in the malE-malF intragenic region -128-requires ATP hydrolysis and the presence of RhlB, but not PAP I (Py et al, 1996). The rpsTxriRNA, encoding ribosomal protein S20, is a small, monocistronic mRNA for which there is a tested secondary structure model (Mackie, 1992). This R N A serves as an excellent paradigm to investigate the relative roles of each component of the degradosome as well as the role of PAP I in the degradation of structured RNAs. The rpsT mRNA is a well characterized substrate for RNase E both in vivo and in vitro (Mackie, 1989; Mackie, 1991; Rapaport and Mackie, 1994). Cleavage of two slightly different synthetic rpsT transcripts (365 or 372 nucleotides) by either crude or purified RNase E generates a number of discrete products. In both cases, the most prominent RNase E-mediated cleavage product spans 147-residues, is coterminal with the substrate's 3'-end, and is identical to a degradative intermediate found in vivo (Mackie, 1989). Final degradation of this product is dependent on PNPase and accumulates in pnp mutants. In crude extracts, complete degradation of this otherwise stable 147-residue fragment is dependent on the presence of ATP, phosphate and PNPase activity (Coburn and Mackie, 1996b). We show here that the decay of the rpsTvriRNA can be reconstituted in vitro with a small number of purified enzymes. To our knowledge this is the first example of a resolved in vitro decay system, from any organism, which accurately describes the decay of an entire mRNA species. Most of these results have been published elsewhere (Coburn and Mackie, 1996b; Coburn and Mackie, 1998a). 4.2 Materials and methods 4.2.1 Enzymes and chemicals Purified recombinant RNase II (Fraction V, Figure 2.2; lane 8), purified PNPase (Fraction IV, Figure 2.5; lane 5), and purified degradosomes (Fraction V, Figure 2.8; lane 5) were prepared as -129-described in section 2.2.5. E. coli poly(A) polymerase I (PAP I) was purchased from Amersham-Pharmacia or Gibco-BRL. All other enzymes and chemicals were as described in sections 2.2.1 and 3.2.1. 4.2.2 Bacterial strains and plasmids The E. coli strains MG1693 (F- thyA715) and SK7988 (F- thyA715 X- pcnB A l:\m\m-kan kan*) were obtained from Dr. S. Kushner (University of Georgia, Athens, GA). Plasmid pCH77 (McLaren et al, 1991), containing 375 nucleotides of intergenic spacer sequence from the malE-malF mRNA under the control of a T7 promoter, was obtained from Dr. C. Higgins (Oxford University, Oxford, UK). The plasmid, pGM87 (rpsT coordinates 92-447; see Figure 4.1), pJG175 (rpsT coordinates 268-447; see Figure 4.1) and the related plasmids pGM87p(A) and pJG175p(A) were described previously in section 3.2.2. 4.2.3 Preparation of RNA substrates 4.2.3.1 Transcription Transcription of the rpsTRNAs and their derivatives was performed on linear D N A templates with T7 R N A polymerase in the presence of [a 3 2P]CTP as described in section 3.2.4. 5'-end-labeled RNAs were prepared as described in section 3.2.4. R N A transcripts were purified by extraction with phenol-chloroform-isoamyl alcohol (25:24:1) and two cycles of ethanol precipitation in the presence of 0.25 MNa-acetate as described in section 3.2.4. Linearized D N A templates and labeled rpsT'RNA substrates are listed in Table 4.1. -130-Table 4.1 Summary of plasmid-encoded rpsT transcripts. Plasmid Linearization RNA Substrates Coordinates1 Poly(A) Modified RNA Substrates2 pGM87 Dral rpsT/365 92-447 r/w77395-poly(A) Hindlll rpsT/333Aterm 92-415 /7w77363Aterm-poly(A) Sau3M rpsT/221 92-303 /7w77251-poly(A) pJG175 Dral rpsTI\19 268-447 >7w77209-poly(A) pGM87p(A) Xbal rps77395-poly(A) 92-447+poly(A)30 pJG175p(A) Xbal r/>.sT/209-poly(A) 268-447+pOly(A)30 1. rpsT coordinates from Mackie, 1992 and Figure 4.1 2. Poly(A) modified substrates were prepared by post-transcriptional enzymatic addition of 25-45 A residues. 4.2.3.2 Polyadenylation Polyadenylated rpsT R N A transcripts (30-40 pmoles) were prepared in a 60 pi reaction volume containing 25 mM HEPES-NaOH, pH 7.5, 5% (v/v) glycerol, 5mM MgCl 2 , 100 mMNFLCl , 60 mM KC1, ImM DTT, 0.02 mM EDTA, and 2mM ATP. The various poly(A) modified RNAs are summarized in Table 4.1. Purified E. coli poly(A) polymerase (0.3U) (Amersham-Pharmacia or Gibco-BRL) was added last and incubations were performed for 45 minutes at 37°C. Oligoadenylated transcripts were purified by extraction with phenol-chloroform-isoamyl alcohol (25:24:1) and collected by ethanol precipitation. The rpsT transcripts were extended by approximately 25-45 -131-nucleotides as determined by sizing on a sequencing gel. Alternatively, two of the polyadenylated substrates, 7ps773 95-poly(A) R N A and r/tt77209-poly(A) RNA, were transcribed directly from plasmids pGM87p(A) and pJG175p(A) which had been previously linearized with Xbal. A schematic representation of the various R N A substrates used throughout this study is shown in Figure 4.2. 4.2.4 Ribonuclease Assays Assays for PNPase and RNase II were assembled as previously described (Coburn and Mackie, 1996; see section 3.2.5 and section 3.2.6 in a 40 pi reaction volume containing 50-60 nM R N A substrate. Purified recombinant PNPase and RNase II were added to a final concentration specified in the Figure legends and incubations were performed at 37°C. Samples were withdrawn at various times and quenched in 3 volumes of loading buffer containing 90% formamide, 22 mM Tris, 22 mM boric acid, 0.5 mM EDTA, 0.1% xylene cyanol FF, and 0.1% bromophenol blue. The products were resolved on 5% or 6% polyacrylamide gels containing 8 M urea and visualized by autoradiography or with a Molecular Dynamics Phosphorlmager system. Al l assays were repeated a minimum of two to three times to ensure reproducibility. Degradation assays expressed as rates of substrate decay or product formation (see text) were calculated as the average rates from at least two experiments. Assays containing degradosomes and other ribonucleases and/or poly(A) polymerase were assembled in 40 pi reaction volumes containing 20 nM of labeled R N A in a buffer containing 25 mM HEPES-NaOH, pH 7.5, 5% (v/v) glycerol, 5 mM MgCl 2 , 100 mM NH 4C1, 0.02 mM E D T A and 60 m M K C l (Mackie, 1991). Enzymes and cofactors were added to the concentrations indicated in the Figure legends and incubations were performed at 30°C. The concentrations of the degradosome and -132-the 3'-exonucleases used in all experiments were limiting (e.g. -75-80% digestion in 15-30 minutes). PAP I was added to a final concentration that would add z 30 A M P residues per R N A in 15 minutes. The products of reactions were analyzed as described above. Al l assays were repeated a minimum of two to three times to ensure reproducibility as described above. 4.2.5 RNA extraction and analysis Fresh saturated cultures of MG1693 and SK7988 were diluted 400-fold into fresh media and grown with aeration at 30 °C until reaching an A^ of 0.2 (early log phase). L B growth medium was supplemented with 20 pg/ml thymidine and 50 ug/ml kanamycin for strain SK7988. Rifampicin, when used for half-life determination, was added to a final concentration of 200 pg/ml prior to sampling. In all cases, growth was arrested by transferring 5 ml of culture into 10 ml of 20 mM NaN 3 containing 350 pg/ml of chloramphenicol. Cells were harvested by centrifugation and R N A was prepared by the method of Dennis and Nomura (1975) with some minor modifications (Mackie, 1987; Mackie, 1989). RNAs were dissolved in DEPC-treated dH 2 0, quantified by A260 and stored at -70 °C until use. Approximately 5 pg of total E. coli R N A was boiled for 2-3 minutes in standard loading buffer containing 90% formamide and tracking dyes and separated electrophoretically through 6% polyacrylamide gels containing 8 M urea. The gels were soaked briefly in I X T B E (Tris-borate-E D T A ) to remove most of the urea and the R N A was transferred to Hybond-N nylon membranes (Amersham-Pharmacia) with a Mini Trans-Blot cell (Bio-Rad). Transfers were performed in 0.5X TBE for 1 hour at a constant current of 350 mA. The R N A was fixed to the Hybond membranes by U V photo-crosslinking (150 mJoules) with a GS Gene Linker (Bio-Rad). R N A blots were prehybridized at 55 °C for 60 minutes in annealing solution containing 50% formamide, 5X SSC -133-(standard saline citrate), 5XDenhardt's solution (Sambrook et al, 1989), 250 pg/ml yeast R N A and 60 ug/ml sheared calf thymus DNA. Membranes were subsequently hybridized in fresh annealing mix containing lx 107 cpm [32P]-labeled cRNA probe (49E; Mackie, 1989) overnight at 55 °C. R N A blots were washed four times in I X SSC, 0.1% SDS for 10 minutes and the hybridized R N A was visualized and quantitated with a Molecular Dynamics Phosphorlmager system. 4.3 Results 4.3.1 Sensitivity of the rpsT mRNAs to 3'-exonucleolytic digestion To determine whether the rpsT mRNA is a substrate for the 3-exonucleases, the purified enzymes were incubated with a 365-nt synthetic rpsT substrate, hereafter called rpsT/365 R N A (see Figure 4.1 and 4.2). During a 15 minute incubation, rpsT/365 R N A was resistant to attack by purified RNase II under conditions where single-stranded RNAs would be rapidly degraded (Figure 4.3 A). Even after prolonged incubations, up to 60 minutes, the rpsT/365 R N A remained resistant to digestion by RNase II (data not shown). Likewise, the rpsT/365 R N A remains intact and resistant to digestion by PNPase after 60 minutes of digestion (Figure 4.3B). A second substrate, rps773 3 3Aterm R N A (see Figure 4.2), lacks the Rho-independent terminator stem-loop structure (stem V U in Mackie, 1992; see Figure 4.1) and the adjacent stem-loop (stem VI in Mackie, 1992; see Figure 4.1) ending instead at the 3' base of stem VIb (residue U415). This R N A was incubated with the purified 3-exonucleases. The r/?s773 3 3Aterm R N A was largely resistant to digestion by purified RNase II (Figure 4.4A) as less than 10% of the substrate was degraded to a somewhat shorter, but stable, product denoted as " D " during a 15 minute digestion (Figure 4.4A). As will be shown later, intermediate D (see Figures 4.6 and 4.7) represents the product -134-Ilia U - A c - a a - X A D-A a 1Mb V 0 -A O-A D-A C-Q D 220 - * A 0-A a O-C - 2 4 0 c - a a - c 0-A A D O A a c o u . . . c A C a A D A D C O A a 320 - C A 0 - > i n D 0 » « n O A a C D c a c Z 1 ° c - a " ° i \ / D O 3 6 0 - a A a A , x o a - c I V c - a A A 1 7 0 c a III s : ; -«o ll \ c c 0-C A A C A - 0 A O c A A S a o c - o ' 3 3 0 c - a II Via D A 0 A 1 4 0 - A D 0 A A A c A A 0 A-150 C a . c a C a ' a c a c a 0 c D 0 A a C 0 A 160-C a c a a 0 200. C O A O A | A O - A - 3 7 0 0 - C 0 - A 2 9 0 ^ C C A - 0 a - c A | D - A 3 1 0 O-C C I A O A O 0 - C \ / l A O C - a 2 7 Q 2 8 0 C O 0 - A VI 3 5 0 - C - O 0 - C / ' ° " ~ 0 - C w 0 - C 340 a - c Y O-  0c a A A » 340 A I D - A C A A A A O C A C A A O C A C A O A A A C C O A O A A A A C A A A O C C a *bo 1 9 0 3oo o o o o o c a 5 0 D O A A 0 0 - A N 4 2 0 A O O 0 - A 4 1 0 - a - C 3 8 0 O 0 - A 0 - A » O A C A a o - A o - c a S O O O A C A C A A O A O A O A C C O O O O A A a O O O C C 0 0 0 - A 120 i i o 100 e 4 4 0 - a - c O-A O 0 -C 0 - A u 0 - C A C 5 ' - g a a u m c & c g g a a u C - a - 4 3 0 A - 0 0 -C C - O - 3 9 0 A a 4 0 0 - c - a A c a - c VII C D o o A A Vlb Figure 4.1 Model for the structure of the rpsT mRNA. Residues are numbered following the convention that residue 1 is transcribed from the natural PI promoter. The model shown corresponds to the more abundant P2 mRNA which would initiate at residue 92 and terminate at residue 447 to generate a 356 residue nascent transcript. Lowercase letters denote several additional residues transcribed from the vector. In the case of the rpsT/365 substrate, the vector-specified residues are: 5'-gggaauucc-3' (Mackie and Genereaux, 1993). The two most prominent RNase E cleavage sites are denoted by arrows. -135-Figure 4.2 The rpsT RNA substrates. A schematic representation of the rpsT gene encoding ribosomal protein S20 is shown. The coding sequence is denoted by a box whereas the major site of RNase E cleavage between residues 300/301 is marked with a large arrow. Various transcripts derived from the P2 ApsTmRNA are depicted below. Transcription of R N A substrates was directed from D N A templates (pGM87 or pJG175) which had been previously linearized with the appropriate restriction enzyme shown below the rpsT sequence (as described in section 4.2.3.1). The rpsT/365 RNA, encompasses the entire P2 mRNA terminating with the Rho-independent terminator stem-loop structure (stem VII; see Figure 4.1). The /ps773 3 3Aterm R N A terminates with an imperfect stem-loop structure (stem VTb; see Figure 4.1) followed by 4 unpaired residues ending at residue U415. This R N A lacks the Rho-independent terminator (stem VII; see Figure 4.1) and the majority of stem VI. The rpsTIHX R N A terminates with a weak stem-loop structure (stem IV) followed by 8 unpaired nucleotide residues ending at residue C303. The substrate, rpsT/179 RNA, encompasses residues 268-447 terminating with the Rho-independent terminator stem-loop (stem VII; see Figure 4.1). In addition to the substrates described above, additional R N A substrates containing 30 residue poly(A) tails were also synthesized as described in section 4.2.3.2 (r/«77395-poly(A), /7w77363Aterm-poly(A), rpsT/25l-poly(A), and r/«77209-poly(A) RNAs). The previously characterized 147-nt degradative intermediate (P0) generated by RNase E cleavage at residue 300 (Mackie, 1989) is shown below for comparison. -136-92 rpsT 300 447 3—£• 365 Sau3^/ Hindlll Dral Stem VII ^ 395 Stem VII *j Stem V l b S r 333 Stem V l b S ? 363 AAAA(A) n AAAA(A) n 221 Stem IV 1 251 Stem IV ft rpsT/365 (P2) rpsT/395-poly(A) rpsT/333Aterm rps77363Aterm-poly(A) rpsT/221 rps77251-poly(A) AAAA(A) n o Stem VII 179 rpsm 79 Stem VII 209 o rps7/209-poly(A) AAAA(A) n Stem VII O 147 147 (P0) -137-Figure 4.3 Digestion of the rpsTmRNA by exonucleases. The rpsT/365 R N A substrate (P2 mRNA) was incubated with 4 mU of purified RNase II (25 ng/ml) (panel A) or with 0.6 mU of purified PNPase (375 ng/ml) (panel B) at 37 °C as described in sections 3.2.5 and 3.2.6.1 respectively. Aliquots were removed at the times indicated (in minutes) and the reaction was quenched as described in section 4.2.4. The products were resolved by electrophoresis through a 6% polyacrylamide gel under denaturing conditions, then visualized by autoradiography and quantified using a Phosphorlmager system. The structure of the 3-terminus of the rpsT/365 R N A substrate (terminating 3' to the base of stem VII at residue 447; see Figure 4.1 and 4.2) is represented schematically in the margin to the right. -138-of exonucleolytic removal of 75 residues from the 3-terminus of r/?s773 3 3Aterm ending at the base of stem V (Mackie, 1992; see Figure 4.1 and 4.7). Similarly, the /7Js77333Aterm RNA was almost completely resistant to digestion by PNPase as less than 1% of the substrate disappeared over 15 minutes of incubation (Figure 4.4B). Although two stem-loop structures have been deleted from the 3'-terminus of the r/?s773 3 3Aterm RNA, this substrate remains largely resistant to digestion by the purified 3-exonucleases, mirroring the behaviour of the full-length rpsT/365 RNA (compare Figure 4.3 to 4.4, panels A and B). The resilience of these RNAs to exonucleolytic degradation is due, almost certainly, to the presence of stable 3'-terminal secondary structures which perturb or inhibit the binding of one or other of the exonucleases to these RNAs. During the course of our examination of the decay of the rpsT mRNA and its derivatives we observed that the resistance of some structured RNAs to digestion by PNPase may be overcome at elevated enzyme to substrate ratios. The r/?s773 3 3Aterm substrate which is almost completely resistant to digestion by PNPase under conditions where enzyme is limiting (Figure 4.4B), was incubated with 3 mU of PNPase activity (5-fold higher than that used in the experiment shown in Figure 4.4B). During 10 minutes of digestion, >80% of the r/?s773 3 3Aterm RNA substrate was degraded, whereas the /7w773 3 3Aterm RNA is completely stable in incubations which employ lower concentrations of PNPase (compare Figures 4.4B to 4.4C). The substrate, rpsT/221 RNA, terminating at residue C303, mimics the 220 nucleotide intermediate which would be generated by a single RNase E cleavage at residues 300/301 (see Figure 4.1 and 4.2). This "pre-cleaved" substrate was incubated with purified RNase II and approximately 75% of the substrate was degraded after 15 minutes of digestion (Figure 4.5 A). Three discrete intermediates (A-C) could be detected in this assay. Using 5-end-labeled substrates, the ends of these -139-F i g u r e 4.4 Digestion of rpsT/333Aterm RNA by exonucleases. The r/?s77333Aterm R N A which lacks the Rho-independent terminator stem-loop (stem VII in Mackie, 1992) and the adjacent stem (stem VI in Mackie, 1992) was incubated at 37 °C with 4 mU of purified RNase II (25 ng/ml) (panel A) or with 0.6 mU (375 ng/ml) (panel B) and 3 mU (1.25 pg/m) (panel C) of purified PNPase as described in sections 3.2.5 and 3.2.6.1 respectively. