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Composition and adpatation of the E. coli RNA degradosome Prud’Homme Genereux, Annie 2004

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Composition and Adaptation of the E. coli RNA Degradosome by ANNIE PRUD'HOMME GENEREUX B.Sc. (Hons.), McGill University, 1997 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES Department of Biochemistry and Molecular Biology We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA February 2004 © Annie Prud'homme Genereux, 2004 ABSTRACT Although transcription and translation are recognized mechanisms for regulating gene expression, control over RNA stability can also accomplish this task. In E. coli, bulk mRNA degradation is largely carried out by a complex of enzymes called the degradosome. It is composed of the endonuclease RNase E, the exonuclease PNPase, the helicase RhIB, and the glycolytic enzyme enolase. A role in mRNA decay has been assigned to the first three proteins, but the function of enolase is currently unknown. The hypothesis that it senses the metabolic state of the cell and alters the activity of the degradosome accordingly was tested. Assays using model substrates were performed with degradosomes reconstituted from purified components, in the presence and absence of enolase. The presence of this enzyme in the assay had no apparent effect on the activity of either RNase E, PNPase, or RhIB. Simulation of possible signals impinging upon enolase, such as binding of 2-phosphoglycerate or phosphoenolpyruvate, or phosphorylation, exerted only a very modest effect on degradosome activity. While purified RNase E, PNPase, and RhIB reconstitute a complex when incubated together, enolase appears to require the assistance of PNPase and/or another factor to assemble with RNase E. These studies have failed to identify a role for enolase in adapting the activity of the degradosome to metabolic signals. Other mechanisms for altering the function of the degradosome were investigated by studying the complex during and after cold shock. Under those conditions, CsdA, an ATP-dependent RNA helicase enters the complex. Three separate types of exper iments support the presence of C s d A in the cold shock degradosome: co-purif ication, co-immunoprecipi tat ion, and reconstitution from purified components . This enzyme is capable of replacing RhIB in the in vitro degradat ion of a substrate that requires the coordinated activity of both P N P a s e and RhIB for degradat ion. The composi t ion of the degradosome is altered in response to a temperature downshift, and a novel mechan ism of regulating the activity of the degradosome, and presumably gene express ion , has therefore been d iscovered. TABLE of CONTENTS ABSTRACT ii LIST OF FIGURES ix LIST OF TABLES x LIST OF ABBREVIATIONS xi ACKNOWLEDGEMENTS xiii CHAPTER I - INTRODUCTION 1 1 mRNA DEGRADATION 1 1.1 mRNA instability 1 1.2 Importance of mRNA degradation 2 1.3 Properties of mRNA decay 4 1.4 Models of mRNA decay 4 1.4.1 Ribonucleases 4 1.4.2 The Apirion model 5 1.4.3 Factors affecting mRNA decay 7 1.4.4 Current Model 8 1.5 Enzymes of mRNA decay 9 1.6 The E. coli RNA degradosome 9 1.6.1 Purification 14 1.6.2 RNase E 14 The protein 14 Catalytic activity 16 Scaffolding region 19 1.6.3 PNPase 19 The protein 19 Catalytic activity 21 1.6.4 RhIB 24 The protein 24 ATPase activity 26 Helicase activity 27 Association of RhIB with PNPase and poly(A) polymerase I 27 iv 1.6.5 Enolase 29 Glycolytic enzyme 29 Possible roles in decay 30 1.6.6 Other components 31 Polyphosphate kinase 31 Poly(A) polymerase I 33 S1 ribosomal protein 35 DnaK 36 GroEL 36 EIF, CspE, and RNA polymerase p and P' subunits 37 1.6.7 Mass and stoichiometry of the degradosome 39 1.6.8 Advantages of association of enzymes into a degradosome 41 1.6.9 Arguments against its existence in vivo 43 1.6.10 Arguments in favour of its existence in vivo 45 1.6.11 RNA decay machines in other organisms 48 RNase E-based degradosomes 49 The exosome 51 The mitochondrial degradosome .... 51 The chloroplast degradosome 52 1.7 Adaptation of mRNA decay 53 1.7.1 Global responses 53 1.7.2 Specific responses 55 2 GOALS 57 CHAPTER II - MATERIALS & METHODS 59 2.1 INTRODUCTION 59 2.2 SOURCE of ENZYMES and CHEMICALS 59 2.3 BACTERIAL STRAINS AND PLASMIDS 59 2.3.1 Overexpressing strains 60 2.3.2 pnp and csdA mutant strains 64 2.4 BACTERIAL CULTURES 65 2.4.1 Growth media 65 2.4.2 Rescue of strains from frozen stocks 66 2.4.3 Induction of cultures 66 2.4.4 Recovery of cells by centrifugation 67 2.4.5 Bacterial transformation 67 v 2.5 ENZYMES and ANTIBODIES 68 2.5.1 Enzyme purification 68 Degradosome purification 68 Rne purification 73 Pnp purification 76 Enolase Purification 79 CsdA Purification 80 2.5.2 Antibody sources 81 2.6 PROTEIN ANALYSIS 83 2.6.1 Ammonium sulfate concentration of proteins 84 2.6.2 Acetone precipitation of proteins 84 2.6.3 Protein quantification 84 2.6.4 Protein / SDS-PAGE gel 85 2.6.5 Western blot 86 2.6.6 Identification of proteins by mass spectrometry 86 2.7 RNA PREPARATION and PROTOCOLS 88 2.7.1 Preparation of template DNA 88 2.7.2 Linearization of plasmids for transcription 89 2.7.3 In vitro synthesis of RNA transcripts 90 2.7.4 Electrophoretic analysis of RNA 92 2.8 TRANSMISSION ELECTRON MICROSCOPY 93 2.9 IMMUNOPURIFICATION of DEGRADOSOMES from CELL 93 EXTRACTS 2.9.1 Using antibodies to specific proteins 93 2.9.2 Using FLAG-Rne and anti.-FLAG antibodies 95 2.9.3 Using His-Rne and a TALON Column 96 2.10 CO-IMMUNOPRECIPITATION EXPERIMENTS USING 97 PURIFIED PROTEINS 2.10.1 Proteins without tag 97 2.10.2 Using FLAG-RNase E 98 2.11 R E C O N S T I T U T E ASSAYS 98 2.11.1 RNase E assay 99 2.11.2 PNPase and RhIB assay 99 2.12 ATPASE ASSAYS 100 2.13 ENOLASEASSAY 101 2.14 COMPUTATIONAL METHODS 101 vi CHAPTER III - E N O L A S E & T H E RNA D E G R A D O S O M E 103 3.1 BACKGROUND 103 3.2 RESULTS 104 3.2.1 Enolase Purification and Activity 104 3.2.2 RhIB Purification 105 3.2.3 Addition of enolase to in vitro degradosome assays 109 3.2.4 Effects of enolase ligands on degradosome activity 113 3.2.5 Physical association of RNase E and enolase in vitro 116 3.2.6 Physical association of RNase E and enolase in vivo .... 118 3.3 DISCUSSION 123 3.3.1 Does enolase influence degradosome activity? 123 3.3.2 Does enolase bind RNase E? 125 3.3.3 Possible role of enolase in "internal entry" 127 CHAPTER IV - CHARACTERIZATION of a " C O L D S H O C K " 130 D E G R A D O S O M E 4.1 BACKGROUND 130 4.1.1 Overview of effects of cold shock 131 4.1.2 Stages of cold shock 135 4.1.3 mRNA decay during cold shock 136 4.1.4 CsdA 140 4.1.5 DEAD-Box Helicases 143 4.2 RESULTS 148 4.2.1 Time course of CsdA induction during cold shock 148 4.2.2 Existence and composition of the cold shock degradosome 150 Biochemical purification of cold shock degradosomes 150 Immunopurification of cold shock degradosomes 154 Co-immunoprecipitation of purified RNase E, PNPase, and CsdA 160 4.2.3 Functional interaction between CsdA and PNPase 160 4.2.4 ATPase activity of CsdA 164 4.3 DISCUSSION 167 4.3.1 Evidence for CsdA in cold shock degradosomes 167 4.3.2 Reconstitution of CsdA-containing "minimal degradosomes" 172 vii 4.3.3 mRNA decay and cold shock 174 4.3.4 A model for expression and function of CsdA during 177 cold shock CHAPTER V - CONCLUSIONS 180 5.1 The role of enolase in the degradosome 180 5.2 Cold shock and the RNA degradosome 180 5.3 Perspectives on adaptation 182 REFERENCES 184 viii LIST of FIGURES Figure Description Page 1.1 Apirion model of mRNA decay 6 1.2 Current model of RNA degradation 10 1.3 Proposed model of RNA-degradosome interactions 12 1.4 Regions of interest on RNase E primary sequence 13 1.5 Environmental determinants of PNPase activity 25 2.1 Purified proteins 69 3.1 Assay of enolase activity 106 3.2 RhIB purification on Resource S column 108 3.3 Diagram of the reactions and intermediates created in reconstitution assays 110 3.4 Effect of enolase on the activity of degradosome proteins in vitro 111 3.5 Effect of enolase ligands on degradosome activity 114 3.6 in vitro association of enolase with RNase E 117 3.7 Degradosomes purified from cell extracts by different methods all contain enolase 119 3.8 Immunoprecipitation of the degradosome from Apnp cells 122 4.1 Rate of bulk and cold shock protein synthesis following temperature downshift 137 4.2 Alignment of CsdA and RhIB amino acid sequences 146 4.3 Time course of CsdA induction during cold shock 149 4.4 Degradosomes isolated from cells grown at 37°C or 15°C 152 4.5 Co-immunopurification from unlabeled cell extracts 156 4.6 Co-immunoprecipitation from radiolabeled cell extracts 158 4.7 Physical interaction of Rne, Pnp, and CsdA in vitro 161 4.8 Functional equivalence of RhIB and CsdA in vitro 163 4.9 ATPase activity of CsdA and degradosomes 165 4.10 Model of CsdA activity during cold shock 179 ix LIST of TABLES Table Description Page 1.1 Degradosome stoichiometry 40 2.1 List of bacterial strains 61 2.2 List of plasmids 62 2.3 Physical properties of purified proteins 70 2.4 List of antibodies 82 4.1 List of all currently known E. coli proteins expressed in response to cold shock 132 4.2 Identification of cold shock degradosome proteins by mass spectrometry 153 x LIST of ABBREVIATIONS ADP Adenosine diphosphate ARRBD Arginine-rich RNA binding domain AS26 Precipitated fraction from a 26% (w/v) ammonium sulfate extraction ATP Adenosine 5'-triphosphate BSA Bovine serum albumin °C Degree Celsius CsdA The csdA (cold shock DEAD-box protein A) gene product csp Cold shock protein C-terminal Carboxyl-terminal DEAD (aspartate-glutamate-alanine-aspartate) DEPC Diethyl pyrocarbonate DNA Deoxyribonucleic acid DNase Deoxyribonuclease DTT 1, 4-dithiothreitol E. coli Escherichia coli EDTA Ethylenediamine tetraacetate EM Electron microscopy FLAG (aspartate-tyrosine-lysine-aspartate-aspartate-aspartate-aspartate-lysine) FPLC Fast protein liquid chromatography GET Glucose-EDTA-Tris HEPES N-2-hydroxy ethyl piperazine-N'-2-ethanesulfonic acid His Histidine igG Immunogloblulin G IP Immunoprecipitation IPTG lsopropyl-|3-D-thiogalactopyranoside kDa kiloDalton LB Luria-Bertani broth MALDI Matrix-assisted laser desorption ionization MalEF Intergenic spacer region of the MalE-MalF mRNA mRNA Messenger ribonucleic acid MW Molecular weight MWCO