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The /ps773 3 3Aterm R N A substrate terminates at residue U415 with an imperfect stem-loop structure (stem VIb in Mackie, 1992; see Figure 4.1 and 4.2). The structure of the 3-terminus of the /*/?.s77333Aterm R N A is represented schematically in the margin to the right. Intermediate D, which terminates at residues 337-344 (31 to the base of stem V ; see Figure 4.1), represents the exonucleolytic removal of approximately 75 nucleotide residues from the 3'-end of the Ay>s773 3 3Aterm R N A (see text and Figure 4.7). - 1 4 0 -rps77333Aterm 0 0.5 1 2.5 5 7.5 10 1 rps773 3 3Aterm D -141-A 0 2.5 5 7.5 10 15 • t t t i t - * m m m B 0 2.5 5 7.5 10 15 rpsT/221 Figure 4.5 Digestion of rpsT/221 RNA by exonucleases. The rpsT/221 R N A which mimics the 220 nucleotide fragment that would be generated by a single cleavage by RNase E at residue 300/301 was incubated with 4 mU of purified RNase II (25 ng/ml) (panel A) or 0.6 mU of purified PNPase (375 ng/ml) (panel B) at 37 °C as described in sections 3.2.5 and 3.2.6.1 respectively. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The rpsTI22\ RNA substrate terminates with a weak stem-loop structure (stem IV in Mackie, 1992; see Figure 4.1) followed by 8 unpaired residues ending at residue C303. A schematic representation of the 3'-terminus of the rpsTI22\ R N A is shown in the margin to the right. The letters A - C in the margin represent clusters of degradation intermediates generated by RNase II digestion. The end points of intermediates A-C map to the following positions: A, residue 182-184 (3' to the base of stem II); B, residue 249-252 (3' to the base of stem Ilia); C, residue 266-270 (3' to the base of stem III) (see text and Figure 4.6 and 4.7). -142-Figure 4.6 Mapping of intermediates generated by RNase II digestion. 5' end-labelled RNAs 77^77333Aterm R N A (lanes 1-4) and rpsfllll R N A (lanes 5-8), prepared from pGM87 which had been linearized by HindUI or Sau2> AI respectively (refer to "Experimental Procedures") were treated as follows: lanes 1 and 5, partial alkaline digestion, pH 9.0 for 5 minutes at 100°C; lanes 2 and 6, partial digestion with 1.2 TJ of RNase T l in 8 M urea, 20 mM sodium citrate, pH 5.0 for 6 minutes at 50 °C; lanes 3 and 7, no treatment. End-labelled /ps773 3 3Aterm R N A (lane 4) and rpsT/221 R N A (lane 8) at a concentration of 100-200 nM were digested with 41 mU and 16.4 mU of purified RNase II respectively at 37°C. After 10 minutes of incubation portions of the incubation mixture were removed and diluted into 10-20 volumes of 0.25 M Na-acetate, containing 25 pg/ml of yeast RNA, 10 mM EDTA, extracted with phenol-chloroform-isoamyl alcohol (25:24:1) and precipitated with ethanol. The digestion products were separated electrophoretically through a 6% polyacrylamide sequencing gel containing 8 M urea then visualized by autoradiography. Selected G residues are indicated in the left margin while degradation intermediates generated by RNase II are indicated in the right margin as groups A-D (see Figure 4.7). -143-Group D Group C Group B Group A mm „ 12 3 4 5 6 7 8 -144-Figure 4.7 Degradative intermediates formed by exonucleoytic digestion of the rpsT RNAs. A schematic representation of the secondary structure model of the rpsT mRNA is shown at the top (Mackie, 1992). The degradative intermediates formed by stalling of RNase II and PNPase (see legend to Figure 4.6) are denoted by the letters A-D in the margin to the right. The size of the degradative intermediates are indicated (in nucleotides) and the position of the digestion end points are also shown. The major RNase E cleavage sites are also denoted by the arrows. -145-intermediates were subsequently mapped on a sequencing gel (Figure 4.6). A schematic diagram of the digestion end points is provided in Figure 4.7. Intermediates A-C could be detected as products of both r/?s77221RNA and 77?s773 3 3Aterm RNA, albeit in much lower abundance in digests of the latter. The most prominent 3'-ends cluster in several groups, designated A-D in the margin to Figure 4.6. Group A includes residues 182-194 which are several residues 3' to stem II (refer to Figures 4.6 and 4.7). Likewise group B (residues 249-252) maps to the 3'-base of stem IIIA, group C (266-270) maps 3' to stem III, and group D (337-344) 3' to stem V (Figures 4.6 and 4.7). The "pre-cleaved" rpsT/221 R N A was also incubated with purified PNPase (0.6 mU) for 15 minutes. The rpsTI22\ RNA, which terminates with a stem-loop of marginal stability (stem IV; zlG=-2.7 kcal/mol; Mackie, 1992; see Figure 4.1), was digested somewhat slowly compared to single-stranded substrates as approximately 50% of the substrate remained intact throughout the incubation (Figure 4.5B). A shortened intermediate, similar in size to group A (~ 100-nts) in RNase II digestions, was generated to very low levels (< 1% of the total substrate) during the assay. Intermediates (B-D) did not accumulate to detectable levels (compare Figures 4.5A and 4.5B). The sub stoichiometric recovery of discrete products is consistent with previous findings which suggest that PNPase is less susceptible to inhibition by R N A structure than RNase II (McLaren et al, 1991; Pepe et al, 1994). 4.3.2 Polyadenylation stimulates 3'-exonucleolytic digestion X u and Cohen (1995) have demonstrated that polyadenylation of RNA1, the anti-sense regulator of colEl-type plasmid replication, by poly(A) polymerase stimulates the activity of PNPase against RNA1 in vitro. In view of their data, we have tested whether prior polyadenylation of the rpsTKNAs could stimulate the activity of either RNase II or PNPase against these substrates. The -146-0 2.5 5 7.5 10 15 rps7/395-poly(A) rpsT/365 I o AAAAAAAAA B 0 1 2.5 5 7.5 10 15 rpsT/395-poly(A) rpsTI36S J Ji J AAAAAAAAA Figure 4.8 Digestion of rpsT/395-poly(A) RNA by exonucleases. Polyadenylated r/w77395-poly(A) R N A was prepared as described previously (see section 4.2.3.2). The substrate was incubated with 4 mU of purified RNase II (25 ng/ml) (panel A) or 0.6 mU of purified PNPase (375 ng/ml) (panel B) at 37°C as described as described in section 3.2.5 and 3.2.6.1 respectively. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The structures of the 3-termini of the r/?s773 95-poly(A) R N A substrate (containing a 30 residue poly(A) tail 3' to stem VTI at residue 447), the rpsT/365 intermediate (terminating 3' to the base of stem VII; residue 447), and intermediate "D" (terminating 3' to stem V at residues 337-344) are shown in the margin to the right (see Figure 4.7). -147-appropriate substrates were synthesized either by post-transcriptional enzymatic addition of approximately 25-45 A M P residues to the rpsT transcripts or by transcription of templates which were modified to contain 30 T residues on the template strand immediately 3' to the natural Rho-independent transcription terminator (see Table 4.1 in section 4.2.3.2 or Figure 4.2). Polyadenylated /"/?s77395-poly(A) R N A was incubated with either purified RNase U (Figure 4.8A) or PNPase (Figure 4.8B) for 15 minutes. In digestions of the former, the 30-nt poly(A) extension is gradually removed from r/?s773 95-poly(A) R N A to regenerate the 365-nucleotide rpsT transcript which is not significantly shortened further by RNase II (Figure 4.8 A). Similarly, poly(A) tails were digested from /7?s773 95-poly(A) R N A by PNPase to produce the rpsT/365 intermediate. Although the rpsT/365 intermediate remains relatively resistant to further digestion by PNPase, an additional degradative intermediate D, representing the exonucleolytic removal of about 107 residues from the 3'-end of the rpsTKNA distal to a stable stem-loop (Figure 4.7; Coburn and Mackie, 1996b), accumulates to relatively low levels during the assay (Figure 4.8B). Interestingly, no more than 15% of the polyadenylated substrate is completely degraded, unlike the situation reported for RNA1 (Xu and Cohen, 1995). Similar experiments were performed with two additional 3'-terminally truncated substrates. In contrast to the situation for the rpsT/365 RNA, addition of a single poly(A) tail to the otherwise resistant substrate, r/w773 3 3Aterm R N A (r/?s773 63Aterm-poly(A) RNA; see Figure 4.1), enhances its susceptibility to 3-exonucleolytic digestion. In the case of RNase II digestions, the rate of disappearance of /"/?s77363Aterm-poly(A) R N A is stimulated 4-5-fold compared to the non-adenylated r/?s773 3 3Aterm R N A (compare Figure 4.4A to 4.9A). In addition, degradative intermediate D accumulates linearly to > 4-fold greater levels than in the experiment of Figure 4.4A -148-0 2.5 5 7.5 10 15 • • • • • • • rps7/363Aterm-poly(A) • D J I AAAAAAAAAAAA B 0 2.5 5 7.5 10 15 4^ {ft 4 | j | m rpsr/363Aterm-poly(A) J J AAAAAAAAAAAA Figure 4.9 Digestion of rpsT/363Aterm-poly(A) by exonucleases. Polyadenylated r/«77363Aterm-poly(A) R N A was prepared as described previously (see section 4.2.3.2). The substrate was incubated with 4 mU of purified RNase II (25 ng/ml) (panel A) or 0.6 mU of purified PNPase (375 ng/ml) (panel B) at 37°C as described in section 3.2.5 and 3.2.6.1 respectively. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The structures of the 3'-termini of the r/w773 63Aterm-poly(A) RNA substrate (containing a 30 residue poly(A) tail 3' to stem VIb at residue U415) and the intermediate " D " (terminating 3' to the base of stem V at residues 337-344) are shown in the margin to the right (see Figure 4.6 and 4.7). -149-(see also Figures 4.9A and 4.9B). The accumulation of intermediate D is consistent with the stalling and dissociation of RNase II near the base of stem-loop V (Mackie, 1992; Figure 4.1). Conversely, addition of a poly(A) tail to /ps77333Aterm R N A makes it fully susceptible to destruction by purified PNPase. During 15 minutes of digestion, the oligoadenylated substrate was almost completely degraded (see Figure 4.9B) whereas the non-adenylated substrate remains intact and fully resistant to digestion by PNPase (Figure 4.4B). These results are not unlike the situation reported for RNA1 (Xu and Cohen, 1995). The resistance of the rpsTKNA to exonucleolytic digestion, therefore, is due largely, if not entirely to the presence of the Rho-independent terminator, a stem-loop containing 11-base pairs closed by a G C A A tetraloop (stem VII; Mackie, 1992; see Figure 4.1), which can impede both the binding and processive action of the 3-exonucleases. Finally /p.s77251-poly(A) RNA, a substrate which encompasses the 5-two-thirds of the rpsT mRNA, was prepared and incubated with the purified 3-exonucleases. During 15 minutes of digestion the rpsTHS l-poly(A) R N A was degraded by both RNase II and PNPase approximately 2-fold faster than the non-adenylated rpsT/221 RNA. Degradative intermediate A accumulates to relatively low levels in PNPase assays (see Figure 4.10B) whereas intermediates A, B and C accumulate in RNase II digestions as before (Figure 4.10A). The stimulation of degradation of Aps77251-poly(A) by the presence of the poly(A) tail is modest, but still significant. It demonstrates that even relatively weak stem-loops (stem IV; AG=-2.7 kcal/mol, Mackie, 1992; Figure 4.1) can perturb the binding of either exonuclease to 3'-ends (compare data in Figure 4.10 to those in Figure 4.5). In order to demonstrate whether, or not, the 3-exonucleases show a preference for poly(A) tails, competition experiments containing equimolar amounts of rpsTIHX R N A and Ayw77251-poly(A) R N A were performed. In assays containing RNase II (Figure 4.11 A) or PNPase (Figure 4.1 IB) both -150 -0 2.5 5 7.5 10 15 - r / 2 5 1 - p c l y ( A , JL AAAAAAAA c B B 0 2.5 5 7.5 10 15 • • * • * * # * » * * rpsT7251-poly(A) JL AAAAAAAA F i g u r e 4.10 Digestion of rpsT/251-poly(A) by exonucleases. Polyadenylated r/?.s77251-poly(A) R N A was prepared as described previously (see section 4.2.3.2). The substrate was incubated with 4 mU of purified RNase II (25 ng/ml) (panel A) or 0.6 mU of purified PNPase (375 ng/ml) (panel B) at 37°C as described as described in section 3.2.5 and 3.2.6.1 respectively. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The structure of the 3'-terminus of the rpsTHS l-poly(A) R N A substrate (containing a 30 residue poly(A) tail 3' to stem IV at residue C303; see Figure 4.1) is shown in the margin to the right. Intermediates A-C are denoted in the margin to the right. The end points of intermediates A-C map to the following positions: A, residue 182-184 (3' to stem II); B, residue 249-252 (3' to stem Ilia); C, residue 266-270 (3' to stem III) (see text and Figure 4.6 and 4.7). The asterisk denotes the position of unadenylated/deadenylated substrate. -151-0 2.5 5 7.5 10 15 l a i t i t rps7/251-poly(A) rpsT/221 c B B 0 2.5 5 7.5 10 15 m m rps77251-poly(A) rpsT/221 Figure 4.11 Competition between rpsT/221 RNA and rpsT/251-poly(A) RNA for exonucleases. Polyadenylated /7Js77221-poly(A) R N A was prepared as described previously (see section 4.2.3.2). Equimolar concentrations (50 nM of each substrate) of rpsT/221 and /7?s77251-poly(A) RNA substrates were incubated with 4 mU of purified RNase II (25 ng/ml) (panel A) or 0.6 mU of purified PNPase (375 ng/ml) (panel B) at 37°C as described in section 3.2.5 and 3.2.6.1 respectively. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The substrates and intermediates A-C are indicated in the margin to the right. The end points of intermediates A-C map to the following positions: A, residue 182-184 (3' to stem II); B, residue 249-252 (3' to stem Ilia); C, residue 266-270 (3' to stem III) (see text and Figure 4.6 and 4.7). -152-the rpsTI22\ R N A and 7ps77251-poly(A) RNAs are gradually degraded over 15 minutes of digestion. However, the rate of disappearance of the oligoadenylated /"/>s77251-poly(A) R N A is approximately 2-fold greater than the rate of disappearance of the non-adenylated substrate. Thus, as shown previously (compare the data shown in Figures 4.9 and 4.10 with 4.11), a single round of polyadenylation can result in a modest increase in the susceptibility of rpsT/221 R N A to degradation by either exonuclease. The finding that non-adenylated rpsT/221 R N A is digested during competition experiments demonstrates that polyadenylation is not a prerequisite for its decay unlike the situation for the r/w773 3 3Aterm R N A (compare Figures 4.3 and 4.9) or RNA1 (Xu and Cohen, 1995). 