Molecular weight cut off NaF Sodium fluoride OD 6 0o Optical density at a wavelength of 600 nm PAGE Polyacrylamide gel electrophoresis PAP Poly(A) polymerase PBS Phosphate buffered saline PEI Poly(ethylenimine) Pi Phosphate PMSF Phenylmethylsulfonylfluoride Pnp The pnp (polynucleotide phosphorylase) gene product xi PNPase Polynucleotide phosphorylase Poly(A) Polyadenylate (polyadenylic acid) psi Pounds per square inch REP Repetitive extragenic palindrome RhIB The rhIB (RNA helicase-like gene A) gene product RNA Ribonucleic acid RNase Ribonuclease Rne The Rne (ribonuclease E) gene product rpm Revolutions per minute rRNA Ribosomal RNA S30 Supernatant of 30,000 x g centrifugation SDS Sodium dodecyl sulfate TAE Tris-acetate-Na EDTA TBE Tris-boreate-Na EDTA TE Tris-EDTA TEMED N, N, N', N'-tetramethylethylenediamine TLC Thin layer chromatography Tris Tris(hydroxymethyl) aminomethane tRNA Transfer RNA U Units UV Ultra-violet v/v Volume/volume w/v Weight/volume xii ACKNOWLEDGEMENTS First and foremost, I would like to thank Dr. George Mackie to have made this work possible. George, your hard work, dedication, patience, encouragement, knowledge and fairness are what made this project a positive experience. I would also like to thank the people who have advised me on my work, as well as those who have reviewed this dissertation and provided helpful comments and criticism: Dr. Peter Candido, Dr. Patrick Dennis, Dr. Leann Howe, Dr. Ross MacGillivray, Dr. Ivan Sadowski, and my university and external examiners, whomever you may be... The cold shock work is a result of a collaboration of our laboratory with Dr. Robert Simons and his then Ph.D. student Dr. Rudolf Beran at U.C.L.A., so I would like to acknowledge their contribution. The EM work was done at the Bioimaging Facility at U.B.C., with much help from Garnet Martens. The mass spectrometry and protein identification was performed by the University of Victoria - Genome British Columbia Proteomics Centre. I cannot forget the two rabbits who have given up their lives for the production of antibodies for these experiments. I am so sorry for subjecting you to this treatment, and hope you suffered very little or not at all. May your little bunny spirits soar high and free in the afterlife... I would have accomplished very little in the laboratory without constant help from labmates, so I would like to thank Janet Hankins, Catherine Spickler, Stephanie Masterman, Dr. Glen Coburn, Dr. Xin Miao, Dr. Douglas Briant, soon to be Master Robert Edge, Dr. George Jones, and the numerous (but equally important) undergraduates whom have graced the lab with their curiosity and smiles through the years (with special mention of Michael Cook, since he did spend so long with us...). Last, but definitely not least, I would like to thank our lab dishwashers, Roy and Eric, for their diligence and hard work (and pies!), and for allowing me to work even more efficiently... All work and no play makes for a very dull Ph.D., so I wish to thank a few friends who have made my life such a great time in Vancouver: Alisen, Ammen, Annick, Dave, Edwin, Emily, Eveline, Isabel, Jason, John, Kati, Kyle, Larissa, Laurie (and Bob), Lili, Marieke, Sonia, Spencer, Susan, and Warren. Special mention should be made of Gordon, who has encouraged me in all I do for the past 3 years. Thanks to all my dive buddies, and to the Aquarium, Aquasoc and VNHS crew who share my enthusiasm for underwater life. Although he probably won't understand this, thanks to Cinnamon for being so sweet and such a good stress reliever - don't worry, I'll never inject you with antibodies or serve you up on a platter at Easter dinner... And while we're at it, thanks to that special sealion for kissing me at Hornby Island... Finally, I would like to thank the members of my family for their constant and unconditional support. M e r c i b e a u c o u p Lise e t Jean-Louis p o u r m ' a v o i r d o n n e l e s o u t i e n m o r a l ( e t p a r f o i s f i n a n c i e r ) d o n t j ' a i e u b e s o i n a u c o u r s d e c e s 6V2 d e r n i e r e s a n n e e s . M e r c i a u s s i a m e s d e u x g r a n d - m e r e s Fernande e t a Tony pour l e u r a f f e c t i o n e t p o u r m ' a v o i r r e p e t e d e n o m b r e u s e s f o i s l e u r f i e r t e e n v e r s m e s r e u s s i t e s . V o i r e a m o u r r e s t e d a n s m o n c o e u r e t e s t u n e f o r c e q u i m ' a p e r m i s ( e t m e p e r m e t t r a ) d e p e r s e v e r e r d a n s d e s m o m e n t s d i f f i c i l e s t o u t a u l o n g d e m a v i e . . . xiii CHAPTER I INTRODUCTION 1 mRNA DEGRADATION 1.1 mRNA instability In 1961, the existence of an unstable intermediate between deoxyribonucleic acid (DNA) and proteins was proposed to explain how gene expression can be reprogrammed rapidly (Brenner et al., 1961; Gros et al., 1961). The instability of messenger ribonucleic acid (mRNA) was therefore anticipated before the discovery of its coding properties, highlighting the importance of this property to its function. It is now known that although roughly 50% of cellular transcription is dedicated to making mRNAs, these transcripts only account for 2-3% of the RNA mass of the cell (Levinthal etal., 1963; Kennell, 1968; Nierlich, 1978). This is a consequence of their instability. In fact, in prokaryotic species, long polycistronic mRNAs may begin to decay before their synthesis is complete (Chow & Dennis, 1994; Cannistraro & Kennell, 1985). Typical mRNA half-lives in Escherichia coli [E. coli), where this phenomenon has been most extensively studied, range from 20 seconds to 50 minutes, with an average half-life of 1-2 minutes (Belasco, 1993; Nierlich & Murakawa, 1996). 1 1.2 Importance of mRNA degradation Bacteria are very efficient at utilizing and conserving energy, so it may first seem counterintuitive that long nucleotide polymers are synthesized only to be degraded shortly thereafter. However, the energetic cost of mRNA turnover is only 7% of the total cost of protein expression1 (Nierlich & Murakawa, 1996). In exchange for this minor loss, the cell gains a much tighter control over its gene expression, because mRNA stability is as important in determining the amount of protein produced as transcription and translation (Mott et al., 1985; Deutscher, 1988; Jain & Belasco, 1995; Grunberg-Manago, 1999). Given two transcripts with identical promoter strengths and translational efficiencies but differing stabilities, the one with the longer lifespan will be available for translation by ribosomes for a longer period of time, thus generating more protein product. As foreseen in 1961, mRNA instability is also crucial in the rapid adaptation of gene expression to changing environmental conditions (Brenner et al., 1961; Gros et al., 1961; Jacob & Monod, 1961). Once transcription ceases in response to environmental signals, protein expression continues only as long as mRNAs are present. Thus, labile mRNAs permit rapid changes in gene expression. 2 mRNA decay can account for the differential gene expression of some polycistronic mRNAs. Since signals within RNAs determine their susceptibility to degradation (Section 1.4.3), distinct sections of polycistronic mRNAs can have different stabilities (Newbury et al., 1987a; Baga et al., 1988; Nilsson & Uhlin, 1991; Yajnik & Godson, 1993; Cam et al., 1996). RNA degradation returns nucleotides to the cellular pool, and it has been estimated that roughly half of the nucleotides required for biosynthesis are provided by this recycling pathway (Deutscher & Reuven, 1991; Kennell, 2002). mRNA decay is also useful in quality control. Removing incorrectly folded or terminated mRNAs from the cell is important, as these could compete with wild-type transcripts for ribosomes. A few biotechnological and medical applications have emerged from the study of mRNA decay. First, the key enzymes and factors affecting RNA decay have been identified and can now be used to optimize the production of proteins in bacterial overexpression systems (Rosenberg et al., 1987; lost & Dreyfus, 1994; Lopez etal., 1999; Smolke & Keasling, 2002; Zhan etal., 2004). Second, some proteins implicated in RNA decay are also necessary for virulence in certain pathogenic bacteria, making them alluring targets for the creation of novel antibiotics (Cheng etal., 1998; Clements et al., 2002). Finally, natural anti-sense 1 The cost of incorporating one amino acid into a protein (including amino acid activation and ribosome-associated elongation steps) is 4 high energy phosphate bonds. The cost of incorporating three nucleotides into a transcript to make a codon is 3 NTPs, or 6 high energy bonds. Since an mRNA is translated an average of 20 times in E. coli (Salser et al., 1968; Bremer & Dennis, 1996), the cost of making a codon is spread over 20 translation events, so it is 6/20. The cost of translating a codon is equal to the cost of making it plus the cost of translating it, so it is equal to 4 + 6/20. Only 6/20 high energy bonds will be lost in the degradation of a codon, which corresponds to 7% ( = [(6/20) * 100 / (4 + 6/20)]) of the energy used in protein 3 nucleic acids, used by the cell or by infecting pathogens to target the degradation of specific RNAs have given rise to both experimental and therapeutic strategies for reducing the amount of specific nucleic acids in cells (Tomizawa et al., 1981; Simons & Kleckner, 1983, 1988; Inouye, 1988; Eguchi era/., 1991; Hilleren & Parker, 1999; Bass, 2000; Maquat & Carmichael, 2001). 1.3 Properties of mRNA decay mRNA decay seems to be an all-or-none phenomenon (few intermediates are ever found) (Coburn & Mackie, 1999). It appears to progress in a 5'->3' direction on the RNA (Morikawa & Imamoto, 1969; Morse et al., 1969; Forchhammer era/., 1972; Yamamoto & Imamoto, 1975; Cannistraro & Kennell, 1985; Cannistraro etal., 1986; Baga etal., 1988; Emory & Belasco, 1990; Emory et al., 1992; Goodrich & Steege, 1999; Steege, 2000). This is perhaps not surprising given the prevalence of 5'->3' directionality in nucleic acid biology (for example, DNA synthesis, RNA synthesis, translation...), and also the fact that the 3'-end of RNA may be sequestered by RNA polymerase or stable secondary structures when degradation is initiated (Chow & Dennis, 1994; Cannistraro & Kennell, 1985). The advantages of degrading mRNA in a 5'->3' direction are that all new translation is blocked, and ribosomes which have already initiated synthesis are allowed to finish correctly (no erroneous proteins are created) (Kennell, 2002). expression of that region. (N.B.: This estimate only holds true for unstructured regions (see Section 1.4.4)). 4 1.4 Models of mRNA decay 1.4.1 Ribonucleases RNA phosphodiester bonds are very stable, with estimated half-lives in the order of millions of years (Sreedhara & Cowan, 2001). Enzymes must therefore play a pivotal role in regulating the amount of transcripts present in the cell. These can be grouped into three categories: the endoribonucleases that cleave phosphodiester bonds within an RNA molecule, the exoribonucleases which progressively remove nucleotides from the end of a transcript, and a third group, with varied functions but no nuclease activity that help the ribonucleases (RNases) accomplish their task. 1.4.2 The Apirion model In the 1970s, based on observations he and others had made on the kinetics of mRNA turnover, David Apirion proposed a model of mRNA decay in bacteria (Apirion, 1972, 1973). This model postulates that the initiating event is an endonucleolytic cleavage, followed by rapid scavenging of the newly-created intermediates by 3'->5' exonucleases (Figure 1.1). The identity of the initiating endonuclease, mechanisms for dealing with stable secondary structures, an explanation of the 5'->3' directionality of mRNA decay, of the different susceptibilities of various RNAs to degradation, or of the all-or-none phenomenon were lacking from this model. Despite these short-comings, Apirion's model has proved generally correct, and is incorporated in newer models. 5 .3' ^1 . Endonucleolyt ic c leavage ^2. Exonucleolyt ic scaveng ing K Figure 1.1 Apirion Model of mRNA Decay David Apirion envisaged a model of RNA decay (Apirion, 1972, 1973). The initiating event (1.) is an endonucleolytic cleavage within the RNA. This is followed by removal of the two intermediates created by 3'->5' exonucleases (2.). The identity of specific enzymes, reasons for differences in stability of various RNAs, and mechanisms for dealing with stable secondary structures were not explained by this model. 6 1.4.3 Factors affecting mRNA decay What are the factors affecting mRNA decay? First, there are RNA determinants, such as substrate structure and sequence. Cleavages are not random, as RNAs of identical length can exhibit quite varied half-lives (Blundell et al., 1972; Nilsson et al., 1984). Hairpins at the 5'-end stabilize transcripts (Chen et al., 1991; Emory et al., 1992; Bouvet & Belasco, 1992; Hansen et al., 1994; Arnold & Belasco, 1998). Similarly, 3' stem-loops protect transcripts from decay (Nossal & Signer, 1968; Mott et al., 1985; Belasco et al., 1986; Wong & Chang, 1986; Mackie, 1987; Newbury et al., 1987a, 1987b; Chen et al., 1988; Plamann & Stauffer, 1990; McLaren et al., 1991; Petersen, 1991; Blum et al., 1999). Most mRNAs in E. coli end in such a structure, whether from a rho-independent transcription terminator or a repetitive extragenic palindrome (REP) sequence (Washio et al., 1998). In fact, REP sequences are found in the extragenic region of about 25% of transcriptional units (Higgins et al., 1982; Stem et al., 1984; Gilson et al., 1984). Finally, the phosphorylation state of the 5'-end of a transcript affects its decay (Lin-Chao & Cohen, 1991; Lin-Chao et al., 1994; Mackie, 1998, 2000; Tock et al., 2000; Spickler et al., 2001). Monophosphorylated transcripts can be up to 30 times more susceptible to RNases than triphosphorylated ones. Given that RNAs are initially synthesized with a triphosphate at the 5'-end, intact transcripts are more stable than their partially degraded counterparts (which acquire a 5' monophosphorylated end as a result of cleavage), explaining the all-or-none phenomenon. 7 There are also factors external to the RNA that control the rate at which it is degraded. Translational efficiency seems to be correlated with stability: the more an mRNA is translated, the more stable it is likely to be (reviewed in Petersen, 1993; Dreyfus & Joyce, 2002). Growth conditions and environmental signals also influence transcript stability (Section 1.7). Finally, the relative abundance and specificities of RNases and other proteins involved in RNA degradation determine the rate with which an RNA is degraded (Jain etal., 2002). 1.4.4 Current model One of the unforeseen problems in Apirion's Model is that all of the 3'->5' exonucleases involved in mRNA degradation are single-strand-specific. As most E. coli transcripts end in a stable stem-loop structure (Section 1.4.3), their 3'-ends block the action of exonucleases. While it may be important to protect mRNAs for short periods to enable them to carry out their function, mRNAs must ultimately be destroyed, so these 3' barriers to decay must be overcome. This can be achieved in one of two ways. The exonucleases can be helped by polyadenylation of the 3'-end, which provides a ramp, or "toehold" enabling the exonucleases to efficiently bind the 3'-end and proceed processively toward the stem-loop (He etal., 1993; Xu etal., 1993; Hajnsdorf etal., 1994, 1995; O'Hara etal., 1995; Xu & Cohen, 1995; Coburn & Mackie, 1996a, 1998; Haugel-Nielsen 8 et al., 1996; Cao et al., 1997; Blum et al., 1999; Mohanty & Kushner, 1999, 2000a). Repeated cycles of polyadenylation, combined with hairpin "breathing" will eventually allow the exonucleases to progress beyond the structure. Alternatively, an RNA helicase can unwind the stem-loop, yielding single-strands that can be easily degraded by the exonucleases (Py et al., 1996, Coburn & Mackie, 1998; Vanzo era/., 1998; Coburn et al., 1999). Figure 1.2 summarizes the current prevailing view of mRNA decay, and names the enzymes known to be involved. 1.5 Enzymes of mRNA decay There are 15 known ribonucleases in E. coli, seven of which operate on mRNA: RNase E, RNase G, RNase P, RNase II, RNase III, polynucleotide phosphorylase (PNPase), and oligoribonuclease (Deutscher, 1988; Goodrich & Steege, 1999; Ghosh & Deutscher, 1999; Nicholson, 1999). Two of these, the endonuclease RNase E, and the exonuclease PNPase associate together with the DEAD-box RNA helicase RhIB, and the glycolytic enzyme enolase in a complex called the degradosome (Py et al., 1994; Carpousis et al., 1994; Py et al., 1996). This complex is thought to be responsible for the degradation of most RNAs in E. coli. 1.6 The E. c o l i RNA degradosome Many important processes in the cell are carried out by large multi-component protein machines, like the proteasome, spliceosome, ribosome, and 9 Figure 1.2 Current Model of RNA Degradation 1. The rate-limiting step is an endonucleolytic cleavage by RNase E. It is believed that RNase E first recognizes or "senses" the phosphorylation state of the 5'-end of the mRNA (it prefers 5'-monophosphorylated RNAs). 2. Cleavage by RNase E creates a new 3'-end, as well as an intermediate RNA with a 5'-monophosphate. This intermediate is more susceptible to further RNase E cleavage than an uncut RNA. 3. All 3'-ends are digested by 3'->5' exonucleases (RNase II and/or PNPase). If stable secondary structures are encountered, the exonucleases stall. 4. Two mechanisms can help stalled exonucleases. 4.a. PAPI can add a poly(A) tail to the hairpin, thereby providing a "ramp" for the exonuclease. Combined with hairpin breathing, the stem-loop can eventually be overcome. 4.b. In the degradosome, RhIB can unwind the stem-loop and give the single-strand product to PNPase for digestion. RhIB activity requires a 3' single-strand extension (Blum e t al., 1999, Coburn e t al., 1999), which may be provided by PAPI, PNPase (Section or the natural 3'-end of the substrate. 10 the nuclear pore and RNA polymerase complexes. The E. coli RNA degradosome was discovered in 1994 during attempts to purify RNase E (Py et al., 1994; Carpousis et al., 1994), and immediately generated considerable interest as the potential functional implications of grouping enzymes of mRNA decay together in a molecular machine were explored (Figure 1.3; Py et al., 1996; Alberts, 1998). At its core is the major endonuclease of E. coli, RNase E/Rne, which provides a scaffold for the assembly of other proteins (Figure 1.4). The other main components of this complex are PNPase/Pnp, RhIB, and enolase (Py et al., 1996, Carpousis et al., 2001; Carpousis, 2002). Other proteins co-purify with these four enzymes, although they differ based on the purification method, and their recovery tends to be sub-stoichiometric. These include polyphosphate kinase (PPK), poly(A) polymerase I (PAPI), the protein chaperones DnaK and GroEL, the ribosomal protein S1 , the cold-shock protein CspE, the exonuclease impeding factor (EIF), and possibly the RNA polymerase p and P' subunits (Sohlberg et al., 1993; Causton et al., 1994; Miczak et al., 1996; Blum etal., 1997; Raynal & Carpousis, 1999; Feng etal., 2001; Marchand et al., 2001). Although these "auxiliary proteins" could conceivably be functionally important, the prevailing view is that they are contaminants. The presence of RNA (mRNA, rRNA, small regulatory RNA) has also been detected in this complex (Miczak etal., 1996; Bessarab etal., 1998). 11 Figure 1.3 Proposed model of RNA-degradosome interactions The degradosome contains at least 3 proteins capable of interactions with RNA: RNase E (long, two-part, shaded object), PNPase (the "Pacman"), and RhIB (triangle). PNPase, RhIB, and enolase (circle) interact with the C-terminal half of RNase E . Bearing in mind the proposed role of each of these enzymes in mRNA degradation (Figure 1.2), an RNA molecule was positioned on the complex to reflect possible interactions with degradosome proteins. The 5' end is retained in the proposed "phosphate-binding pocket" (5'-end recognition site) of RNase E, which is thought to be in the N-terminal part of the protein (Spickler era/., 2001). A stem-loop structure is placed over RhIB. The activity of this helicase generates single-stranded RNAs which are "fed" to PNPase, positioned nearby and held in the correct orientation by virtue of its association with RNase E. 12 RNase E (Rne) 1061 ^ Endonucleolvtic Activity v . ^498 S1 Pomain sip_ 35^~ 25 ARRBD 608^^35 Self-Interaction 500 528 S^elf-lnteractiQn 752 501 509 498 RhIB 7 3 4 ^ 7 3 8 Enolase 739^ ^ 4 5 ^ PNPase ^ PAPI DnaK 844 843 1045 844 S1 765 Figure 1.4 Regions of Interest on RNase E Primary Sequence The catalytic region of RNase E (Cormack etal., 1993; Taraseviciene etal., 1995; Kido etal., 1996; McDowall & Cohen, 1996; Huang etal., 1998), its S1 and ARRBD domains (Cormack et al., 1993), as well as sites of interaction for self-assembly and for association with the degradosomal proteins RhIB, enolase, PNPase, PAPI, DnaK and S1 are mapped onto the RNase E primary sequence (Kido etal., 1996; Kaberdin et al., 1998; Vanzo et al., 1998; Raynal & Carpousis, 1999; Feng etal., 2001; Callanghan etal., 2003). Numbers at the ends of each arrow indicate which RNase E amino acids are implicated in the binding of each protein. 