4.3.3 Northern analysis of the rpsT mRNAs Polyadenylation of bacterial mRNAs is believed to promote their destabilization by facilitating 3'-exonucleolytic attack (Cohen, 1995; Xu and Cohen, 1995; Coburn and Mackie, 1996b). In apparent contradiction, however, the preceeding data show that the intact rpsT mRNA is relatively resistant to purified 3-exonucleases in vitro, even when polyadenylated. In order to determine whether the pcnB gene product plays a role in the decay of the rpsT mRNA in vivo, we examined both steady-state levels and half-lives of the two rpsT transcripts by northern analysis. In the former (Figure 4.12), equivalent amounts of total R N A extracted from either strain MG1693 (wild type) (lane 3) or strain SK7988 (ApcnB) (lane 4) were compared. The steady-state levels of the rpsT mRNAs, derived from the PI and P2 promoters (447- and 356-residues respectively), were essentially identical between the two strains; however, the levels of the P 0 R N A fragment, which represents the 3'-terminal 147-nt fragment generated by RNase E cleavage at residues 300/301/302 (Mackie, 1991) varied markedly. In strain SK7988 (ApcnB), it accumulated to levels approximately 50-fold greater -153-CQ C o 1632 517 396 298 220 i 2 2 PI P2 o 154 - $ - p 1 2 3 4 Figure 4.12 Effect ofpcnB on the steady-state levels of the rpsT mRNAs. Cultures of MG1693 and SK7988 were grown at 30 °C to mid-exponential phase. Total R N A was extracted and separated by electrophoresis through a 6% polyacrylamide gel containing 8 M urea. The rpsT mRNAs were visualized by Northern blotting of 5 pg of total RNA as described previously (Rapaport and Mackie, 1994). Lane 1 contains denatured end-labeled DNA fragments from pBR322 digested with Hinfl (M); lane 2, empty; lane 3, R N A isolated from strain MG1693 (wt); lane 4, R N A isolated from strain SK7988 (ApcnB). PI and P2 denote the 356- and 447-nt mRNAs respectively derived from tandem promoters for the rpsT gene, while P D (Mackie, 1989) represents the major 147-nt degradative intermediate generated by RNase E cleavage at residue 300/301. -154-than those in the wild-type strain where it was undetectable (Figure 4.12, compare lanes 3 and 4). The half-lives of the PI and P2 rpsT mRNAs in strain MG1693 (wild-type) and strain SK7988 (ApcnB) were determined by northern blotting of RNAs extracted from exponentially growing cultures after addition of rifampicin. Interestingly, the half-lives of the rpsT mRNAs extracted from strain SK7988 (ApcnB) were similar to those in the wild-type strain (PI: 90 s, P2: 118 s) (Mackie, 1989). The half-life of the P 0 fragment (Mackie, 1989) from strain SK7988 (ApcnB) was approximately 24.5 minutes whereas the half-life of this fragment from strain MG1693 (wild-type) could not be determined reliably since it did not accumulate to levels significantly above background. These data show that only the decay of the structured 3'-terminal 147 residues of the rpsT mRNA but not the decay of the full-length species mRNA or 5-terminal fragments depends on PAP I activity. 4.3.4 Sensitivity of the rpsT/365 RNA and derived products to degradosomes Since purified PNPase is unable to degrade the rpsT mRNA efficiently, even when the latter is oligoadenylated, we have investigated the ability of the degradosome (Carpousis et al. 1994; Py et al, 1994; Py etal, 1996; Miczak et al, 1996; Blum etal, 1997), to cleave the rpsT/365 RNA. In the presence or absence (not shown) of ATP and phosphate, degradosomes readily attack rpsT/365 R N A to generate the expected stable 147-nt RNase E cleavage product which accumulates throughout the assay (Mackie, 1991; Mackie and Genereaux, 1993) (Figure 4.13). An additional stable 106-nt fragment, corresponding to a cleavage by RNase E at a previously characterized minor site (residue 340/341) within the 3'-one-third of the rpsTmRNA (Mackie and Genereaux, 1993) also accumulates during the assay. All other endonucleolytic fragments disappear completely after 20 minutes of incubation. The spectrum of cleavage products does not differ significantly from that -155-0 2.5 5 7.5 10 15 20 30 45 f f f f f -HIV < * » • " " * » 4 * 4 » . * 4 » « * » 4 » « » 147 . w * « # * • • 106 Figure 4.13 Digestion of the rpsT mRNA by degradosomes. The substrate rpsT/365 R N A was incubated with purified degradosomes (final concentration of 6.25 pg/ml) at 30 °C in the presence of 3mM ATP and 10 mM potassium phosphate as described in section 4.2.4. Aliquots removed at the times indicated (in minutes) and were processed as described in the legend to Figure 4.2. The numbers in the right margins represent the size of the 365-nt substrate and the two major decay intermediates in nucleotides. -156-obtained from crude extracts, from partially purified RNase E or from electrophoretically purified Rne protein (Mackie, 1991; Cormack etal, 1993). We also tested the ability of the degradosomes to digest a second substrate, rpsTI\19 R N A (see Figure 4.1), encompassing residues 268-447. The rpsTI\19 R N A is known to be a poor substrate for RNase E (Mackie et al, 1997), presumably due to the formation of a 5'-terminal stem-loop. This substrate shares the same 3-terminus with the 147-nt RNase E product and should, therefore, be as resistant to exonucleolytic activity as the 147-nt product. During 30 minutes of digestion in the presence of ATP and phosphate, RNase E activity in the degradosome generated the 147-nt fragment to relatively low levels, <, 5% of theoretical yield, as anticipated (Figure 4.14B; compare 0 and 30 minute time points). Since rpsTI\19 R N A is largely resistant to exonucleolytic digestion by PNPase and to cleavage by RNase E, this substrate truly mimics the 147-nt fragment. Previous results have shown that addition of ATP and phosphate results in the rapid elongation and subsequent disappearance of the otherwise stable 147-nt R N A fragment generated by RNase E cleavage of rpsT/365 R N A in crude extracts (Coburn and Mackie, 1996b). In order to separate the ATP requirement for polyadenylation from that of ATP-dependent unwinding of R N A secondary structure by putative R N A helicases such as RhlB, we incubated purified degradosomes with a "pre-adenylated" substrate, rJps77209-poly(A) R N A which contains 30 terminal A residues (Figures 4.1 and 4.14A). In the presence of ATP and phosphate, the 30 residue poly(A) tail was rapidly removed to regenerate a product essentially identical to rpsTI\19 R N A (indicated in the margin under Figure 4.14). This decay intermediate was a poor substrate for purified degradosomes as > 75% remained intact after 30 minutes of incubation (Figure 4.14 A; compare the 30 minute time point to the 0 time). This result is similar to that obtained with purified PNPase and rpsT/365 R N A -157-Figure 4.14 Effect of a single round of polyadenylation on the decay of the 3'-end of the rpsT mRNA. The following substrates r/w77209-poly(A) R N A (residues 268-447 containing a 30-nt poly(A) tail 3' to the base of stem VII at residue 447) (panel A) and rpsTI\19 R N A (residues 268-447) (panel B) were incubated with purified degradosomes (final concentration of 6.25 pg/ml) at 30 °C in the presence of 3 mM ATP and 10 mM potassium phosphate as described in section 4.2.4. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The numbers in the right margins represent the size of the substrates and decay intermediates in nucleotides. The substrate and intermediates are depicted schematically in the margin below. -158-(Figure 4.2B) but differs significantly from the situation reported for a fragment oimalE-malF mRNA. The latter could be degraded effectively by purified degradosomes in the presence of ATP (Py etal, 1996). 4.3.5 Continuous polyadenylation and 3'-exonucleolytic decay Since a single addition of a poly(A) tail to rpsT/365 R N A or to rpsTI\19 R N A failed to stimulate the degradation of these RNAs significantly, we investigated whether sustained oligoadenylation by commercially available poly(A) polymerase (PAP I) might potentiate the activity of the degradosomes and/or PNPase against such otherwise refractory RNAs. The r/?s77209-poly(A) R N A substrate was incubated with degradosomes (Figure 4.15 A) or purified PNPase (Figure 4.15B) in the presence of the ATP analog, ATPyS, and 10 mM phosphate. This analog contains a natural a-P phosphoanhydride linkage, and can serve as a substrate for PAP I which releases thiopyrophosphate. The data in Figure 4.15A show that both /7?s77209-poly(A) R N A and the rpsTI\19 intermediate were completely degraded by purified degradosomes in the presence of PAP I even though the concentration of degradosomes was one-sixth of what was used in Figure 4.14A. Moreover, compared to assays which contain degradosomes and ATP alone, addition of PAP I stimulated the action of the degradosomes against the rpsTI\19 intermediate by 5-fold as judged by the rate of disappearance of the rpsTI\19 intermediate (compare Figure 4.14A and 4.15A). This experiment was repeated, but purified PNPase was substituted for degradosomes. In this case as well, purified PAP I was also able to stimulate the activity of purified PNPase against the rpsTI\19 intermediate by 7-fold compared to assays which contained PNPase alone (see Figure 4.15B). In contrast to the result shown in Figure 4.14A, where rpsTI\19 R N A is stable, > 95% of rpsTI\19 -159-0 1 2.5 5 7.5 10 15 20 30 o AAAAAAAAA rpsT/209-poly(A) rpsT/179 1 B 0 1 2.5 5 7.5 10 15 20 30 • l l * ! o AAAAAAAAA rpsT/209-poly(A) rpsT/179 1 Figure 4.15 Effect of continuous polyadenylation on decay of the 3'-end of the rpsT mRNA. In panel A, substrate rpsT209-po\y(A) R N A (residues 268-447 containing a 30-nt poly(A) tail 3' to the base of stem VII at residue 447) was incubated with either purified degradosomes (1 pg/ml) or in panel B with 0.2 mU of purified PNPase (125 ng/ml). Digestions also contained 40 mU of PAP I activity and were performed at 30 °C in the presence of 3 mM ATPyS and 10 mM potassium phosphate as described in section 4.2.4. Aliquots removed at the times indicated (in minutes) were processed as described under the legend to Figure 4.3. The 3'-termini and size (in nucleotides) of substrates and intermediates are depicted schematically in the margin to the right. -160-R N A was completely degraded by either purified PNPase or PNPase complexed in the degradosome (compare the 30 minute time points in Figure 4.14A with Figures 4.15A and 4.15B). All reactions were carried out in the presence of 10 mM phosphate and either ATP or ATPyS, both of which support oligoadenylation of R N A substrates by PAP I. Previous results, however, showed that ATPyS could not substitute for ATP in the decay of malE-malFREP sequences by the degradosome (Py etal., 1996), presumably because ATPyS cannot serve as a substrate for ATPases. In contrast, substitution of ATP with the non-hydrolyzable analog in assays using purified degradosomes or purified PNPase as the source of 3-exonuclease activity had no effect on the rate of disappearance ofrpsT/179 RNA. The data in Figure 4.15 show that degradation of the 3'-terminus of the rpsT mRNAs depends on PAP I, PNPase, ATP and phosphate and is independent of P-y hydrolysis of ATP and thus ATPase activity. The effect of continuous polyadenylation on the degradation of the full-length rpsT/365 R N A by the degradosome was also examined. Substrate R N A was incubated with degradosomes and PAP I in the absence of ATP. Samples were removed at 5 minute intervals. Both the 147-nt and 106-nt products of RNase E cleavage accumulate under these conditions (Figure 4.16A and 4.16B, 0-25 minute time points). After addition of ATP at T=25 minutes, both the 147-nt R N A and 106-nt fragments completely disappear within 7.5 minutes (compare Figure 4.16A, 25-32.5 minute time points to Figure 4.13). Slightly different results were obtained when ATPyS was substituted for ATP at T=25 minutes (Figure 4.16B). Elongation of the 147-nt product was much more pronounced in digestions supplemented with the non-hydrolyzable analog, as demonstrated by the more dramatic smearing of the 147-nt product. Moreover, only 50% of the 147-nt product disappeared within 7.5 minutes of -161-ATP A * 0 5 10 15 20 25 26 27.5 30 32.5 35 f t * •365 ATPyS B 1 t * 0 5 10 15 20 25 26 27.5 30 32.5 35 •147 •106 f t ! •365 J • » » - * -147 +poly(A) -<-147 Figure 4.16 Effect of continuous polyadenylation on decay of the rpsT mRNA. The substrate rpsT/365 RNA was incubated with purified degradosomes (6.25 pg/ml) and 100 mU of PAP I activity at 30 °C in the presence of 10 mM potassium phosphate. At T=25 minutes ATP (panel A) or ATPyS (panel B) was added to a final concentration of 3 mM. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The numbers in the right margins represent the size of substrates and decay intermediates in nucleotides. -162-addition ATPyS, whereas both the 147-nt and 106-nt products were rapidly elongated and completely degraded within the same time interval after addition of ATP (compare Figure 4.16B). Interestingly, in digestions supplemented with the ATPyS, the full-length rpsT/365 R N A continues to be digested by the degradosome whereas addition of ATP appears to stabilize significantly a fraction of the rpsT/365 R N A (compare Figures 4.16A and 4.16B).The significance of this observation is not clear. We also tested whether purified PNPase could substitute for degradosomes in the decay of the rpsT/365 RNA. The data in Figure 4.17A show that complete degradation of rpsT/365 R N A can also be accomplished in the absence of degradosomes by PNPase and PAP I in the presence of ATP and 10 mM phosphate. In contrast, rpsT/365 R N A is stable in the presence of either PNPase (Figure 4.2B) or PAP I (Figure 4.17C) alone, although it is clearly elongated in the latter. Moreover, in the presence of both PNPase and PAP I but in the absence of ATP, rpsT/365 R N A is also completely stable (Figure 4.17B). Taken together, these results demonstrate the crucial importance of continuous polyadenylation in overcoming the resistance of structured RNAs to degradation. Genetic data have suggested that RNase II and PNPase are functionally redundant (Donovan and Kushner, 1986). Accordingly, we have examined the effect of continuous polyadenylation on RNase II-mediated decay of Aps77209-poly(A) RNA. The data in Figure 4.18A show that poly(A) tails (30 residues) are rapidly hydrolyzed by RNase U to produce a slightly elongated rpsTI\19 decay intermediate. This intermediate maintains approximately 10-15 adenylate residues, as estimated by its mobility on polyacrylamide gels, and is almost fully resistant to digestion over 30 minutes (Figure 4.18A). Thus, RNase II cannot substitute for PNPase, even in the presence of continuous polyadenylation. Normally, RNase II activity is in considerable excess over that of PNPase in vivo (Deutscher and Reuven, 1991). Therefore, we have examined whether RNase II would affect PAP -163-0 10 20 30 40 50 60 75 90 rps77365 B C 0 30 60 90 0 30 60 90 • • • • +rPsmes f • f • - rpsm^m rps 7/36 5 Figure 4.17 Effect of continuous polyadenylation on 3'-decay of the rpsT mRNA. In panel A, the substrate rpsT/365 R N A was incubated with 0.4 mU of purified PNPase (250 ng/ml). Digestions also contained 100 mU of PAP I activity and were performed at 30 °C in the presence of 3 mM ATP and 10 mM potassium phosphate as described in section 4.2.4. Similarly, in panel B, rpsT/365 R N A was incubated as described above with PNPase and PAP I but in the absence of added ATP. In panel C, rpsT/365 R N A was incubated with 100 mU of PAP I activity. This incubation also contained 3 mM ATP and 10 mM sodium phosphate as described above. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The position of the various substrates and products are shown in the margin to the right. -164-I-dependent PNPase-mediated degradation of r/w77209-poly(A) RNA. Addition of purified RNase II to assays containing PAP I and degradosomes resulted in a 5-fold stabilization of the rpsTI\19 intermediate (compare Figure 4.18B (+RNase II) to Figure 4.15A (-RNase II)). Moreover, rpsTI\19 R N A is stable in the presence of RNase II (Figure 4.18B) whereas it disappears completely in Figure 4.15 A in the absence of RNase II (compare the 30 minute time points in Figure 4.15A and 4.18B). The slightly elongated rpsTI\19 intermediate produced by RNase II digestion of 77?s77209-poly(A) R N A (compare Figures 4.18A and 4.18B) does not accumulate to significant levels in the presence of degradosomes. Presumably this is due to non-processive removal of the 10-15 remaining 3'-terminal nucleotides by PNPase (Littauer and Soreq, 1982). Thus, paradoxically, RNase II inhibits PNPase-mediated degradation of rpsTI\19 by removing the essential poly(A) tail. This provides a clear explanation for the phenomenon of RNase II-mediated, stem-loop-dependent, stabilization of the rpsO mRNA (Hajnsdorf etal, 1994) and of RNA-OUT, the anti-sense regulator ofTnJO/ISJO transposition (Pepe et al, 1994), in agreement with our earlier hypothesis (Coburn and Mackie, 1996a). For completeness, we have performed additional assays containing degradosomes and PAP I and various combinations of RNase II and ATP. First, in the presence of the degradosome, PAP I and RNase II, but in the absence of ATP, the rpsT/365 R N A is rapidly digested to generate the 147-nt and 106-nt products which accumulate as expected (shown in Figure 4.19A). The spectrum of decay intermediates does not vary significantly from that observed in Figure 4.13. In a second experiment, ther/«773 65 R N A was incubated with the degradosome, PAP I and ATP alone (shown in Figure 4.19C). In the absence of RNase II activity, the rpsT/365 R N A gradually disappears over 40 minutes of digestion. Interestingly, degradative intermediates, such as the 147-nt product, are -165-0 0.5 1 2.5 5 7.5 10 15 30 t f^i rpsT/209-poly(A) • rpsT/179 1 1 A A A A A A A A A B 0 1 2.5 5 7.5 10 15 20 30 Jk m •<— rpsT/209-poly(A) • fc « * • » • * • • • • • P o A A A A A A A A A Figure 4.18 Effect of RNase II on the decay of the 3'-end of the rpsT mRNA. In panel A, the /ps77209-poly(A) RNA (residues 268-447 containing a 30-nt poly(A) tail 3' to the base of stem VII at residue 447) was incubated with 8 mU of purified RNase II (50 ng/ml) and 100 mU of PAP I activity. In panel B, the 7ps77209-poly(A) RNA was incubated with purified degradosomes (1 pg/ml), 40 mU of PAP I activity and 60 mU of RNase II (375 ng/ml). Digestions were performed at 30 °C in the presence of 3 mM ATP and 10 mM potassium phosphate as described in section 4.2.4. Aliquots removed at the times indicated (in minutes) were processed as described under the legend to Figure 4.3. The 3'-termini of the substrate and intermediates are depicted schematically in the margin under panel B. -166-Figure 4.19 Effect of continuous polyadenylation on decay of the rpsT mRNA in the presence of RNase II. In panel A, the substrate rpsT/365 R N A was incubated with purified degradosomes (1 pg/ml), 164 mU of RNase II (1 pg/ml) and 30 mU of PAP I activity at 30 °C as described in section 4.2.4. In panel B, the rpsT/365 R N A substrate was incubated with degradosomes, RNase II and PAP I as above. This reaction, however, was supplemented with 3 mM ATP. In panel C, the substrate rpsT/365 R N A was incubated with degradosomes and PAPI in the presence of 3 mM ATP as above. A l l incubations contained 10 mM sodium phosphate. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The numbers in the right margins represent the size of substrates and decay intermediates in nucleotides. -167-0 5 10 15 20 25 30 40 + Degradosomes + RNase II + PAP I -ATP « # « * Complete S . ^"365 147 106 m .* n ii w n M B 0 5 10 15 20 25 30 40 -*-365 ?!! I I I # • • «( 147 106 365 Complete - RNase II H (BP ^ «j -168-relatively transient and do not accumulate to significant levels throughout the incubation suggesting that endonucleolytic fragments are preferentially degraded over the full-length rpsT/365 RNA. Finally, the rpsT/365 R N A was incubated with the degradosome, PAP I, RNase II and ATP. Surprisingly, in the experiment shown in Figure 4.19B, the rpsT/365 substrate was digested approximately 2-3-fold faster than in incubations which lacked RNase II (compare Figure 4.19B and 4.19C). The apparent stimulation of endonucleolytic activity may be attributed to the exonucleolytic removal of cleavage products and/or the trimming of poly(A) tails by RNase II which compete with the 5'-end for binding to one or more components of the degradosome. Under these conditions, the 147-nt product accumulates quickly and remains constant for 10-15 minutes of digestion and then disappears gradually during the following 30 minutes of digestion (Figure 4.19B). As in the case for the rpsTI\19 intermediate, the 147-nt product is slightly more resistant to digestion by the degradosome in the presence of RNase II (compare Figures 4.18B and 4.19B). 4.3.6 Continuous polyadenylation and degradation of the malE-malF REP sequence Previous work in C. Higgins' laboratory (Oxford) has shown that the decay of a fragment of the malE-malFmRNA, containing a REP sequence (shown in Figure 4.20), requires ATP hydrolysis and the degradosome but not PAP I (Py et al, 1996). In order to ensure that our degradosome preparation was comparable to theirs, we have re-investigated the decay of the malE-malF REP sequence under the previously described conditions (McLaren et al, 1991; Py et al, 1996). In the experiment shown in Figure 4.21A, the 375-nt REP sequence was incubated with purified degradosomes in the absence of ATP as described in section 4.2.4. During 60 minutes of digestion the REP R N A was gradually digested to generate a previously characterized intermediate, referred -169-GAA C A c c A G A U A A A—U C — G A — U G A U—A U A C — G C — G G—C G — C U C G A C — G G — c G — C C — G C — G G—C A - U G—C C U C U C — G - C A G A—U G — C U—A G—U U—A U—A C — G A C - G C C — G G—U A C U—G U—A A—U G—C G — C U - A C — G C — G C — G G—C A A A U G — C C U A C G U A G 0 - C AGUU C — G u AG = -21.7kcaVmol AG = -22.8 kcaVmol G — C U—A G G—C C — G G—U G — C C — G G — C U—A A—U G—C G A C — G C — G G _ C A A A U — * G U U AG = -56 kcal/mol Figure 4.20 The REP stem-loop structure. The sequences depicted are from the malE-malF intergenic region. In this region there are two copies of the REP sequence, inverted in orientation to one another. Thus, they can potentially form a single large hairpin structure or, alternatively, the two REP sequences can form two smaller structures (adapted from Fliggins et al, 1993). -170-Figure 4.21 Digestion of the malEREPRNA by the degradosome. The malE REP R N A substrate (McLaren et al, 1991) was incubated with degradosomes (5 pg/ml) alone (panel A) or in the presence of ATP (panel B) or ATPyS (panel C) under the conditions previous described in Py et al. (1996). All incubations were supplemented with 10 mM sodium phosphate and the reactions were performed at 30 °C as described in section 4.2.4. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The positions of the substrate, malE REP, and two previously characterized decay intermediates *, and RSR (REP Stabilized RNA) are indicated in the margin to the right of panel A (see Figure 4.22 for a schematic diagram of their structures). The intermediates * and RSR represent the exonucleolytic trimming of approximately 43 and 77 nucleotide residues respectively from the malE REP RNA. -171-ATP 0 2.5 5 10 15 20 30 45 60 ^ # • • • • — * « — malE REP — RSR B 0 2.5 5 10 15 20 30 45 60 + ATP \ £ m 4 — ™IE REP • * » * » - « * — RSR + ATPyS 0 2.5 5 10 20 30 45 60 — ma/£REP — RSR -172-to as the RSR (REP Stabilized RNA) (McLaren et al, 1991; see schematic in the margin to Figure 4.22). This intermediate is stable and remains fully resistant to digestion for 60 minutes (Figure 4.20, lanes 1-9). A second, faint intermediate, denoted by an asterisk (*) in Py et al. (1996), also appears transiently within the first 5 minutes of the assay (Figure 4.21 A). Although the structure of the REP R N A has yet to be determined empirically, the formation of the degradative intermediates are presumed to be the result of PNPase stalling at the base of stable stem-loop structures, which are inferred from the R N A sequence (McLaren et al, 1991; Py et al, 1996). Upon addition of ATP to assays containing degradosomes, the malE REP R N A was rapidly digested to generate the * and RSR intermediates as before (Figure 4.2 IB). In contrast to the result shown in Figure 4.21 A, where the RSR intermediate is stable, £ 80% of the RSR intermediate was completely degraded by the degradosome in the presence of ATP (compare the 60 minute time points in Figure 4.21A with Figures 4.2 IB). Assays containing the malE REP R N A and degradosomes were also performed in presence of the non-hydrolyzable analog, ATPyS (Figure 4.21C). After 60 minutes of incubation, the RSR intermediate was largely stable, whereas in incubations containing ATP, the RSR intermediate was almost completely degraded after 30 minutes (compare Figures 4.21A and 4.2IB). Thus, ATPyS cannot substitute for ATP in the decay of the RSR intermediate by the degradosome confirming that ATP hydrolysis is required for complete degradation of the malE REP R N A as previously reported (Py et al, 1996). Since PNPase and PAP I are sufficient for complete decay of the rpsT mRNA in vitro, we have tested whether this is also true for the malE-malF R N A under conditions of continuous polyadenylation. In the experiment shown in Figure 4.22 (lanes 10-11), the REP R N A substrate was incubated with PNPase and PAP I in the absence of ATP. During 90 minutes of digestion the REP -173-0 10 20 30 40 50 60 75 90 0 90 lis? -Figure 4.22 Digestion of the malE REP RNA by PNPase. The 375-nt malE-malF REP RNA substrate (McLaren et al, 1991) was incubated with 0.4 mU of purified PNPase (275 ng/ml) in the presence of 100 mU PAP I activity. Incubations contained 10 mM sodium phosphate and were performed in the presence (lanes 1-9) or absence (lanes 10-11) of 3 mM ATP at 30 °C as described in section 4.2.4. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The substrate, malE REP, and three previously characterized decay intermediates **, * and RSR (REP Stabilized RNA) (Py et al, 1996) are indicated in the margin to the right of panel A. The formation of these degradative intermediates, 14-nts (**), 43-nts (*), and 77-nts (RSR) from the 3'-end of the malE REP RNA, is likely the result of exonucleolytic stalling 3' to the base of stable stem-loop structures which can be inferred from the nucleotide sequence. A schematic representation of the malE REP RNA and the decay intermediates is shown in the margin below. -174-R N A substrate was shortened by approximately 3-5 nucleotides, as estimated by the mobility of the product on plyacrylamide gels. This previously characterized intermediate (Py et al, 1996) (denoted as ** in Figure 4.22) remains completely stable for 90 minutes of digestion (Figure 4.22, lanes 10-11). In the presence of PAP I and ATP, PNPase is able to digest the REP R N A to generate the intermediate * (McLaren et al, 1991; Py et al, 1996) (Figure 4.22, lanes 1-9). This degradative intermediate remains almost fully resistant to further shortening, as less than 1% of the total REP R N A is converted to the shorter RSR intermediate. Unlike the situation for the rpsT mRNA, degradation of the malE R N A by PNPase cannot be potentiated by the presence of continuous polyadenylation (compare Figure 4.17A with Figure 4.22, lanes 1-9). We have also examined the decay of the malE REP R N A by the degradosome under our assay conditions. Our assay buffer contains more M g 2 + (5 mM versus 1 mM) and significantly more monovalent cation (160 mM versus 50 mM) than that employed by Py et al. (1996). Presumably the elevated levels of cations, which stabilize R N A stem-loops, are responsible for the enhanced sensitivity of PNPase to secondary structure observed in Figure 2.22. In the presence or absence of ATP, the degradosome shortens the R N A substrate by 3-5 nucleotides, similar to that observed for digestions containing PNPase (compare the ** intermediate in Figures 4.22 with lanes 10-11 in Figure 4.23 A and 4.23B). Upon addition of ATP to assays containing the degradosome and PAP I, > 95% of the malEKNA. substrate disappeared within 60 minutes of incubation, whereas in the absence of ATP the shortened REP R N A is completely stable after 90 minutes (Figure 4.23 A, compare lanes 7 with 11) . Degradative intermediates * and RSR did not accumulate to high levels under these conditions (McLaren et al, 1991; Py et al, 1996). Moreover, smearing of the intermediates, which may be indicative of successive rounds of polyadenylation/deadenylation, is notable throughout the -175-A + A T P - A T P 0 10 20 30 40 50 60 75 90 0 90 „ malE REP • t l t l f ! f « RSR 1 2 3 4 5 6 7 8 9 10 11 + A T P y S - PAPI D 0 10 20 30 40 50 60 75 90 0 90 malE REP • • • • • • • • • ( , 6 7 8 9 10 11 Figure 4.23 Effect of continuous polyadenylation on the decay of the malE REP RNA by the degradosome. The malE REP R N A substrate (McLaren et al., 1991) was incubated with degradosomes (1.25 ug/ml) and 50 mU of PAP I activity in the presence (panel A, lanes 1-9) or absence of ATP (panel A, lanes 10-11). In panel B (lanes 1-9), the malE REP R N A was incubated with degradosomes and PAPI as above in the presence of 3 mM ATPyS. In panel B (lanes 10-11), the malE REP R N A was incubated with degradosomes alone in the presence of 3 mM ATP. Al l incubations were supplemented with 10 mM sodium phosphate and the reactions were performed at 30 °C as described in section 4.2.4. Aliquots removed at the times indicated (in minutes) were processed as described in the legend to Figure 4.3. The positions of the substrate, malEKEP, and three previously characterized decay intermediates **, * and RSR (REP Stabilized RNA) are indicated in the margin to the right of panel A (see Figure 4.22 for a schematic diagram of their structures). -176-assay. Since the data of Py et al. (1996) and the data shown in Figure 4.21 indicate that ATP hydrolysis is required for complete decay of the malE REP RNA, we have substituted the non-hydrolyzable analog ATPyS for ATP in digestions containing the degradosome and PAP I (Figure 4.23B, lanes 1-9). After 90 minutes of incubation, only 50% of the REP R N A was digested to the shortened * intermediate. In contrast, the REP R N A was almost undetectable after 60 minutes in assays containing ATP (compare lanes 1-9 in Figures 4.23A and 4.23B). Thus, unlike the situation for structured portions of the rpsTmRNA, the non-hydrolyzable analog ATPyS does not readily substitute for ATP in the decay of the malE-malFREP sequence. 