13 1.6.1 Purification The degradosome can be purified by different means. Some laboratories employ a purely biochemical approach (Py etal. 1994; Carpousis et al., 1994; Py et al., 1996; Coburn & Mackie, 1998), while others prefer to use artificial tags to aid in the purification (Miczak et al., 1996; Blum etal., 1997). The interactions between components are strong enough to enable its reconstitution in vitro from purified components: mixed RNase E, PNPase, and RhIB re-assemble into a fully functional "minimal degradosome" (Coburn et al.. 1999). The interactions between proteins have been studied, mapped, and confirmed using a variety of methods (yeast and E. coli two-hybrid, far-Western analysis, surface plasmon resonance, immunoprecipitation, functional assays, e tc . ) , all of which reinforce the existence of this complex in vivo. 1.6.2 RNase E The protein RNase E/Rne is a 1061 amino acid protein encoded by the rne/ams/hmp/smbB gene (Cormack et al., 1993; Casaregola et al., 1994). Although its mass is 118,000, it migrates as a 180-kDa protein on an SDS-PAGE gel due to its three proline-rich regions (Casaregola et al., 1992; McDowall & Cohen, 1996; Cohen & McDowall, 1997). It bears some sequence homology to myosin and other contractile proteins, and both myosin and RNase E cross-react with some antibodies raised against the other protein (Casaregola et al., 1992; McDowall et al., 1993). 14 Recent biophysical studies suggest that RNase E is a homo-tetramer, and that association of the monomers is required for catalytic activity of the protein (Mackie et al., 1997; Vanzo et al., 1998; Briant et al., 2003; Callaghan et al., 2003). Multimerization domains are present in the N-terminal part of the protein (Vanzo et al., 1998; Callaghan et al., 2003). Each tetramer can bind up to four RNA molecules. There are two suspected RNA binding sites on RNase E. The first, present between residues 35 and 125 is an S1 or OB-fold domain, which has homology to each of the six repeated segments in the middle and C-terminus of ribosomal protein S1 (Bycroft era/. , 1997). Whether this putative single-strand RNA-binding domain is required for cleavage is unresolved, although two thermolabile RNase E mutants have been mapped to this region (Diwa et al., 2002; Miao et al., in preparation; Schubert et al., in preparation), and the S1 domain of RNase E has been implicated in 5'-end recognition (Mackie, 1998; Jiang et al., 2000; Tock et al., 2000; Miao et al., in preparation). The second is an arginine-rich RNA binding domain (ARRBD), consisting of a strongly basic region (greater than 40% arginine) flanked by proline-rich sequences located between residues 608 and 635 (Cormack etal., 1993; Taraseviciene etal., 1995; McDowall & Cohen, 1996). This domain, with similarity to the RNA binding site of human U1 snRNP protein, also has RNA-binding properties in vitro (McDowall & Cohen, 1996, Taraseviciene et al., 1995; Leroy etal., 2002). However, as for the S1 domain, the essentiality of this domain for enzymatic activity is the subject 15 of some debate (Taraseviciene et al., 1995; McDowall & Cohen, 1996; Ow etal., 2000; Redko etal., 2003). Electron microscopic studies have revealed that RNase E is a membrane-bound protein (Liou et al., 2001). This association is mediated by its first 602 amino acids. This is consistent with a requirement for detergents during the purification of RNase E or degradosomes (Carpousis et al., 1994), and the fact that RNase E fractionates with the inner membrane (Miczak etal., 1991). Catalytic activity RNase E is the principal intracellular endoribonuclease of E. coli. It catalyzes the rate-limiting cleavage in the degradation of most mRNAs, and is involved in the processing of tmRNA, tRNA and rRNA (Kuwano et al., 1977; Ghora & Apirion, 1978; Misra & Apirion, 1979; Ono & Kuwano, 1979; Ray & Apirion, 1981; Arraiano et al., 1988; Mudd et al., 1990a, 1990b; Babitzke & Kushner, 1991; Taraseviciene et al., 1991; Melefors & von Gabain, 1991; Li et al., 1999; Lin-Chao et al., 1999; Wachi et al., 1999; Klein & Evguenieva-Hackenberg, 2002; Li & Deutscher, 2002; Ow & Kushner, 2002). RNase E's catalytic activity resides within the first 498 amino acids of the protein (Cormack et al., 1993; Taraseviciene et al., 1995; Kido et al., 1996; McDowall & Cohen, 1996; Huang etal., 1998; Jiang etal., 2000). RNase E is a single-strand-specific endonuclease with only limited sequence specificity, which can best be described as AU-rich regions at least eight nucleotides from the 5'-end (Mackie, 1991, 16 1992; Ehretsmann et al., 1992; Mackie & Genereaux, 1993; Lin-Chao et al., 1994; McDowall etal., 1994, 1995; Mackie etal., 1997; Kaberdin, 2003; Redko et al., 2003). The activity of RNase E is strongly stimulated on 5' monophosphorylated RNAs compared to 5' triphosphorylated ones (Lin-Chao & Cohen, 1991; Lin-Chao et al., 1995; Mackie, 1998, 2000; Tock et al., 2000; Spickler et al., 2001). Cleavage by RNase E releases a 5' monophosphorylated decay intermediate, which is preferred by the enzyme. This property therefore ensures that the enzyme works "processively" on a substrate rather than initiating cleavage on intact ones. RNase E has a paralogue in E. coli named RNase G/Caf A. RNase G is over 49.5% similar (34% identical) to the first 498 amino acids of RNase E (Dahlberg etal., 1978; Wachi etal., 1991, 1997, 1999; McDowall et al., 1993; Okada et al., 1994; Li & Deutscher, 1999; Tock et al., 2000; Umitsuki et al., 2001). While the two proteins are catalytically very similar, it appears that RNase G cannot substitute for RNase E in the cell, despite an earlier report that it might (Jiang et al., 2000; Lee et al., 2002; Briant et al., 2003; Ow et al., 2003). Homologues of the N-terminal catalytic domain have been identified in archaeal, plastid, and other bacterial genomes (Reith & Munholland, 1995; Franzetti et al., 1997; Hagege & Cohen, 1997; Kaberdin et al., 1998; Kokoska & Steege, 1998; Coburn & Mackie, 1999; Aravind & Koonin, 2001; Baginsky et al., 2001; Lee & Cohen, 2003). 17 RNase E's activity is essential for cell survival (Gegenheimer et al., 1977; Ghora & Apirion, 1978). A minimum of 10-20% of the activity of the N-terminal domain of RNase E is required to maintain viability (Jain & Belasco, 1995; Taraseviciene et al., 1995; Kido et al., 1996; Jain et al., 2002). Early speculations focused on either rRNA processing or the degradation of critical mRNAs, but tRNA processing is the most likely reason that this enzyme is essential (Li & Deutscher, 2002; Ow & Kushner, 2002, although see Deana & Belasco, 2004). RNase E is not a very abundant protein (Nierlich & Murakawa, 1996). It is estimated that in exponentially growing cells, there are roughly 320 to 960 molecules of RNase E (Kido et al., 1996). This number is tightly maintained by a process of autoregulation (Mudd & Higgins, 1993; Jain & Belasco, 1995; Diwa et al., 2000; Sousa et al., 2001). Meanwhile, the cell produces approximately 16,500 RNase E substrates every minute2. Under normal growth conditions, RNase E appears to be near substrate saturation, as modest increases in the amount of a substrate RNA can easily overwhelm RNase E's ability to cleave it (Sousa et al., 2001). This indicates that RNase E's turn-over rate is approximately 1-4 sec 3. This estimate is supported by the recent kinetic studies 2 This assumes that there are approximately 1,315 pre-rRNAs (each with at least three RNase E sites), 12,200 tRNA precursors (each with a minimum of one RNase E site), and 180-480 mRNA (which are substrates for RNase E) produced every minute in the cell (Sousa ef al., 2001). The number of rRNA and tRNA is based on the assumption that the approximately 26.3 X 103 ribosomes and 244 X 103 tRNAs present in the cell need to be replicated each doubling time, which is about 20 min (Bremer & Dennis, 1996). 3 If there are 320-960 molecules of RNase E catalyzing the cleavage of 16,500 substrates each minute, then a single protein can cut 17-51 RNAs each minute. This means that a cleavage reaction occurs every 1 -4 seconds. 