4.4 Discussion 4.4.1 Reconstitution of mRNA degradation with purified components A major outcome of this chapter is a demonstration that degradation of the rpsT'mRNA can be reconstituted by a small number of purified enzymes. As shown schematically in Figure 4.24 A and 4.24B (steps 2A and 3), degradation of the 5'-two-thirds of the rpsTmRNA can be accomplished by the action of RNase E and either RNase II or PNPase (Coburn and Mackie, 1996b; this work). Although modification of 3'-ends by PAP I is not required at this stage, it does significantly stimulate the decay of R N A fragments derived from the 5'-two-thirds of rpsTmRNA (Coburn and Mackie, 1996b). In contrast, fragments originating from the 3'-one-third of the r ^ J m R N A exhibit additional requirements. PAP I, PNPase, ATP and phosphate alone are necessary and sufficient to catalyze the degradation of the smaller intermediates which are generated by RNase E cleavage and which retain the stable Rho-independent terminator stem-loop structure (Figure 4.24A and 4.24B, steps 4A, 5-7). Data obtained in vivo (Mackie, 1989; this work) are fully consistent with this minimal set of enzymes. -177-Figure 4.24 Degradation of the rpsTmRNA. A schematic representation of the rpsT gene encoding the mRNA for ribosomal protein S20 is shown in panel A (top). The coding sequence is depicted by the large box (residues 133-395). The transcription start sites PI (residue 1) and P2 (residue 92) are denoted by bent arrows while the Rho-independent terminator, composed of 11 paired residues closed by a G C A A tetraloop, is represented by a stem-loop structure. RNase E cleavage sites are denoted by arrows. In panel A, solid horizontal arrows depict regional susceptibility of the rpsT mRNA to the indicated enzymes. Dotted horizontal arrows denote regions of the rpsTmRNA which are likely susceptible to the indicated enzymes. In panel B, a model for the degradation of the rpsT mRNA is shown. A potential mechanism for PAP I-PNPase dependent degradation of structured RNAs is shown in steps 4-7 (see text). -178-r + r P s T I I 1 1 UUG UAAI 1 1 1 92 133 300 395 447 RNase E + RNase II / PNPase •< • — > • PNPase + PAP I 5' 2A RNase E 5" I RNase II or PNPase PAP I (?) Mononucleotides + Short Oligos 1A PAP I 2B RNase II 1B 5'-• I A RNase II or 4A PAP I | | p N R a s e 4 B 5 PNPase •_8 \ r - ! AAAAAAAAAA 5 PAP I 5 PNPase AAAAAAAAAA Mononucleotides + Short Oligos AAAAAAAAAA -179-It is likely, however, that a significant quantity of intracellular PNPase exists in a complex with RNase E and other proteins (Carpousis et al, 1994; Py et al, 1994; Py et al, 1996; Miczak et al, 1996; Blum et al, 1997). Thus under physiological conditions PNPase activity may be supplied by degradosomes rather than by free PNPase. Our data are completely consistent with this likelihood as purified degradosomes can fully substitute for PNPase on an activity basis (compare Figure 4.15A with4.15B). Interestingly, the full-length ApsrrnRNA can also be degraded in vitro, albeit inefficiently, by PAP I, PNPase, ATP and phosphate alone. This observation shows that other components of the degradosome (e.g. RNase E and RhlB) do not necessarily contribute to its 3'-decay. The contribution of this minor pathway to decay of the rpsT mRNA in vivo remains unknown. Our data reveal, however, an unexpected role for RNase II in suppressing such poly(A)-dependent 3-exonucleolytic decay mediated by PNPase (Figure 4.24B; steps I A and IB; see also Hajnsdorf et al, 1996). RNase II will hydrolyze 3'-single-stranded extensions including poly(A) tails distal to stable stem-loops. RNAs terminated by a sufficiently stable duplex are resistant to the action of the 3'-exonucleases (Mott et al, 1985; Coburn and Mackie, 1996a, 1996b). As a consequence, RNase II activity will convert RNAs from a form susceptible to PNPase, and possibly to other 3-exonucleases, to one which is relatively resistant. We believe that this property of RNase II is highly relevant to mRNA decay in vivo. Indeed, RNase II activity has been reported to stabilize two different RNAs in vivo (Pepe et al, 1994; Hajnsdorf et al, 1994) consistent with the preceding explanation. Additionally, in the absence of poly(A), degradosomes should be excluded from initiating 3-exonucleolytic attack dependent on PNPase and would be channelled into the authentic 5'-dependent endonucleolytic pathway involving RNase E (Figure 4.24B; steps 2A and 2B). The latter is believed to be the pathway -180-followed by most mRNAs (reviewed in Nierlich and Murikawa, 1996). In accordance with the preceeding hypothesis, some preliminary evidence from the expression of Desulfovibrio vulgaris eye mRNA in E. coli, suggests that many of the degradative intermediates which accumulate in strains deficient for RNase II also accumulate in strains deficient for RNase E activity (Cruz et al, 1996). 4.4.2 Ispoly(A) polymerase I required to initiate mRNA decay? Several authors have suggested that polyadenylation facilitates 3-exonucleolytic degradation of highly-folded R N A fragments (Cohen, 1995; Xu and Cohen, 1995; Coburn and Mackie, 1996b; Hajnsdorf et al, 1996), while others have implied that polyadenylation may trigger decay of full-length mRNAs (O'Hara et al., 1995; Ingle and Kushner, 1996). Two lines of evidence suggest that degradation of the rpsT mRNA is initiated by events which are independent of 3-oligoadenylation, presumably involving one or more cleavages catalyzed by RNase E. First, cleavage of the rpsT mRNA by RNase E in vitro is independent of ATP, PAP I or the presence of a poly(A) tail (Mackie, 1991; Cormack etal, 1993; Coburn and Mackie, 1996b). Second, and more importantly, both the in vivo half-lives and the steady-state levels of the two rpsT mRNA species do not appear to be altered significantly in strains deficient for PAP I. In addition, there are several strong arguments against polyadenylation playing a general role in initiating the decay of most mRNAs. First, the pcnB gene is dispensable (Masters et al, 1993), although the presence of an additional poly(A) polymerase (PAP II) (Kalapos et al, 1994) may compensate. Second, degradation of many long polycistronic mRNAs is initiated prior to 3'-end formation (Chow and Dennis, 1994). Third, among the handful of RNAs which has been examined thus far, only modest effects on the half-lives of full-length RNAs have been observed in strains -181-deficient for PAP I. The greatest impact of eliminating PAP I activity is observed only in strains which are also deficient for a number of additional ribonucleases (O'Hara et al, 1995; Hajnsdorf et al, 1995). Finally, the requirement for continuous polyadenylation in the degradation of highly-folded, distal portions of the rpsT mRNA argues against a role for poly(A) in initiating decay as suggested by others (O'Hara et al, 1995; Ingle and Kushner, 1996). Nonetheless, RNAs which are not substrates for RNase E (e.g. RNA-OUT; Pepe et al, 1994) may depend on PAP I and 3'-exonucleases for initiation of their decay. 4.4.3 Hierarchies of RNA secondary structure, poly (A) polymerase I, and 3 '-exonucleolytic decay R N A secondary structure, an important determinant of mRNA stability, is known to impede the processive action of RNase II and to a lesser extent PNPase (Mott et al, 1985; McLaren et al, 1991). In addition, RNAs which lack >12 unpaired 3'-terminal residues cannot be engaged in a processive mode of decay by RNase II or PNPase (Coburn and Mackie, 1996a; Littauer and Soreq, 1982), although some "nibbling" at the 3'-end may occur, possibly by a non-processive mechanism previously described for PNPase (Littauer and Soreq, 1982). We hypothesize that there is a hierarchy in the susceptibility of structured 3'-termini to attack by either RNase II (Coburn and Mackie, 1996b) or PNPase. First, unpaired 3-ends preceded by weak secondary structure (e.g. residues 5' to residue 300 in the rpsTmRNA) are directly accessible to both 3-exonucleases without prior modification (see Figure 4.24A and 4.24B; steps 2A and 3). Second, more stable stem-loop structures, including those found at the 3'-termini of R N A I (Xu and Cohen, 1995) and /7w773 3 3Aterm, reduce the rate of exonucleolytic attack but can be degraded once modified by a single round of polyadenylation. Third, very stable structures, such as the rpsT terminator stem-loop, are completely resistant to attack by -182-RNase II and, surprisingly, by PNPase (either free or complexed) even in the presence of a poly(A) tail (e.g. rpsT/209-uo\y(A)). These RNAs require the continuous action of PAP I and PNPase for their complete decay (see Figure 4.24A and 4.24B; steps 4-7). Finally, the most stable structures such as those within the malE R N A fragment (Py et al, 1996) require the degradosome, ATP hydrolysis, and PAP I activity. We believe that the relative thermodynamic stability of stem-loop structures located at the immediate 3'-terminus defines the number polyadenylation-PNPase cycles required for complete decay of a structured R N A and accounts for the differences observed between R N A I, malE REP sequences and the rpsT mRNA. In our model, upon stalling 3' to the base of a stable duplexed RNA, PNPase will dissociate releasing the R N A substrate similar to the situation reported for RNase II (Coburn and Mackie, 1996a; 1996b) (Figure 4.24B; 4B). The more stable the stem-loop, the greater the likelihood the enzyme will dissociate before digesting through the stem-loop. Oligoadenylation of 3-ends catalyzed by the action of PAP I, and possibly PAP II, is then required to permit re-engagement and processive phosphorolysis by PNPase (Figure 4.24B; step 4A). Localized melting at the base of the stem-loop structure ("breathing") would eventually allow PNPase to invade a stem-loop structure prior to its dissociation (Figure 4.24B, step 5). Incremental removal of residues from the 3'-(descending) strand of a duplex involving successive rounds of polyadenylation, breathing and processive phosphorolysis would progressively weaken the Rho-independent terminator below a threshold stability resulting in its eventual destruction (Figure 4.24B, steps 5- 7). The finding of heterogeneous sites of polyadenylation in several RNAs including the Ipp (major outer membrane lipoprotein) mRNA (Cao and Sarkar, 1992b), R N A I (Xu et al, 1993) and the rpsO mRNA (Haugel-Nielsen et al, 1996) is strong evidence in support of reiterative events at 3-ends. -183-Figure 4.25 Alternative mechanisms ofpoly(A)-mediated decay of highly folded RNAs. The Rho-independent terminator stem-loop of the rpsT mRNA (stem VII in Mackie, 1992; Figure 4.1) is denoted as intermediate I in both panels A and B. Addition of a poly(A) tail to the stem-loop (intermediate II) catalyzed by PAP I (or PAP II) permits the binding of PNPase (or RNase II) to an otherwise inaccessible 3'-end and may lead to thermodynamic destabilization of the stem-loop by one of two models. In the first (panel A), unimolecular rearrangement driven by base pairing between the poly(A) tail and the A - U rich base of the stem (intermediate III), may lead to a significant loss in base stacking resulting in the destruction of the stem-loop by PNPase. In the second model (panel B), addition of a poly(A) tail to the stem-loop (intermediate II) results in the co-localization of PAP I and PNPase. Interaction between PAP I (denoted by the black circle) and PNPase (denoted by the "pac-man") may result in a conformational change which enhances the ability of PNPase to processively phosphorylyze structured RNAs. -184-A G C — G G — C C — G C — G C — G G * U A — U A — U A — U A — U • G A - U U I 1B RNase II PNPase -< • P A P I -+ ATP 1A C — G G — C C — G C — G C — G G * U A — U A — U A — U A — U • G A — U U A A A A A A A A A A A A A 4 Rearrangement • c G C — G G — C C — G C — G C _ G A A A G U U U U U U U * A I 11 I 11 I A A A A A A A A A A A A A 3 PNPase • A A A A • G A A A A A A A A A A A A A A A A A A III C — G G — C C — G C — G C ~ G . A A G U U U U U U U A " A A 1 111 I I I A A A A A A A A A . . A A A A A • G A A A A A A A A A A A A A A A A A ^ A A A " 0 -C — G G — C C — G C — G C — G G U U U U A A A A G A 0 IV B C A C * C * G A G G C — G C — G C — G G - C PAP I G ~ c PNPase/PAP I + ATP C - G Complex C - G C — G C — G G » U G U U U U A — U A A — U A A - U „ W A - U W > k A • G A — U U G A — U U A A A A A A A A A A A A A A A A > 1 G A II Ml C — G C — G C — G > G » U A — U A — U A — U A — U -185-Additional models that describe alternative mechanisms by which poly(A) addition might overcome the resistance of structured RNAs to PNPase degradation have also been proposed (Coburn and Mackie, 1998b). In the first model, shown in Figure 4.25A, a poly(A) tail invades the base of the terminator stem, pairing with U residues in the descending arm of the stem, forming an alternative, lower-energy structure (Figure 4.25A, step 2). This would melt part of the stem and permit PNPase to advance through the structure via incremental removal of residues from the 3'-(descending) strand of an otherwise stable duplex (Figure 4.25A, step 4). A second, hypothetical model implies that PAP I not only provides a poly(A) tail for binding to the 3'-end but also induces a conformational change in PNPase which enhances the processive activity of the enzyme (Figure 4.25B). An example of this kind of conformational change has been described previously for the assembly of the NusA-dependent anti-termination complex which increases the processivity of R N A polymerase allowing it to transcribe through Rho-dependent and intrinsic terminators (Richardson and Greenblatt, 1996). The decay pathway for the rpsT mRNA requires the continuous activity of poly(A) polymerase to "edit" the 3'-ends of structured R N A fragments so that PNPase can bind otherwise resistant RNAs and enter a processive mode of degradation (Xu et al, 1993; X u and Cohen, 1995; this work). The ratio of ATP hydrolyzed per molecule of R N A degraded cannot yet be calculated. It is undoubtedly considerable, particularly in view of the likelihood that RNase II, the more active of the two major 3-exonucleases involved in mRNA decay, constantly removes adenylate residues from the 3'-termini of mRNAs. Conceivably, RNase E in the degradosome may also contribute to 3' shortening (Huang et al, 1998). In contrast, degradation of a portion of the malE mRNA exhibits additional requirements. The free-energy of ATP hydrolysis, which is presumed to drive the -186-unwinding of REP sequences, is required such that PNPase in the degradosome can gain access to the otherwise buried residues (Py et al, 1996). Although the degradation of both RNAs share a requirement for ATP and PNPase, the decay of the rpsT mRNA is supported by the non-hydrolyzable analog, ATPyS, and can be accomplished in the absence of the other degradosomal components arguing against an absolute requirement for an ATPase and, by implication, against RhlB. 4.4.4 malE REP RNA decay revisited Over the course of investigating the decay of the malE REP sequence in vitro, several discrepancies have appeared in the literature which merit some comment. The original data of McLaren et al. (1991) demonstrated that REP sequences could stabilize upstream R N A by temporarily impeding the action of PNPase. In fact, the rather weak stabilization that these structures provided in vitro led to the hypothesis that 3-ends must be protected