18 of Redko and coworkers (2003), who calculated that RNase E's K c at value is 1.1-1.4 sec"1, depending on substrate. This value is similar to those obtained for other phosphotransferases (Kanaya et al., 1990; Waters & Connolly, 1994; Baldwin et al., 1999). This rate of catalysis is rather slow, but is not unexpected of an enzyme that stays bound to its substrate after cleavage for repeated cycles of catalysis (i.e. once RNase E makes the initial cleavage, it prefers to continue cutting the intermediate instead of attacking an uncut RNA). Scaffolding region The latter half of the protein, though not involved in enzyme catalysis, harbours distinct binding sites for each of the other degradosomal proteins, and hence has been dubbed the "scaffolding domain" (Kido et al., 1996; Blum et al., 1997; Kaberdin etal., 1998; Vanzo etal., 1998; Raynal & Carpousis, 1999; Feng et al., 2001). Figure 1.4 illustrates the domain organization and shows the binding sites for each degradosome protein. The scaffolding domain is not well conserved in other organisms (Kaberdin etal., 1998; Lee & Cohen, 2003) 1.6.3 PNPase The protein PNPase is a 711 amino acid homo-trimeric protein with a mass of roughly 258-kDa (Valentine et al., 1969; Regnier era/., 1987; Grunberg-Manago, 1989; Symmons et al., 2000). Its predicted monomeric size is 78-kDa, but it migrates as a 86-kDa peptide on an SDS-PAGE gel (Soreq & Littauer, 1977; Regnier et 19 al., 1987). Depending on the purification protocol, PNPase sometimes co-purifies with a dispensable (3 subunit, which has been identified as enolase (Portier, 1975; Carpousis et al., 1994; Py et al., 1996). However, E. coli and yeast two-hybrid analyses, far-Western blots, and co-immunoprecipitation experiments were all unable to find a direct physical association of the two proteins, suggesting that the initial co-purification probably resulted from the presence of contaminating partially degraded degradosomes in the PNPase preparation (Carpousis etal., 1994; Py etal., 1996; Vanzo etal., 1998; Kuhnel & Luisi, 2001; Liou etal., 2002). Based on immunogold labeling and electron microscopy, PNPase is localized to the cytoplasm and is not enriched in cell membranes (although it is present there, whereas a strictly cytoplasmic control is not) (Py et al., 1994; Liou et al., 2001). This is perhaps not surprising in view of the fact that only 10-20% of cellular PNPase is thought to associate with the degradosome (there is an excess of PNPase compared to RNase E) (Carpousis et al., 1994; Liou et al., 2001). The PNPase sequence contains two RNase PH-like domains (named PH and PH') which encompass the catalytic core and give the protein a duplicated symmetry (Bateman etal., 2000; Symmons et al., 2000). Once assembled into a trimer, PNPase is therefore a "trimer of dimers" (Symmons et al., 2000). The crystal structure of Streptomyces antibioticus PNPase has been resolved to 2.6 20 A resolution, revealing a ring-shaped trimer. In addition, PNPase contains two single-stranded RNA-binding domains. The S1 domain lies between residues 619 and 691 (Godefroy-Colburn & Grunberg-Manago, 1972; Regnier etal., 1987; Bycroft et al., 1997; Littauer & Grunberg-Manago, 1999). As mentioned in Section, S1 is a single-strand RNA-binding clasp that can bind RNA with a certain degree of sequence specificity (Ringquist et al., 1995; Bycroft et al., 1997). The S1 domain of PNPase is suspected of participating in interactions with the C-terminal tail of RNase E, and thus to be important in the assembly of the degradosome (Miao et al., in preparation). The KH domain (with similarity to hnRNP K protein) is found between residues 557 and 591 (Gibson et al., 1993; Mattaj, 1993; Symmons et al., 2000; Grishin, 2001). Although they were not resolved in the crystal structure, the KH domains have been modeled to lie above the central channel, and the S1 domains facing outward from the channel (Symmons etal., 2000). Catalytic activity PNPase is a widely conserved polynucleotide nucleotidyl transferase, catalyzing the reversible reaction (Littauer & Grunberg-Manago, 1999; Raijmakers etal., 2002): (NMP)n + Pi o (NMP)n-i + NDP Given the high (roughly 10 mM) phosphate concentration in E. coli cells, and a K m for phosphate of 1.8 mM, it is presumed that the exonucleolytic 21 (forward) reaction is favoured (Singer & O'Brien, 1962; Shulman et al., 1979). Thus PNPase is considered a single-strand-specific 3'->5' phosphorolytic "exonuclease4" (Littauer & Kornberg, 1957; Grunberg-Manago, 1963; Godefroy-Colburn & Grunberg-Manago, 1972; Littauer & Soreq, 1982). Its activity is strongly inhibited by RNA secondary structure (stem-loops containing more than six G-C base pairs (Spickler & Mackie, 2000)). A single-strand extension of roughly 10 nucleotides is required for efficient binding and for enzymatic processivity (Nossal & Singer, 1968; Littauer & Soreq, 1982; Newbury et al., 1987; Plamann & Stauffer, 1990; McLaren etal., 1991; Cannistraro & Kennell, 1994; Coburn & Mackie, 1996b). PNPase activity is also dependent upon the presence of a divalent cation such as Mg + + , which has lead to the suggestion that it cleaves nucleotides using a two-metal-ion mechanism where phosphate rather than hydroxyl ion is the attacking nucleophile (Burgers & Eckstein, 1979; Steitz & Steitz, 1993). PNPase mutants are unable to grow below 30°C (Luttinger et al., 1996; Grunberg-Manago, 1999). At more elevated temperatures, PNPase activity is not essential in the cell, although a mutant in the genes for both PNPase and RNase II, a 3'->5' hydrolytic exonuclease is lethal (Kinscherf & Apirion, 1975; Donovan & Kushner, 1986). This suggests that RNase II and PNPase are functionally redundant, although there is evidence that RNase II cannot handle secondary structures as well as PNPase (Mackie, 1989; Guaneros & Portier, 4 For the purposes of this discussion, PNPase will be referred to as an exonuclease, although this is not strictly correct. Nucleases are defined as hydrolytic enzymes, whereas PNPase is 22 1991; McLaren et al., 1991; Pepe et al., 1994; Braun et al., 1996; Coburn & Mackie, 1996a; Mohanty & Kushner, 2000a; Kushner, 2002). As up to 90% of the exonucleolytic activity in E. coli is hydrolytic, it has been proposed that RNase II is the major exonuclease in E. coli (Chaney & Boyer, 1972; Deutscher & Reuven, 1991). However, Kushner has pointed out that if RNase II exhibits a different specificity than PNPase (as indeed there is some evidence (Mackie, 1989; Guaneros & Portier, 1991; McLaren etal., 1991; Pepe etal., 1994; Braun et al., 1996; Coburn & Mackie, 1996a; Mohanty & Kushner, 2000a; Kushner, 2002)), for instance preferring rRNA, and keeping in mind that mRNA makes up less than 10% of the RNA mass, then it is still possible that PNPase is the major exonuclease of mRNA decay (Kushner, 2002). Using PNPase is more energetically favourable for the cell, as it creates NDPs, instead of NMPs (Deutscher & Reuven, 1991). In fact, Bacillus subtilis, which normally inhabits energy-poor environments, contains only a phosphorolytic 3'->5' exonuclease (Deutscher & Reuven, 1991; Higgins etal., 1993; Wang & Bechhofer, 1996). The presumption that PNPase functions solely as an exonuclease in the cell was recently revised, as evidence emerged that PNPase might also exhibit polymerization activity in vivo (Reuven et al., 1997; Li (Q.S.) et al., 1998; Mohanty & Kushner, 2000b; Yehudai-Resheff et al., 2001; A. Prud'homme Genereux, unpublished results). It has been found that even at a concentration of 20 mM phosphate, addition of 1 mM ADP blocks the forward (exonuclease) reaction (Yehudai-Resheff et al., 2001). This finding has interesting phosphorolytic. 23 consequences for mRNA decay. If the microenvironment around PNPase contains a sufficiently high concentration of ADP, then the enzyme could revert to its polymerase activity, despite the high overall phosphate concentration in the cell (Mohanty & Kushner, 2000b). Thus, in the absence of a stable stem-loop, PNPase (in the degradosome) will remove single-stranded nucleotides and create NDPs. Once the degradosome encounters a stable stem-loop, RhIB will hydrolyze ATP to ADP in an effort to unwind the structure (Section 1.4.4). Both PNPase and RhIB will be creating a local and transient pool of ADP surrounding the degradosome, which could trigger the switch to polymerase activity in PNPase. At these difficult stem-loops, PNPase would use ADP to add a 3' tail, which is known to facilitate its activity and make the enzyme more processive. Once PNPase has exhausted the local surplus pool of ADP, it would revert to its exonucleolytic activity and attempt to degrade the weakened stem once more (Figure 1.5). 1.6.4 RhIB The protein BNA helicase-]ike protein B (RhIB) was identified in a search of the E. coli genome for DEAD-box helicase sequences (Section 4.1.5; Kalman et al., 1991). It is 421 amino acids long, and has a molecular mass of 48-kDa. Whether rhIB is an essential gene is currently the subject of some debate. The first report on this matter claimed that the rhIB gene was only essential in certain genetic backgrounds (Kalman et al., 1991). However, later studies found that this gene 24 C D 1. 2. 3. 4. 5. 6. + NDPs NDPs —\ Figure 1.5 Environmental determinants of PNPase activity PNPase catalyzes the reversible reaction: (NMP) n + Pi * (NMP)n.., + NDP. The amount of NDP present in the microenvironment surrounding PNPase is proposed to control whether PNPase works as a polymerase or as a nuclease. 1. PNPase, in the context of the degradosome, encounters an RNA, and proceeds to remove 3' nucleotides by phosphorolysis, (2.) creating a pool of NDPs around the degradosome. 3. The presence of a stable stem-loop induces RhIB to unwind the hairpin, creating A D P (by A T P hydrolysis) in the process. 4. The high concentration of NDPs in the immediate vicinity of PNPase causes the protein to catalyze the reverse reaction, i.e. add nucleotides to the 3' tail. 5. After a period, NDPs are depleted from the area, and PNPase reverts to its exonucleolytic mode of action. 6. The 3' extension provides PNPase with a ramp that may promote its processivity. This, combined with the action of RhIB, helps PNPase overcome the stem-loop. Repeated cycles of polyadenylation/exonucleolytic action may occur on a given substrate. 25 is essential in all backgrounds (Py et al., 1996; Blum et al., 1997). This issue is complicated further by the recent claim of Dr. A.J. Carpousis (Laboratoire de microbiologie et genetique moleculaire, Toulouse) of the creation of a viable rhIB deletion mutant (personal communication; Khemici & Carpousis, 2004). The debate on RhIB is not limited to the status of its essentiality: one report claims that RhIB remains monomeric in solution (Callaghan et al., 2003), while yeast two-hybrid and plasmon resonance analysis on BIAcore apparatus have found that the protein can interact with itself (Vanzo et al., 1998; Liou et al., 2002). There appears to be a roughly equimolar amount of RNase E and RhIB in the cell, and RhIB co-localizes to the inner cell membrane with RNase E (Liou et al., 2001). This co-localization appears to be dependent upon an association with RNase E (Liou et al., 2001). The RhIB protein is the target of kinases and is phosphorylated in vivo (Marchand etal., 2001). ATPase activity By itself, RhIB has no ATPase activity. However, RhIB can bind RNA in vitro, and the presence of RNA stimulates its ATPase activity over three-fold (Py et al., 1996). An additional 15-fold stimulation is observed in the presence of the RNase E region to which it binds (RNase E amino acids 628-843) (Py et al., 1996; Vanzo etal., 1998). This stimulated activity saturates at about one RNase E molecule for every RhIB monomer. It has been suggested that RhIB's association with RNase E folds the arginine-rich and randomly-coiled C-terminus 26 of RhIB into a functional RNA-binding region, thereby activating the enzyme (Vanzo et al., 1998). Helicase activity RhIB can unwind short RNA duplexes in an ATP-dependent fashion, and therefore appears to be a true helicase (Vanzo et al., 1998; Liou et al., 2002). This activity is observable in the absence of RNase E, but requires a 500-fold excess of RhIB over RNA to be detected (Liou et al., 2002). In vitro assays have demonstrated that, like its ATPase activity, RhIB's helicase activity is dependent upon an association with RNase E (Coburn et al., 1999). Indeed, this activity has been shown to assist in the degradation of stable terminal stem-loop structures by PNPase (Py et al., 1996; Coburn et al., 1999). RhIB cannot facilitate RNase ll's ability to degrade hairpins, and functions only in degradosomes (Coburn et al., 1999). Association of RhIB with PNPase and poly(A) polymerase I Initial investigations into RhIB's association with the degradosome revealed a physical interaction between RNase E and RhIB (Vanzo et al., 1998). The possibility that RhIB might interact with PNPase was tested by yeast two-hybrid analysis, far-Western, and both in vivo and in vitro immunoprecipitation experiments, but no such association was ever uncovered (Vanzo et al., 1998; Coburn et al., 1999; Ow et al., 2000). However, a recent study that used E. coli two-hybrid analysis and surface plasmon resonance on a BIAcore biosensor 27 detected a weak interaction between residues 194 and 421 of RhIB and PNPase (Liou et al., 2002). This interaction was four times weaker than the RNase E-RhlB or the RhIB-RhIB associations. Unfortunately, the experiment did not control for a possible RNase E or RNA bridge. As RhIB was absent in the immunoprecipitate of an RNase E mutant lacking the PNPase binding site (Ow et al., 2000), these results hint that RhIB may require the assistance of PNPase to associate with RNase E. Despite these pieces of evidence, given that there is a roughly equimolar (and by inference, stoichiometric) amount of RNase E and RhIB in the cell (Section 1.6.7), and that PNPase does not simulate RhIB's activity (Liou et al., 2002), it would appear that RhIB's affinity for PNPase is not physiologically relevant. A far-Western analysis of the interactions between PAPI and the degradosome unexpectedly revealed that PAPI can bind RhIB weakly in vitro, in the absence of an RNA bridge (Raynal & Carpousis, 1999). This interaction could not be confirmed by yeast two-hybrid analysis, but the addition of PAPI enhanced five-fold the ability of RhIB bound .to an RNase E fragment (amino acids 501-843) to hydrolyze ATP (Raynal & Carpousis, 1999). Perhaps this functional effect simply reflects the fact that RhIB needs a single-strand extension on an RNA duplex to have activity, rather than a physical interaction between it and PAPI (Blum etal., 1999). 28 1.6.5 Enolase Glycolytic enzyme Enolase (formerly called 2-phosphoglycerate dehydratase) is a very well conserved glycolytic enzyme necessary for both gluconeogenesis and glycolysis (Verma & Dutta, 1994). Mutations in its gene transform E. coli from a facultative anaerobe into an obligate aerobe (Irani & Maitra, 1977). It catalyzes the reversible dehydration reaction that converts 2-phosphoglycerate (also called glycerate-2-phosphate) into the energy-rich phosphoenolpyruvate. Enolase present in degradosomes has as much specific activity as purified protein (Py et al., 1996). It is estimated that only 5-10% of cellular enolase is associated with the degradosome (Py et al., 1996). Electron microscopy shows that although enolase is a cytoplasmic protein, it is often found near the inner membrane (Angiolella et al., 1996; Edwards et al., 1999; Liou et al., 2001). This might be expected of a degradosome protein present in excess over RNase E. A crystal structure of E. coli enolase has recently been solved (Kuhnel & Luisi, 2001). Enolase is a homo-dimer of two 46-kDa subunits combining to form a protein with a molecular mass of 92,000 (Spring & Wold, 1971; Dannelly & Reeves, 1988; Vanzo et al., 1998; Kuhnel & Luisi, 2001). The enzyme appears to change conformation when binding to its substrate (Kuhnel & Luisi, 2001). Interestingly, there is a structural similarity between a helical domain of PNPase and the N-terminal domain of enolase, but its significance is unknown (see Section (Symmons etal., 2000; Kuhnel & Luisi, 2001). 29 Possible roles in decay The functional role of enolase in the degradosome has baffled the field of mRNA decay since its discovery. Degradosomes can be reconstituted in vitro in the absence of enolase without any apparent lack of activity (Coburn et al., 1999). The prevailing hypothesis is that enolase is a regulator of degradosome activity, adjusting it to the cell's metabolic need (Kuhnel & Luisi, 2001). In the presence of glucose, enolase is phosphorylated on serine residues and its glycolytic activity is stimulated. This modification does not occur when cells are grown on acetate (Dannelly et al., 1989; Dannelly & Reeves, 1989; Marchand et al., 2001). This modification could serve as a regulatory signal for degradosomes. Indeed, the stabilities of a few mRNAs are known to fluctuate with growth medium composition (Kahn et al., 1982; Meyer & Schottel, 1991; Albertson & Nystrom, 1994; Woo & Lin-Chao, 1997; Barlow et al., 1998; Bernstein et al., 2002; Le Derout et al. 2002). Bulk RNA is also stabilized during anaerobic growth (Georgellis et al., 1993). Since phosphoenolpyruvate is used by pyruvate kinase to synthesize pyruvate (the branch point for aerobic and anaerobic growth), enolase may also be involved in sensing the aerobic conditions of the cell and promoting appropriate changes in degradosome activity. The structural similarity between the a-helical domain of PNPase (which is located between the PH and PH' domains) and the N-terminal domain of enolase 30 has prompted the suggestion that the domain may be involved in RNA binding (Symmons et al., 2000; Kuhnel & Luisi, 2001). Indeed, yeast enolase has demonstrated some ability to bind polynucleotides (Al-Giery & Brewer, 1992), and there are anecdotal reports of E. coli enolase binding weakly to RNA (Kuhnel & Luisi, 2001). However, this property was not observed by North-Western, UV cross-linking, or soaking the enolase crystals with RNA (Py etal., 1996; Kuhnel & Luisi, 2001). Nonetheless, there is some accumulating evidence that the section of RNase E that binds enolase (amino acids 738-845) is necessary for 5'-end-independent entry (called internal entry) of RNase E into mRNAs 5 (Lopez et al., 1999; Marchand et al., 2001; Leroy et al., 2002; G.A. Mackie, personal communication): Whether internal entry requires the portion of RNase E that binds enolase or enolase perse remains to be established. 1.6.6 Other components In addition to the main components of the RNA degradosome, the following proteins have been reported to co-purify with the degradosome by some experimenters. Polyphosphate kinase Polyphosphate kinase (PPK) is a peripheral membrane-bound tetramer of 80-kDa subunits that catalyzes the reversible polymerization of the y-phosphate 5 5'-end-independent entry (internal entry) of RNase E into a substrate is a concept that arose from the observation that RNase E can cleave (albeit slowly) RNA substrates that lack a free 5'-end (Mackie, 1998, 2000). The C-terminal half of RNase E appears instrumental in allowing RNase E to recognize and cleave a substrate without prior interaction with the 5'-end of the RNA. 31 of ATP (and sometimes GTP) into long chain (10 to many 100s) polyphosphates (Kornberg, 1995; Blum etal., 1997; Kornberg etal., 1999): Poly(Pi) + ADP o ATP Polyphosphates occur in E. coli cells at concentrations ranging from 0.1 to 50 mM, and accumulate in response to deficiencies in amino acids, phosphates, nitrogen, the stress of nutrient downshift, or high salt (Kornberg et al., 1999). In the cell, polyphosphates can replace ATP in some kinase reactions, serve as a phosphate reservoir, chelate metals (Mn + 2, Mg + 2 , Ca + 2 ) , buffer against alkalis, are used in making the bacterial capsule, are important for competence, can stimulate proteases, and have a regulatory role in physiological adaptation to growth, stress, and deprivation (Kornberg etal., 1999; Kuroda et al., 2001). As polyphosphate kinase is not essential, it is presumed to have only a facilitating role in the cell, although lack of this enzyme causes cells to die after only a few days in stationary phase (Kornberg etal., 1999). PPK has been shown to interact with RNase E through biochemical co-purification and co-immunoprecipitation. Although it does not co-purify in stoichiometric amount with other degradosomal components, it can account for up to 5% of the protein mass in a degradosome preparation (Blum et al., 1997). Deletion of ppk shows no effect on bulk mRNA decay, but the degradation of specific mRNAs seems to be altered (Blum et al., 1997). Although it was 32 hypothesized that PPK can accomplish this by altering the phosphorylation state of RNA, it is unable to transfer phosphates onto or from either the 5'- or 3'-ends (Blum et al., 1997). Alternatively, PPK might stimulate RNA decay through its enzymatic activity, as poly(Pj) and ADP are known to inhibit RNA decay (Kornberg, 1995; Blum et al., 1997). ADP is constantly produced by the activity of both PNPase and RhIB, and its local accumulation could inhibit the action of both enzymes. By promoting the removal of both poly(Pj) and ADP from the local environment of the degradosome, and by regenerating ATP for RhIB, PPK could ensure the proper function of degradosome proteins (Blum etal., 1997). Poly(A) polymerase I Poly(A) tails are much shorter on prokaryotic mRNA and stable RNA (14-60 As) than on eukaryotic mRNA (80-200 As) (O'Hara et al., 1995; Li (Z) et al., 1998). They are also only found on 0.01-2% of the bacterial mRNA population, probably due to their rapid removal by exonucleases (Cao & Sarkar, 1992; O'Hara etal. 1995; Mohanty & Kushner, 1999). PAPI is responsible for 95% of the polyadenylation activity in