from exonucleolytic attack in vivo by stem-loop binding factors (McLaren et al., 1991; Causton et al., 1994). Later, an apparently conflicting result was reported by the same group (Py et al, 1996). In this case, the REP stabilized R N A remained intact and resistant to further decay by PNPase (Py et al, 1996). One explanation for the differences between these two reports may lie in the quality of the preparation of purified PNPase (McLaren et al, 1991; Py et al, 1996). The presence of a contaminating non-specific nuclease, such as RNase I, might account for the gradual digestion of the REP stabilized R N A intermediate observed by McLaren et al. (1991). This would give the erroneous appearance that PNPase had digested through the REP stem-loop. A second, plausible explanation proposes that degradation through a REP sequence is an artifact of performing digestions at high enzyme to substrate ratios. Since there is a dynamic equilibrium between the folded and partially -187-unfolded states of an R N A stem-loop, when the concentration of enzyme is in excess over that of the substrate RNA, the probability of a random collision between an unfolded 3'-end and PNPase is greatly enhanced. Thus, the natural resistance of structured RNAs to 3'-exonucleolytic decay may be overcome at high enzyme concentrations. Our observations of the decay of the /7?s773 3 3Aterm R N A with high concentrations of PNPase fully support this hypothesis. In agreement with the findings of Py et al. (1996), we show that decay of the malE REP sequence is dependent on the degradosome and ATP. However, our results demonstrate an additional requirement for continuous polyadenylation in the decay of the malE REP R N A under moderate salt conditions. It is likely that this requirement was dismissed by the previous group for several reasons. First, in the preceding study, the effect of only a single round of polyadenylation on malE REP decay was examined. Our results clearly show that a single poly(A) tail does not significantly enhance the susceptibility of structured portions of the rpsT mRNA to digestion. Second, the studies by Py et al. used equimolar (or molar excess) quantities of degradosome to REP RNA. As discussed for PNPase-mediated decay (see above), at high concentrations of degradosomes the requirement for a poly(A) tail in the decay of the malE REP R N A may be by-passed. Finally, compared to the low salt conditions utilized in previous studies (McLaren et al., 1991; Py et al., 1996), the relatively high concentration of M g 2 + (5 mM) and lower temperatures (30 ° C) used in continuous polyadenylation assays serve to significantly stabilize R N A substrates against exonucleolytic digestion. This is most likely the result of an increase in the stabilization of R N A structure which can impede both the binding and processive action of the 3-exonucleases. Stabilization of a stem-loop, particularly at the 3'-terminus, would further strengthen the requirement for a poly(A) tail to initiate exonucleolytic decay. -188-CHAPTER 5 Conclusions and Perspectives 5.1 Summary The work described in this thesis represents a significant advance in our understanding of mRNA decay in Escherichia coli. First, and foremost, a major outcome of this work is a demonstration that degradation of the /psTmRNA can be reconstituted by a small number of purified enzymes. To my knowledge this is the first example of a resolved in vitro decay system which accurately describes the degradation of a model mRNA. Secondly, this work has sharpened significantly the relative roles of the ribonucleases and poly(A) polymerase I in the process of mRNA decay. In addition, we have demonstrated that a hierarchy of R N A secondary structures determines the availability of 3'-ends to exonucleolytic attack and determines the number of polyadenylation/phosphorolysis cycles required for complete decay of otherwise stable structured RNAs. 5.2 Models for investigating mRNA decay As stated previously (see section 1.6), early models of mRNA decay postulated that degradation of an mRNA is initiated by one or more endonucleolytic cleavages followed by digestion of the remaining oligonucleotide fragments to mononucleotides by the 3-exonucleases (Belasco and Higgins, 1988; Higgins etal, 1992; Higgins et al, 1993) . Over the last several years, many aspects of these models have proven valid; however, the rediscovery of polyadenylation in prokaryotes and -189-the finding that RNase E and PNPase are tightly associated have rendered previous models of mRNA decay somewhat obsolete. Moreover, the finding that both the purified Rne/Ams protein and the degradosome show a marked preference for monophosphorylated 5-termini begins to explain the 5'-3' directionality associated with the decay of most mRNAs (G. A. Mackie, manuscript submitted). Several new models which provide greater predictive power and which incorporate new information regarding the properties of the enzymes and the role of R N A secondary structure in directing the action of the ribonucleases are discussed in section 5.2 below. It should be noted that many of the ideas appearing in this section were derived from discussions between George Mackie, myself and other members of the laboratory. These models and the associated paradigms of mRNA decay appear in a recent review (Coburn and Mackie, 1998b). 5.2.1 The 5'-tethering model A model applicable to most RNAs is shown in Figure 5.1 (Coburn and Mackie, 1998b). This model, proposed by George Mackie, was based, in part, on the mechanism of action of Rep D N A helicase (Lohman, 1993). The key predictions and novel features of the 5-tethering model are first, that the Rne/Ams protein is present as a dimer (or higher oligomer) in the degradosome, second, that RNA-binding and catalytic sites are used alternatively and third, that products are preferentially bound to or retained by the degradosome. A similar, but simpler looping model has also been suggested by David Bechhofer (Bechhofer, 1993). Likewise, Alifano et al. have proposed a 5'-dependent nuclease model (Alifano et al, 1994). We hypothesize that the initiating step in mRNA decay is the binding of RNase E as a multienzyme complex to an mRNA at a site (site 1 in Figure 5.1) at or near the 5'-end. Site 1 probably -190-Figure 5.1 The 5'-tethering model The degradosome is a multi-enzyme complex likely containing two subunits of Rne/Ams constituting RNase E (ellipsoid), two trimers of PNPase (shaded circles), two dimers of RhlB (black circles) and two dimers of enolase (not shown; see the text). A high affinity RNA-binding domain (hatched boxes) and putative phosphate binding pockets (small black circles) are indicated on the two Rne subunits (labeled I and U). Open circles represent strong binding of 5'-monophosphorylated RNAs by the phosphate binding pocket. -191-A c t 1. Binding to 5" end of mRNA at site 1. 2. Optional cleavage at site 1 followed by binding to site 2 on mRNA by subunit II. 3. Cleavage at site 2 and 3' exo attack on 5' segment B 1. Recognition of 5' end. 2. Looping out of mRNA and binding to site 1. 3.Cleavage at site 1 followed by release or 3' exo digestion of looped out RNA. c 3 Mononucleotides + short oligos 4.Di ^ fn isplacement of 5' end rom subunit I. 5'-monophosphorytated fragment retained. c «9 5. Binding to site 3 by subunit I. II •as 6. Repeat steps 3-5. " n . M> yOH 'it ATP-PAP I J I RNase II PPi . . . " M O H A A A A A A . . A E3* ^ v * A . «* 3 ^ A A A * ^ >^IIIMHHQ Recycle RNase E + PNPase 7.3' T exo degradation of segment. Mononucleotides + short oligos -192-consists of at least 3 unpaired residues (Bouvet and Belasco, 1992) and is not necessarily cleaved by RNase E although RNA1 is cleaved effectively at a single site 5 residues from its 5'-end (Tomcsanyi and Apirion, 1985; Lin-Chao and Cohen, 1991). In vivo, this step is inhibited by secondary structure at or near the 5'-end (Chen et al, 1991; Emory et al, 1992; Bouvet and Belasco, 1992; Hansen et al, 1994), as shown in Figure 5.2 and possibly by competition from translating ribosomes, depending on the proximity of the 5'-end to the ribosome binding site, as shown in Figure 5.3. The mRNA substrate would be tethered to the degradosome, either by the R-RBD (Taraseviciene et al, 1995; McDowall and Cohen, 1996) or more likely, by the SI domain (Bycroft et al, 1997) in the N -terminus of subunit I of homodimeric Rne/Ams (see Figure 1.1). Subunit II of RNase E would then contact the mRNA at a more distal site (site 2) without significant dissociation. This generates apparent processivity which is important for "all or none" models of mRNA decay (reviewed in Nierlich and Murakawa, 1996). Site 2 would be single-stranded and A + U-rich, typical of RNase E cleavage sites. Binding of the tethered mRNA to the second or subsequent internal cleavage sites would be strongly affected by both translating ribosomes and R N A secondary structure by steric hindrance or by folding a susceptible site into a double-stranded R N A stem-loop (Mackie and Genereaux, 1993). In the model of Figure 5.1, step 1 is clearly reversible. In fact, steps 1-3 could be reversible if the first cleavage occurs at site 2, rather than site 1. A cleavage at site 2 may lead to functional inactivation of the message if ribosome loading becomes impaired. Moreover, this initial cleavage event produces an R N A fragment which contains a 5'-monophosphorylated end and commits the remainder of the mRNA to chemical decay (Bechhofer, 1993). Both purified Rne/Ams protein and the degradosome have a striking preference for monophosphorylated over triphosphorylated 5-ends in vitro (G. A. Mackie, manuscript -193-Figure 5.2 Effect of RNA secondary structure on mRNA decay. In panel A, RNAs which contain fewer than three unpaired residues at the 5'-end are resistant to attack by the degradosome. In panel B , RNase UI cleavage in the stem-loop, shown by the arrows, may initiate the decay of many RNAs by eliminating structural barriers to the degradosome. In panel C, the degradosome skips past stem-loop structures to internal cleavage sites once bound to a free 5'-end on an mRNA. Thus, positioning rather than the presence of stem-loop structures determines whether a potential site will be cleaved by the RNase E component of the degradosome. All symbols are as described in the legend to Figure 5.1. -194-submitted). This might reflect the ability of monophosphorylated ends to enter a phosphate binding pocket on the Rne subunit or it may simply be the result of a reduction in charge repulsion between RNase E in the degradosome and an mRNA substrate. The kinetically preferred binding of 5'-monophosphorylated termini by the Rne subunits could account for much of the apparent processivity of mRNA decay and is consistent with the dependence of the decay of RNA1 in vivo on prior RNase E cleavage (Lin-Chao and Cohen, 1991; Xu er al., 1993; Bechhofer, 1993; Alifano et al., 1994). Whether this preference is sufficient to account for the "all or none" behaviour that characterizes the decay of many mRNAs is unclear. The retention of 3'-products shown in the 5'-tethering model of Figure 5.1 postulates a physical association between the degradosome and 3'-products of RNase E cleavage reinforcing the kinetic preference of RNase E for monophosphorylated RNAs and enjoys some experimental support (Kaberdin etal., 1996). Some evidence also suggests that PNPase may also prefer 5'-monophosphorylated substrates (Xu and Cohen, 1995) The 5'-tethering model or its variants can explain how 5'-stem-loop structures confer stability by mediating resistance to RNase E (Chen et al., 1991; Emory et al., 1992; Bouvet and Belasco, 1992; Hansen etal., 1994). Figure 5.2A shows that such structures occlude the initial site of binding (and possibly cleavage). The combination of a terminal triphosphate and R N A duplex would reduce the ability of RNase E to bind to intact RNAs. Without this initial recognition, internal sites cannot be attacked. Figure 5.2B illustrates the facilitating role played by RNase III in initiating the decay of mRNAs: it removes terminal stem-loops which block the initial binding of the degradosome to an mRNA and could reveal otherwise cryptic RNase E sites. Conversely, "pseudo-processing" of polycistronic mRNAs (shown in Figure 5.4) may contribute to differential stability by re-positioning internal stem-loop structures at the 5'-terminus, resulting in the subsequent stabilization of the -195-Figure 5.3 Effect of translation on mRNA decay. In panel A, translating ribosomes compete with the initiating events of mRNA decay (i.e. 5'-end recognition by Rne/Ams subunit I within the degradosome). In panel B, translating ribosomes compete with secondary events in mRNA decay (i.e. the acquisition of internal cleavage sites by the degradosome). Potential RNase E sites on the mRNA are denoted by black boxes while the Shine-Dalgarno sequence (SD) is denoted by an open box. Al l other symbols are described in the legend to Figure 5.1. -196-downstream mRNA. The processing of mRNAs, which can be referred to as interrupted tethering, may be the basis of RNase P-dependent stabilization of the his operon from Salmonella typhimurium (Alifano et al, 1994) or the stabilization of the 3'-end of the papBA bicistronic mRNA. (Baga et al, 1988). Each cleavage catalyzed by RNase E would generate a new free 3'-end. As drawn in step 3 of Figure 5.1, the model suggests that the action of a 3'-exonuclease removes the 5'-cleavage product from its binding site on Rne/Ams allowing subunit I to engage a more distal site on the mRNA substrate. Alternatively, the cleavage event itself could liberate the 5'-cleavage product from the RNA-binding site of subunit I. The new 3'-end on the 5'-cleavage product could be attacked directly by PNPase in the degradosome or by RNase II, in some cases only after prior oligoadenylation of the 3'-end to facilitate binding of the 3-exonucleases. What dictates the choice of 3'-exonuclease is unknown. RNase II activity predominates over PNPase activity in E. coli (Deutscher and Reuven, 1991) and likely acts on the vast majority of accessible 3'-ends. Tightly folded structures lacking unpaired residues, particularly at 3'-ends, pose a formidable problem since RNase E, RNase II and PNPase are single-strand specific enzymes. This poses the question of how the degradosome degrades structured RNAs ? When the complex encounters an internal stem-loop structure it may skip past by looping out this structure (Figure 5.2C). Examples of RNAs where cleavages pass over highly structured internal regions include 9S R N A where p5S R N A is essentially by-passed (Ghora and Apirion, 1978; Misra and Apirion, 1979; Cormack and Mackie, 1992), the rpsO mRNA (Hajnsdorf er al, 1994) and stem III in the rpsT mRNA which separates two prominent cleavage sites, one at residues 190-191 and a second at residues 300-301 (Mackie, 1991; Mackie, 1992; Mackie and Genereaux, 1993). In principle, once bound to the 5'-end -197-o 5'-tethering as in Fig. 5.1. Degradosome dissociates. Cleavage product is resistant to further attack. Figure 5.4 Interrupted 5'-tethering model After several successive cleavage events, the degradosome may dissociate from its substrate to generate a product which is more stable than the initial primary transcript. The repositioning of an internal stem-loop to the 5'-terminus of the cleaved product would account for the stabilization of some downstream mRNA cleavage products as observed for the papBA mRNA of E. coli (Baga et