bacteria, PNPase making up the remainder (Cao & Sarkar, 1992; Kalapos etal., 1994; Mohanty & Kushner, 1999, 2000b). PAPI is a dispensable monomeric protein of 55-kDa that catalyzes the reaction (Masters et al., 1993): ATP + (NMP)m o PP| +(NMP) m + 1 33 While poly(A) tails protect RNAs from digestion in eukaryotes, they serve as a "toehold" for the processive action of exonucleases in bacteria (He et al., 1993; Xu et al., 1993; Hajnsdorf et al., 1994, 1995; O'Hara et al., 1995; Xu & Cohen, 1995; Coburn & Mackie, 1996b, 1998; Haugel-Nielsen et al., 1996; Cao etal., 1997; Korner & Wahle, 1997; Blum etal., 1999; Mohanty & Kushner, 1999, 2000a). Thus, they are involved in the degradation of structured RNA (Figure 1.2). It has been proposed that polyadenylation affects only the degradation of RNA intermediates, as the initiating event in the degradation of most mRNAs is an endonucleolytic cleavage (Coburn & Mackie, 1998; Goodrich & Steege, 1999; Hajnsdorf & Regnier, 1999; Mohanty & Kushner, 2000a; Dreyfus & Regnier, 2002). Whether PAPI is part of the degradosome is currently the subject of some debate. While PAPI was not detected in the biochemically purified degradosomes described by Py and colleagues (1994, 1996), it is present in at least the first two steps of the biochemical degradosome preparations of Coburn & Mackie (1998) (personal communication). Polyadenylation activity was also observed in degradosomes by Raynal et al. (1996), and the presence of PAPI confirmed by Western blotting. In these experiments, PAPI could be separated from the degradosome by sedimentation on a glycerol gradient containing 0.5M NaCI, suggesting that PAPI is an accessory factor that reversibly associates with the degradosome. PAPI has also been shown by far-Western and yeast two-hybrid analysis to physically interact with RNase E (Raynal & Carpousis, 1999). 34 A weak interaction with RhIB, as well as the ability to stimulate RhIB's ATPase activity 5-fold has also been noted. This stimulation requires the polyadenylation activity of PAPI, as a mutant cannot promote it. RhIB cannot increase PAPI's ability to synthesize poly(A) tails; however, there is some additional evidence for a functional interaction between PAPI and RNase E. PAPI may be sensitive to the phosphorylation state of the 5'-end (it prefers monophosphorylated RNAs), a property that could be conferred by an association with RNase E (Feng & Cohen, 2000). There is also evidence that the specificity of RNase E is altered in a p c n B (the gene for PAPI) mutant, although this may be an artifact of the unusal structure of the substrate used in these experiments (the 5'- and 3'-ends of RNAI are held in an unusually close conformation, possibly yielding a functional interaction between the two ends of this RNA that is atypical of mRNAs in general) (Xu & Cohen, 1995). S1 ribosomal protein Ribosomal protein S1 is part of the 30S ribosomal subunit. It interacts with sequences at the 5'-end of mRNAs and promotes interactions between the Shine-Dalgarno sequence of the mRNA and 16S rRNA. Some reports also claim that S1 is a poly(A) binding protein that interacts physically with both RNase E and PNPase in vitro (Kalapos et al., 1997; Feng et al., 2001). Since all three proteins contain S1 domains, it is possible that the far-Western results are due to antibody cross-reactivity. 35 DnaK DnaK is an abundant 69-kDa heat-shock protein with ATPase activity that interacts with DnaJ and GrpE to form a molecular chaperone that promotes the folding of nascent and newly synthesized proteins (Bardwell & Craig, 1984). The presence of DnaK has been reported in some degradosome preparations (Miczak et al., 1996; Blum er al., 1997; Vanzo et al., 1998). The association of DnaK with the degradosome is most likely due to an interaction with amino acids 509-844 of RNase E (Vanzo etal., 1998). An association with PNPase has been ruled out, although interactions with RhIB and enolase have yet to be studied. GroEL GroEL is a soluble chaperonin acting on proteins smaller than 60-kDa that are prone to rapid and irreversible aggregation (Georgopoulos & Ang, 1990; Baneyx, 1999). GroEL has weak ATPase activity that is necessary for peptide release. It is made up of 14 subunits of 57-kDa each that organize into two stacked doughnut-shaped rings. Although a heat-shock protein, it is required and constitutively expressed at all temperatures. Several indirect lines of evidence point to a potential interaction between GroEL and the degradosome. Even prior to the isolation of the complex, a genetic interaction between RNase E and GroEL was recognized. Overexpression of a fragment of GroEL could reverse the thermolabile growth phenotype of an RNase E mutant (Chandra et al., 1985). This is perhaps not 36 surprising given the nature of GroEL's function in refolding heat-denatured polypeptides, although RNase E is much larger than optimal substrates for GroEL. GroEL was found to associate with temperature-sensitive FLAG-tagged RNase E mutants, but not with the wild-type protein (Miczak et al., 1996). However, analysis of the protein content of a standard GroEL preparation revealed the presence of contaminating PNPase (Ybarra & Horowitz, 1996), and E. coli GroEL can interact specifically with yeast enolase to refold it, suggesting that E. coli enolase is a substrate as well (Kubo et al., 1993). There is only one published report of the co-purification of RNase E and GroEL (Sohlberg et al., 1993). Other laboratories have failed to detect such an interaction, and lack of GroEL in degradosomal preparations yields no discernible phenotype (Py et al., 1994, 1996; Carpousis et al., 1994). GroEL is known to cross-react with many different IgG molecules, which could account for its apparent interaction with RNase E in some experiments (Hajeer & Bernstein, 1993; Sohlberg et al., 1993; Alconada et al., 1994; Baginsky et al., 2001). Nonetheless, a modified GroEL has been implicated in the binding and protection of mRNA against RNase E action (Sohlberg etal., 1993; Georgellis etal., 1995; Vytvytska etal., 1998). EIF, CspE, and RNA polymerase p and P' subunits The exoribonuclease inhibitory factor (EIF) was reported once as a partly purified activity that co-purified with the degradosome and prevented the attack of PNPase at 3' stem-loops, despite its inability to interact with RNA (Causton et 37 al., 1994). The size of EIF suggests that it could be RNase II, whose activity will prevent a productive interaction of PNPase with RNA (Coburn & Mackie, 1996a). CspE, a constitutively expressed 7.3-kDa protein bearing similarity to CspA, the major cold shock protein of E. coli (Section 4.1.1; Bae et al., 1999), can bind to poly(A) tails to prevent their degradation by PNPase (Feng et al., 2001). The ability of CspE to bind poly(A) tails is not surprising, given its preference for AU-rich sequences (Phadtare & Inouye, 2000; Feng et al., 2001). Its RNA binding activity has been implicated in antitermination (Bae et al., 2000; Phadtare et al., 2002), but more significantly also in inhibition of RNase E and PNPase activity (Feng et al., 2001). There is a possible physical interaction between CspE and PNPase revealed by far-Western blotting (Feng etal., 2001). Like RNase E, CspE is a multicopy suppressor of mukB mutations (Hiraga et al., 1989). Finally, the RNA polymerase p and P' subunits have been co-immunoprecipitated with degradosomes while using RNase E antibodies to bring down the complex (Marchand et al., 2001). The authors of this paper claim that this is due to a non-specific cross-reactivity between the RNase E antibody and the RNA polymerase subunits. 38 1.6.7 Mass and stoichiometry of the degradosome The stoichiometry of the degradosome has been somewhat difficult to determine. Table 1.1 shows the stoichiometries that have been reported thus far. Given that until very recently the multimeric state of RNase E was unknown (Callaghan et al., 2003), all stoichiometries are given as ratios related to the amount of RNase E. There are two explanations that could account for the great variability observed between experiments. Either the variability is due to different experimental limitations for each of the techniques used to determine stoichiometries, or else the stoichiometry of the degradosome is variable. If the latter explanation is correct, then there are different forms of the degradosome, perhaps due to changing conditions in the cell. The fact that different experimenters report the presence of some dissimilar proteins in their degradosome preparations supports this hypothesis. Degradosome mass has also been difficult to establish. Various methods have placed it anywhere between 160-2,400-kDa (Ono & Kuwano, 1979; Carpousis et al., 1994; Py et al., 1994; Coburn & Mackie, 1999). The major discrepancies are between methods based on sedimentation and those based on gel filtration. As for stoichiometry, the variability of the mass may reflect differences in the experimental protocols used (or the presence of lipids or detergents which reduce the sedimentation rate), or the presence of alternate forms of the degradosome. 39 Molar Ratios Method & Reference RNase E PNPase RhIB enolase Biochemical purification Radio-densitometry (Carpousis et al., 1994) 1 1.5 0.6 1.7 Biochemical purification Staining and densitometry (Py etal., 1994, 1996) 1 2.7 1.6 2.7 Biochemical purification + immunopurification (note: RNase E is overexpressed) Silver staining and densitometry (Miczak etal., 1996) 1 0.9 0.9 1.8 Biochemical purification Staining and densitometry (Coburn & Mackie, 1999) 1 3 1 2 Immunopurification Staining + Western blot densitometry (Liou etal., 2001) Log phase 1 4 1 8 Stationary phase 1 3 1 22 Table 1.1 Degradosome Stoichiometry The left-most column lists the different purification methods used to isolate degradosomes, the procedure used to quantify the amount of each protein, and the reference of the article that reported these values. The next four columns are the ratios of each protein relative to the amount of RNase E reported for each degradosome preparation. 