al, 1988) and the RNase P-dependent stabilization of the his operon mRNA from S. typhimurium (Alifano et al., 1994). All symbols are as described in the legend to Figure 5.1. -198-of a substrate, the degradosome could engage and cleave a distal site at great distance from the 5'-end. Any internal structured fragments generated in this manner may be left for degradation by the 3-pathway as discussed in sections 5.2.2. Alternatively, the degradosome may encounter a highly stable internal stem-loop structure and stall. The stalled degradosome could enter a 3'- 5' mode of decay as shown in step 7 of Figure 5.1. Oligoadenylation catalyzed by PAP I could provide the signal for this transition to 3'-decay by an unknown mechanism. One could speculate that binding of PNPase to a poly(A) tail may trigger a conformational change in the associated RNase E resulting in the displacement of the end and allowing for exonucleolytic degradation of the 3-segment. In many cases, continuous cycles of polyadenylation may be required as poly(A) tails are extremely labile and are constantly turned over by RNase II, PNPase and possibly other nucleases (Coburn and Mackie, 1998b). Noticeably absent from this model is a putative role for the DEAD-box containing R N A helicase, RhlB. It has been reported that ATP hydrolysis and the presence of RhlB are required in vitro for PNPase-dependent decay of a fragment of the malE-malF R N A containing a strong stem-loop, the REP motif (Py et al, 1996). This can be interpreted to mean that RhlB unwinds REP sequences which normally serves as impediments to the processive action of PNPase (Newbury et al, 1987; McLaren etal, 1991). In contrast, ATP hydrolysis (to ADP and phosphate) and by implication RhlB, are not required to degrade structured portions of the rpsT mRNA in vitro (Coburn and Mackie, 1998b). We have also investigated the decay of the malE REP R N A with degradosomes under the conditions described in Py et al. (1996) and under the conditions of continuous polyadenylation. In both cases, degradation of the malE REP R N A requires ATP and the presence of RhlB (see Chapter 4). Thus, there appears to be a real difference between the degradation of -199-structured portions of the malE-malFRNA and the rpsT mRNA. Although the role of RhlB in the decay process remains unclear, it is possible that R N A unwinding by RhlB is only applicable to a small subset of RNAs. Alternatively, RhlB may play a role in facilitating some other aspect of the decay process (see section 5.2.4). 5.2.2 The latent 3'-decay model RNAs such as the ompA mRNA which contain a 5'-barrier to RNase E (Chen et al, 1991; Emory etal, 1992; Bouvet and Belasco, 1992; Hansen et al, 1994) would be resistant to decay in the 5'-tethering model of Figure 5.1. The model shown in Figure 5.5 demonstrates a purely 3-mode of decay dependent on exonucleases, primarily PNPase, and PAP I. This model may apply to RNAs which lack efficient RNase E sites (e.g., RNA-OUT) or contain an inhibitory 5'-terminal stem-loop to prevent binding of RNase E (e.g., ompA mRNA). The default pathway would require attack from a 3'-end. If the 3'-end of an mRNA is sequestered in a stem-loop structure, binding of 3-exonucleases would be inefficient but could be triggered by polyadenylation catalyzed by PAP I as outlined in Figure 5.5. Potential models of the mechanism of PAP I-dependent PNPase-mediated decay of structured RNAs are shown in section 4.4.3. It should be noted that the 3'-end of any mRNA is sequestered by R N A polymerase until the mRNA is completed. Thus, it is likely that only short completed mRNAs could enter this pathway while intact. The time to synthesize long polycistronic mRNAs such as the rpU-rplL-rpoB-rpoC mRNA can be much longer than the half-life of the mRNA itself (Chow and Dennis, 1994). In such cases, degradation must begin from the 5'-end before the 3-end is complete. -200-o o 5- ppp I OH 3' 5' PPP o o 4t ATP PAP I J | RNase II PPi o 1. Polyadenylation of 3' end by PAP I. 5' PPP Recycle RNase E + PNPase I I I o R AAAAAAAAAAAAAAAAAAAA OH 3' o 2. Binding and digestion of poly(A) tail by PNPase. 3. PNPase may stall 3' to the base of stable stem-loop structures. 4. Additional rounds of polyadenylation may be required. 4 Mononucleotides + short oligos Figure 5.5 The 3 '-latent decay model All symbols are as described in the legend to Figure 5.1. -201-The model in Figure 5.5 is referred to as the latent 3'-decay model because it conceivably can be applied to all mRNAs to some extent, regardless of the state of their 5'-end and their accessibility to RNase E. In particular, it serves as the pathway of small, highly structured RNAs (e.g., RNA-OUT (Pepe etal, 1994), the 3'-end of the rpsTmKNA (Mackie, 1989; Coburn and Mackie, 1996b ), REP stem-loop sequences (Newbury et al, 1987; McLaren et al, 1991) damaged RNAs and endonucleolytic fragments which are not retained by the degradosome. RNAs which are poorly structured at their 3'-termini are directly accessible to the 3'-exonucleases (Coburn and Mackie, 1996b). PNPase is known to undergo a transition from a non-processive (distributive) form to one capable of processive phosphorolysis associated with tight binding when the 3'-single-stranded tail is longer than 12-20 residues (Littauer and Soreq, 1982) (Figure 5.5, step2). Thus, when the initial substrate is modified by PAP I and ATP to a sufficient length, PNPase can bind and initiate processive phosphorolysis. Once in this mode PNPase will release nucleoside diphosphates until it encounters a stable stem-loop structure (Newbury et al, 1987; McLaren et al, 1991). PNPase stalls at such structures, although the life-time of the stalled intermediate is unknown. If the stem-loop "breathes" (i.e. a few base pairs at the base of the stem melt momentarily) then PNPase can renew its activity and remove one or a few exposed residues. Successive rounds of polyadenylation and processive phosphorolysis would weaken the stem-loop structure resulting in its eventual destruction as described previously in section 4.4.3. In addition ATP hydrolysis by RhlB may drive the unwinding of some R N A structures allowing PNPase in the degradosome access to otherwise buried residues. Although not shown in the model in Figure 5.5, possible interactions between the 5'- and 3'-ends mediated by RNase E and PNPase, respectively, as proposed by Xu and Cohen cannot be excluded during the operation of the 3'-latent pathway (Xu et -202-al, 1993). Such interactions would, of course, require that RNase E be engaged at the 5'-end of the mRNA being degraded. It is difficult to envisage how this pathway might operate during translation since active 3'-decay would generate truncated mRNAs and potentially truncated protein products. One key role for RNase II in mRNA decay may be to suppress this pathway by the removal of poly(A) tails. It has also been suggested that RNase E may also contribute to the removal of poly(A) tails (Huang et al, 1998). This would essentially force decay into dependence on a 5'-end as exemplified by the 5'-tethering model as is believed to be the case for most mRNAs (Nierlich and Murakawa, 1996). To my knowledge, competition between translating ribosomes and exonucleolytic digestion of an mRNA has never been examined much less demonstrated. Presumably even a stalled ribosome (e.g., at a termination codon) would prevent 3-exonucleases from progressing into the coding region of the message in a situation analogous to ribosome "toeprinting" by reverse transcriptase (Hartz et al, 1988). In the event that the 3'-latent decay pathway is engaged inappropriately, E. coli has evolved a protective mechanism which employs lOSa R N A charged with alanine to mimic an amino acyl tRNA and tag incomplete polypeptides for degradation (Keiler et al, 1996). It is an uninvestigated possibility that the lOSa R N A (tmRNA) mediated protein decay pathway could be linked to 3' -mRNA decay. Despite the preceding caveat, one can imagine that the 3'-latent pathway might make a significant contribution to the reduction of mRNA levels which accompanies translational repression of some mRNAs (Singer and Nomura, 1985; Zengel and Lindahl, 1994). When a repressor binds to an mRNA and blocks further translational initiation, the only barrier to exonuclease attack is distal R N A structure. Conceivably, translationally repressed mRNAs would be degraded from both ends -203-simultaneously. The models illustrated in sections 5.2.1 and 5.2.2 appear to be rather energy-wasteful due to the repeated use of ATP to form poly(A) tails and/or to drive the action of RhlB. Moreover, poly(A) tails are constantly turned over by RNase U, PNPase and possibly RNase E (Cao and Sarkar, 1992b; Huang etal, 1998). Several exits from this seemingly futile cycle of adenylation/deadenylation have been previously rationalized including scenarios where localized melting or strand-invasion followed by successive removal of nucleotides from the 3-(descending) base of the duplex decreases the overall stability of a 3'-terminal stem-loop structure (see section 4.4.3). Even so, these explanations remain somewhat unsatisfactory. Although polyadenylation in E. coli is thought to be a relatively indiscriminate process in vivo (Cao and Sarkar, 1992b; Sarkar, 1997), it is plausible that the degradosome may recruit PAP I directly to the 3'-end of 5'-monophosphorylated R N A fragments. In this vein, PAP I itself may show a preference for 5-monophosphorylated RNAs, as is the case for the Rne/Ams protein (G. A. Mackie, manuscript submitted). In addition, we have hypothesized that formation of a transient complex between PAP I and PNPase may induce a conformational change which enhances the processivity of the enzyme (see section 4.4.3). Such mechanisms which coordinate the action of PAP I and the degradosome are highly desirable since they would greatly reduce the requirement for ATP in the process of mRNA decay. 5.2.3 The 3'-tethering models. It has been suggested that polyadenylation may be the initial signal which triggers the decay of mRNA (O'Hara et al. 1995; Ingle and Kushner, 1996; Kushner, 1996). A third model to account for this possibility is illustrated in Figure 5.6.; it requires PAP I (or PAP II), and one or more -204-o ATP • PAP I J T RNase II PPi 1. Polyadenylation of 3'-end by PAP I. o a AAAAAAAAAAAAAAAAAAAA OH }' o 2. Binding and digestion of poly(A) tail by PNPase. »rrr\ c o Recycle RNase E + PNPase II t 3A. PNPase delivers RNase E to 5'-end. 3'-end is displaced. o 4A. 5'-decay pathway resumes (see 5'-tethering model). 3B. PNPase delivers RNase E to internal site. 4B. Both 5'- and 3'-ends are retained upon cleavage. Mononucleotides + short oligos Recycle RNase E + PNPase Mononucleotides + short oligos Figure 5.6 The 3'-tethering model Al l symbols are as described in the legend to Figure 5.1. -205-ribonucleases as discussed below. Polyadenylation of an mRNA catalyzed by PAP I would allow PNPase and/or the degradosome (via PNPase) to bind to the 3' end of an mRNA and initiate 3'-exonucleolytic decay. Alternatively, Kushner and his colleagues have suggested that a novel endonuclease dependent on hypothetical poly(A) binding proteins would initiate the decay process (Ingle and Kushner, 1996; Kushner, 1996). At some point the 3'-exonuclease engaged on the mRNA would encounter a stem-loop or a ribosome causing both to stall in the latter case. The RNase E component in a stalled degradosome could then bind to the 5'-end of an mRNA (step 3 A in Figure 5.6) and initiate 5'-3' decay similar to that in the 5-tethering model of Figure 5.1. In Figure 5.6, the 3'-end of the mRNA is displaced from PNPase by unspecified means. It is plausible that conformational changes within PNPase are induced by binding of RNase E elsewhere on the mRNA promoting the release of the 3'-end. Like the 5'-tethering model in Figure 5.1, the "end exchange" would also be sensitive to translation and to the positioning of R N A secondary structure at or near the 5'-end of the mRNA (see Figures 5.2 and 5.3). We could also envisage alternative models where PNPase might deliver RNase E to internal sites as shown in step 3B of Figure 5.6. One disadvantage of this model and its variants (O'Hara et al, 1995; Ingle and Kushner, 1996; Kushner, 1996) is that it deliberately provokes the stalling of ribosomes on an mRNA and consequently, the formation of truncated polypeptide products. Moreover several lines of evidence, described in section 4.4.2, tend to preclude any model which invokes polyadenylation as a common initiating event in mRNA decay. 5.2.4 5'-decay vs. 3'-decay by the degradosome Although a clear dependence of one enzyme on the other remains to be demonstrated, the discovery of a multienzyme complex controlling mRNA decay suggests a number of important -206-questions. Among others, what are the signals that switch the degradosome from a 5'-dependent endonuclease (RNase E) to a 3'-dependent exonuclease (PNPase) ? As discussed previously, polyadenylation catalyzed by PAP I may be one such signal. A second hypothetical signal may involve ATP hydrolysis and RhlB. Although the mechanism by which RhlB enhances the susceptibility of the malE REP R N A to degradation by PNPase remains unknown, it has been suggested that RhlB facilitates R N A degradation by unwinding R N A duplexes directly. We believe, however, that protein function should not be assigned on sequence homology alone. At least one author has suggested that DExD/H helicase proteins may perform functions other than R N A unwinding (Staley and Guthrie, 1998). In this regard, ATP hydrolysis by RhlB may drive conformational changes within the RNA-protein complex which displace the 5'-end and allow the degradosome to switch to a 3'-mode of decay. A potential paradigm for such a mechanism is provided by M o t l , a regulator of transcription, which displaces TBP from the T A T A box in an ATP-dependent fashion (Kadonaga, 1998). Alternatively, ATP hydrolysis may serve to juxtapose the 5' and 3'-ends of the R N A fragment to allow PNPase to attack a poly(A) tail while the degradosome maintains contact with the 5'-end via the Rne subunit. Recent observations of the decay of oligoadenylated RNA1 may suggest a means by which the Rne component of the degradosome could exercise overall control of mRNA decay (Huang et al, 1998). These authors showed that the N-terminus of the Rne/Ams protein contains an activity which is able to shorten poly(A) tails from the 3'-end of oligoadenylated RNA1. Moreover, poly(A) tails, ^7 residues appear to occlude cleavage of RNA1 by RNase E (Huang etal, 1998). A model which incorporates this potentially important phenomenon is shown in Figure 5.7. -207-o 1 5'ppp ~WC I T 3' OH AAAAAAAA \ J H OH 3" o H AAAAAAAAAAA AAAAAAAAA OH 3' \ Binding of subunit I to site 1 Binding of subunit II to poly(A) tail \ Cleavage of poly(A) tail 3' OH AAAAAA Binding and Cleavage at site 2 Figure 5.7 RNase E control of mRNA decay. All symbols are as described in the legend to Figure 5.1. -208-As in the 5'-tethering model of Figure 5.1, an RNA-binding domain in subunit I of the Rne protein binds to the 5'-end of an mRNA independent of 3-oligoadenylation. If internal sites within the mRNA are occluded by ribosomes, subunit II of the Rne protein may skip past the entire mRNA sequence and bind to a poly(A) tail of sufficient length. After successive removal of several adenylate residues from the 3'-end, subunit II may dissociate from the tail and resume "scanning" for internal sites (e.g., site 2). This hitherto unrecognized activity of the Rne/Ams protein may serve to prevent inappropriate engagement of the 3-latent decay pathway by removal of poly(A) tails from intact mRNAs. While the finding that the Rne/Ams protein can shorten poly(A) tails is certainly provocative, one should remain cautious when interpreting the data obtained in the study by Huang et al. (1998). First, only a portion of the Rne /Ams protein, consisting of the first 498 amino acids, was examined. This truncated protein lacks the R-RBD (see Figure 1.1). The presence of this second RNA-binding domain may serve to limit the cleavage specificity of RNase E (S. K. Masterman and G. A. Mackie, unpublished observations). To complicate matters, the degradosome may be the only physiological source of RNase E in the cell. Although it is clear that the Rne/Ams protein is the catalytic subunit of RNase E (Cormack et al, 1993), whether, or not, other components of the degradosome contribute to RNase E specificity remains unknown. Thus, these results should be repeated with both purified full-length Rne/Ams protein and degradosomes to verify the observed poly(A) shortening activity of RNase E. Second, to our knowledge, the effect of an ams/rne mutation on the degree of polyadenylation in vivo has never been examined, much less demonstrated. Moreover, previous work has shown that the degree of polyadenylation and length of poly(A) tails in vivo are dramatically increased in strains which lack both RNase II and PNPase but are otherwise wild-type for RNase E activity (Cao and Sarkar, 1992b; Hajnsdorf et al, 1995; Haugel-Nielsen et al, 1996). These results -209-have led others to conclude that the majority of poly(A) tail trimming in vivo is accomplished by the action of the 3-exonucleases and not RNase E. 5.3 Paradigms for the selective decay of mRNA Only the degradation of a few dozen mRNAs has ever been studied in any detail, a small representation of the 2,000-4,500 genes in Escherichia coli. Moreover, our present knowledge of mRNA decay is based largely on information obtained from the detailed study, either in vivo or in vitro, of a few model messages. To make matters worse, detailed structural information is available for only two RNAs: RNA1 and the rpsT mRNA. Thus, it is unlikely that our models will describe the process of decay for all messages. In short, there is room for additional paradigms, perhaps many. It is believed, however, that many aspects of these models enjoy some degree of experimental support or are amenable to testing. In the following sections, the decay of three well-documented RNAs will be rationalized in light of our models. 5.3.1 RNA1 Degradation of RNA1 is initiated by a single RNase E cleavage five residues from its 5'-end (Tomcsanyi and Apirion, 1985; Lin-Chao and Cohen, 1991; see Figure 5.8). This leads to the rapid 3'-exonucleolytic decay of an intermediate, termed pRNAl . 5 , dependent on PNPase and PAP I (Xu etal., 1993; Xu and Cohen, 1995; Figure 5.8). In contrast, "pre-cleaved" pppRNAl. 5 , generated from a plasmid was not efficiently degraded in vivo (Xu et al., 1993). Taken together, these observations suggest that the initial cleavage event by RNase E engages the degradosome on RNA1 and "delivers" PNPase to the 3'-end. Put differently, RNase E and the degradosome serve as a platform from which -210 -O o pppRNAI 5' PPP ^ RNase E Cleavage o ° o pRNA1^ 5' P — 1 \ Poly(A) tail addition by PAP I 5" P AAAAAAAAAAA PNPase-mediated degradation of polyadenylated RNase E cleaved RNA1 Mononucleotides + short oligos Figure 5 . 8 Summary of the decay of RNAL A schematic diagram summarizing the decay of RNA1 is shown above. The RNase E cleavage site, 5 nucleotide residues from the 5-terminus, is denoted by the black box. -211-the decay of RNA1 is launched. The dependence of decay of RNA1 on RNase E provides strong evidence that 3'-products are preferentially bound to or retained by the degradosome. 5.3.2 The rpsT mRNA The two rpsT mRNAs of 447 and 356 residues are transcribed from two tandem promoters separated by 91 bp and share a common terminus generated by a Rho-independent terminator (Mackie, 1986). Their decay displays characteristics compatible with both the 5'-tethering and the 3'-latent decay models. The half-lives of the two rpsT mRNAs increase significantly (2.5-fold) in strains deficient in PNPase (Mackie, 1989). Moreover, a 147-residue fragment coterminal with the 3'-end of the rpsT mRNAs is highly stabilized by pnp or pcnB mutations (Mackie, 1989; Coburn and Mackie, 1998a).These observations were interpreted to mean that the degradation of the rpsT mRNAs could be initiated by the 3-latent decay pathway (Mackie, 1989). Investigations with purified enzymes have confirmed this supposition: continuous polyadenylation catalyzed by PAP I is sufficient to permit PNPase, alone or in the degradosome, to catalyze the complete decay of a full-length rpsT mRNA (Coburn and Mackie, 1998a). A non-hydrolyzable analog or ATP, ATPyS, supports this process, further evidence that the putative helicase activity of RhlB is not required, unlike the case for the malE REP sequence (Py etal, 1996). In the absence ofPNPase, the rpsT mRNAs still decay, and accumulate a 3'-fragment corresponding to highly folded sequences distal to the most 3' RNase E site in the mRNA (Mackie, 1989). A body of data obtained in vitro is consistent with use of the 5'-tethering pathway involving RNase E and RNase II to degrade the 5-two-thirds of the rpsT mRNAs (Mackie, 1991; Mackie, 1992; Mackie and Genereaux, 1993; Mackie et al, 1997). The most interesting feature, therefore, of the decay of the rpsT mRNAs is that either a 5'- or a 3'-pathway can -212-be followed. It is conceivable that the 3-pathway is, in fact, initiated by the initial binding of the degradosome to the 5'-end of the r^rmRNA followed by its delivery to the 3'-end. 5.3.3 The rpsO mRNA The degradation of the rpsO mRNA in vivo is one of the best characterized of all mRNA decay processes in E. coli. Phillipe Regnier and his colleagues have summarized their work in a recent concise and provocative review (Hajnsdorf et al, 1996). Much of their in vivo work on the rpsO mRNA support our data obtained for the rpsT mRNA in vitro. Moreover, many of their interpretations conform in broad outline to the models we have proposed. In the following section we will discuss how our models might explain the decay of the rpsO mRNA. The rpsO gene can give rise to a number of different mRNA species resulting from termination of transcription at tl or from processing of primary transcripts by RNase III (RIII; Figure 5.9) and/or RNase E (Ml and M2; Figure 5.9) (Portier et al, 1987; Hajnsdorf et al, 1994). Any transcripts which escape termination at tl are processed by the distal RNase III site and may be subjected to exonucleolytic "trimming" by RNase II and possibly PNPase. Such species would be indistinguishable from those which had actually terminated at tl. Interestingly, however, PAP I does not appear to affect the stability of RNase III processed rpsO mRNAs (Hajnsdorf et al, 1995; Hajnsdorf et al, 1996). The main degradation pathway of the rpsO mRNA involving cleavage by RNase E (Hajnsdorf etal, 1994), fits many aspects of the 5'-tethering model. After initial recognition and binding of the degradosome to the 5'-end, the majority of the transcript would be looped out, possibly skipping some internal sites (Braun et al, 1996; Hajnsdorf et al, 1996). The preferred internal site of RNase -213-E cleavage, denoted as M2, lies 10 residues downstream from the stop codon and is immediately 5' to the base of the Rho-independent terminator, tl (Portier et al, 1987; Hajnsdorf et al, 1996; see Figure 5.9). Cleavage at the M2 site would remove the protective barrier to exonucleolytic degradation, particularly by PNPase and possibly by RNase II, afforded by the presence of the Rho-independent terminator.Binding and cleavage by the RNase E component of the degradosome is apparently affected by translating ribosomes (Braun, Le Derout and Regnier, cited in Hahndorf et al, 1996). Ribosomes stalled at the M2 site would provide sufficient steric hindrance to occlude the M2 site. In addition translation would also be expected to inhibit recognition of internal cleavage sites in steps 2-6 of the tethering model in Figure 5.1 A. RNase E also cleaves the rpsO message in vitro at additional internal sites between the 5'-end and the M2 site (Braun et al, 1996). The corresponding cleavage products are not detectable in vivo, raising the possibility that these sites are inefficient in vivo. If so, this provides evidence that some potential sites can be skipped by the degradosome, a possibility compatible with the tethering model. Alternatively, it is possible that these products decay too quickly to be detected by primer extension analyses. In this vein, at least 12 internal cleavage sites have been mapped in the similarly sized rpsT mRNA in vitro, but only two of these can be detected readily in vivo (Mackie, 1991). These observations in both systems point to the need to determine the effect, if any, of actively translating and stalled ribosomes on the ability of RNase E and degradosomes to recognize and cleave mRNAs in vitro. RNase E cleavage at site M2 generates a new 3'-end for exonuclease attack (Figure 5.9). RNase II stalls and presumably dissociates from the rpsO mRNA 3' to the base of potential internal stem-loop structures (Hajnsdorf etal, 1994; Braun etal, 1996). Polyadenylation of structured 3-ends, suggested by the discovery of several internal polyadenylation sites in the rpsO mRNA in strains -214-rpsO pnp 5' O M2 M1 RIM i]LL_ Rill 5' M 2 ^ M 1 P1-RIII 5' 5' PNPase /RNase II PAP I ? Mononucleotides + short oligos 5' P1-t1 and/or processed P1-RIII transcripts RNase E Cleavage at M2 J PAP I PNPase Mononucleotides + short oligos Figure 5.9 Summary of the decay of the rpsO mRNA. A schematic representation of the decay of the rpsO mRNA is shown above. The coding sequence of the rpsO and partial coding sequence of the pnp gene are denoted by the open boxes. The sites of RNase III cleavage are maked by RJII and the RNase E cleavage sites are marked by M l and M2 respectively. P l - t l represents the primary transcript which initiates at promoter PI and terminates at the Rho-independent terminator stem-loop (tl). Pl-RIII represents the RNase III processed transcript initiated at promoter PI . -215-deficient for RNase II and PNPase, may be involved in reinitiating the decay of stalled intermediates (Haugel-Nielsen et al, 1996). Decay of the rpsO mRNA also fits aspects of the 3-latent decay pathway (Figure 5.6). First, the rpsO mRNA is mildly stabilized in pnp mutant strains in vivo (Hajnsdorf et al, 1994). Moreover, the half-life of the rpsO mRNA decreases by two-fold in an rnc strain which overexpresses PNPase by up to 10-fold (Hajnsdorf et al, 1994). These data suggest, that like the rpsT mRNA, the rpsO mRNA can be degraded in a PNPase-dependent pathway in which the degradosome either initiates phosphorolysis directly at the 3'-end of the mRNA, or switches from an early step in the 5'-tethering pathway. Investigations of the role of polyadenylation in the decay of the rpsO mRNA have provided some data which do not fit either the 5'-tethering or 3-latent decay pathways. In an ams rnb pnp triple mutant, the rpsO mRNA decays unexpectedly much faster than in otherwise isogenic ams, ams pnp, or ams rnb strains (Hajnsdorf et al, 1994; Hajnsdorf er al, 1995). Models to explain how polyadenylated R N A might be degraded in the absence of exonucleases involve base pairing of the poly(A) tail with internal sequences in the rpsO transcript (Hajnsdorf et al, 1996). This might provide new secondary structures for an unknown RNase (possibly an RNase III signal sequence). It is not clear how "strand invasion" by a poly(A) tail might occur during translation, however. Other exoribonucleases such as those involved in tRNA maturation could, conceivably, participate in mRNA decay in the absence of RNase II or PNPase. 5.4 Perspectives As alluded to above, our knowledge of the process of mRNA decay is based on only a small -216-sample of all known bacterial mRNAs. This points to the need to define new paradigms which, hopefully, will unlock clues to the mechanism of mRNA decay. Obvious experiments include the application of the in vitro decay system to a number of mRNAs both simple and complex. RNAs which have already been well studied in vivo, such as the Sok-RNA, the anti-sense regulator of plasmid RI encoded programmed cell death, or the rpsO mRNA may be particularly advantageous. At any rate, we believe that a balanced approach of in vivo and in vitro experimentation, coupled to structural information, will be required to accurately describe the decay process for any given mRNA. A demonstration that the degradosome is physiologically relevant is of paramount importance. This may be accomplished, in part, by determining the relative ratios of the degradative enzymes in vivo. Although confirmation of the degradosome components and their stoichiometry is an important first step, a clear dependence of one enzyme on the other in the degradosome remains forthcoming. Further reconstitution experiments will help to clarify the roles and mechanism of action of each component of the degradosome and may give some clues to its assembly. This will, of course, depend on our ability to develop novel strategies for the purification of RNase E. One shortcoming of the in vitro decay system is that it ignores the role of additional factors which may influence the decay process. We are actively pursuing the development of a coupled in vitro translation-mRNA decay system to examine the role of ribosomes, translational repressors and other putative decay factors (e.g., CsrA) in the decay of model mRNAs. In this vein, Kushner and his colleagues have had some limited success studying the decay of polysome associated mRNAs in vitro (Ingle and Kushner, 1996). One interesting observation of R N A metabolism in Escherichia coli is the finding that mRNA decay enzymes are also involved in the processing of stable RNAs such as ribosomal R N A and -217-transfer R N A . It is plausible that additional protein factors may play a role in R N A processing by interrupting both 5 - and 3'-dependent mRNA decay pathways. Similarly, stability factors may contribute to the differential stability of polycistronic messages by this mechanism. We believe that in the next few years the study of mRNA decay will mature from its initial descriptive phase into one which will focus on mechanism: recognition of cleavage sites, catalytic mechanisms, and regulation. It is likely that high resolution structural information of both enzymes and model substrates will play an important role in defining mechanisms. The in vitro system provides an excellent framework for verifying the apparent 5'-3' directionality of mRNA decay, as well as addressing the apparent processive "all or none" mechanism of decay observed for most mRNAs (Nierlich and Murakawa, 1996). We have little doubt, therefore, that the next few years of study will be extremely exciting as we gain new insights into this important cellular process. -218-References Alifano, P., Rivellini, F., Piscitelli, C , Arraiano, C. M . , Bruni, C. B. , and Carlomagno, M . S. (1994) Ribonuclease E provides substrates for RNase P-dependent processing of a polycistronic mRNA. 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