40 1.6.8 Advantages of association of enzymes into a degradosome As will be discussed in Section 1.6.11, the enzymes of mRNA degradation have been found associated in a complex in most of the organisms studied to date. Thus, there must be a selective advantage for the cell to do so. Generally speaking, enzyme complexes form for four main reasons: 1) to enhance the intrinsic catalytic activity of the enzymes, 2) to sequester reaction intermediates, 3) to permit regulation of metabolic fluxes, and 4) to sequester the active site and thereby reduce the rate at which incorrect substrates are cleaved (Hammes, 1981; van Hoof & Parker, 1999). There are several lines of evidence to support the assertion that the degradosome benefits the cell on all four levels. As it pertains to the enhancement of catalytic activity, there have been reports that the activities of RNase E and PNPase are coordinated, and that processing of an mRNA by one enzyme promotes the activity of the other (Xu & Cohen, 1995; Braun et al., 1996). PNPase has been suggested to dephosphorylate the 5'-end of RNA substrates in addition to removing nucleotides at the 3'-end, which would help regulate the 5'-dependent activity of RNase E (Sulewinski etal., 1989). In addition, a functional interaction has been observed between RNase E and PAPI (Section (Feng & Cohen, 2000), and RhIB and PAPI (Section (Raynal & Carpousis, 1999). However, the two best examples of catalytic enhancement are the ability of RNase E to dramatically stimulate the ATPase and helicase activities of RhIB (Sections 41 and, and the coordination of RhIB and PNPase activities necessary to degrade stable terminal structures (Section The all-or-none phenomenon of mRNA decay suggests that the cell does not allow the accumulation of decay intermediates. The deleterious nature of such intermediates or their products is probably attributable to their disturbance of cell metabolism. For example, an mRNA truncated towards its 3'-end could still encode a C-terminally truncated protein. Ribosomes would initiate translation, but the lack of stop codon (removed by RNase cleavage) would trap the ribosome, which would then require the assistance of tmRNA for release (Lin-Chao etal., 1999). Much energy would be wasted in this process. Moreover, the incomplete protein could be toxic. Following the initial endonucleolytic cleavage, an mRNA must be sequestered from the cellular pool of mRNAs, and passed on to PNPase and RhIB for complete degradation. Regulating a pathway is easier if the enzymes involved in this pathway are grouped together. This is indeed the best current hypothesis for the presence of enolase in the degradosome. Enolase may serve as a link between mRNA decay and the cell's energetic needs. Certain conditions, such as cold shock and resumption of translation after a block, are known to lead to rapid, general stabilization of mRNA (Goldenberg et al., 1996; Zangrossi et al., 2000; Beran & Simons, 2001; Mathy et al., 2001; Sousa et al., 2001; Yamanaka & Inouye, 42 2001). Such a response must occur by the rapid collective inhibition of all enzymes of mRNA decay. Given that RNase E catalyzes the initiating event in the degradation of most mRNAs, some factor must prevent PNPase and RhIB (even those present in the degradosome) from attacking intact substrates. Otherwise, degradation would proceed in a 3'->5' fashion. RNase E may position the RNA in such a way that the 3'-end becomes available to or promotes the cooperation of PNPase and RhIB (Figure 1.3) (Khemici & Carpousis, 2004). RNase E may also tag an RNA such that PNPase and RhIB recognize that the substrate is available for degradation. This idea has been proposed in the "Tethering model", where RNase E remains bound to a substrate after cleavage, and "delivers" it to PNPase and RhIB (Coburn & Mackie, 1999). Alternatively, the 5%end of an mRNA may be the only end available (for example, if the 3'-end is still being transcribed), and hence RNA is recognized by the only enzyme in the complex that can bind 5'-ends. RNase E selects which RNAs are to be degraded, and only those it recognizes can enter the decay pathway (Khemici & Carpousis, 2004). 1.6.9 Arguments against its existence in vivo Considerable evidence supports the existence and importance of the degradosome in vivo (Section 1.6.10). However, there are concerns that the degradosome is an artifact of in vitro experiments, that it does not represent the 43 major decay pathway for most mRNAs in E. coli, or that this complex is of relatively little importance given its lack of evolutionary conservation. First, the C-terminal region of RNase E, which contains binding sites for other degradosomal proteins, does not appear to be conserved, even among closely-related bacteria (Kaberdin et al., 1998). Indeed, the scaffolding domain (RNase E residues 593-1061) is dispensable for growth, and cells lacking it exhibit no growth defects (although they are less fit than wild-type strains, see Section 1.6.10) (Kido etal., 1996; McDowall & Cohen, 1996; Vanzo etal., 1998). Furthermore, bulk mRNA decay in strains containing this mutant form of RNase E proceeds at a rate that is only 1.7-times slower than in wild-type cells, and rRNA processing is unaffected (Lopez et al., 1999; Ow et al., 2000). In vitro, this phenomenon is also observed on some RNase E substrates (McDowall & Cohen, 1996). The second line of evidence is that although PNPase is heralded as the 3'->5' exonuclease with the greater ability to remove stable 3' RNA structures (by virtue of its collaboration with RhIB), pnp deletion strains are viable, and display few growth defects or effects on bulk mRNA decay (Portier, 1980; Donovan & Kushner, 1986; Lopez et al., 1999; however, see Mohanty & Kushner, 2003). RNase II must be able to compensate in its absence. 44 Finally, although the concentration of each degradosomal protein is very tightly regulated, their concentrations are not coordinated with each other, as might be expected of proteins associated together in a complex. In fact there is a large excess of PNPase and enolase (Carpousis et al., 1994; Py et al., 1996; Liou et al., 2001). In addition, the composition of the degradosome appears to be somewhat dependent upon the purification method used (Carpousis et al., 1994; Miczak et al., 1996; Py et al., 1996; Coburn & Mackie, 1999; Liou et al., 2001). 1.6.10 Arguments in favour of its existence i n v i v o The arguments listed above need to be addressed to fully comprehend mRNA decay in E. coli. However, despite these, it is generally believed that the degradosome is a real in vivo structure, given the following lines of evidence. First, the proteins of the degradosome physically associate. This is suggested by the fact that the degradosome can be purified using a variety of methods. There have been several biochemical purification methods published (Carpousis et al., 1994, 2001; Py etal., 1994, 1996; Coburn & Mackie, 1998). Similarly, different groups have immunopurified the degradosome using antibodies directed against either RNase E, PNPase, or a tag (histidine or FLAG) present on one of these proteins (Carpousis etal., 1994; Py etal., 1994; Miczak etal., 1996; Bessarab et al., 1998; Vanzo et al., 1998). This association is robust, as purified proteins reconstitute into a functional degradosome in vitro (Coburn et al., 1999). The protein regions involved in complex formation have been mapped to specific 45 residues, so the complexes purified were not due to aggregation (Vanzo et al., 1998). RNase E and RhIB also co-localize to the inner membrane, and 70-80% of RNase E, RhIB and enolase is located at or near the cytoplasmic membrane (Liou et al., 2001). In addition, in a mutant strain where RNase E is missing the RhIB binding site, RNase E is found at the membrane whereas RhIB is not (Liou et al., 2001). PNPase is more evenly distributed throughout the cell, although this might be expected of a cytoplasmic protein present in excess over its complexed form. In addition to physical interactions, the proteins of the degradosome functionally interact. To begin, monomers of RNase E do not appear to have any functional activity: a self-interaction is required (Callaghan et al., 2003). RhIB's activity is greatly increased by its association with RNase E (Py et al., 1996; Vanzo etal., 1998; Coburn etal., 1999). PNPase and RhIB work collaboratively to degrade stable 3' stem-loops, and cannot work in trans (i.e. they must both be bound to RNase E) in order to carry out this function (Coburn et al., 1999). RNase II cannot replace PNPase in this capacity (Coburn et al., 1999). There is also a report that the activities of RNase E and PNPase affect one another (Xu & Cohen, 1995). Given that RNase E mutants lacking the scaffolding domain are viable and exhibit only a two-fold stabilization of bulk mRNAs (Kido et al., 1996; Vanzo et al., 1998; Lopez et al., 1999; Ow et al., 2000), what are the roles, significances 46 and advantages of the degradosome? One hypothesis is that the degradosome is only necessary under conditions other than those used in the laboratory (37°C, LB broth). Indeed, while the generation time of RNase E mutants lacking various regions of the scaffolding domain was relatively unaffected at 37°C, much larger effects were observed at 44°C in minimal media (Lopez et al., 1999; Ow et al., 2000). Perhaps degradosome function becomes critical under different conditions, such as when RNA structures are stabilized, or when nutrient availability is limiting. Whatever the reason, the degradosome clearly confers a selective advantage to cells, as demonstrated in competition experiments between wild-type and mutants strains of RNase E (Leroy et al., 2002). Growing initially equivalent amounts of the two strains together for 100 generations, it was found that at the end of the experiment, RNase E mutants lacking the whole scaffolding region were present in 100,000-times lesser amount than wild-type, strains in which the RhIB and enolase binding sites were missing were present in 1,000-fold lesser amount, and those lacking a PNPase binding site were 100-fold less abundant than wild-type cells. Thus, bacteria in which degradosome formation is disrupted cannot grow as efficiently as wild-type cells and are quickly lost from the population pool. This is supported by the finding that although RNase E strains lacking residues 499-1061 are viable, the protein needs to be expressed eight-fold more than the wild-type protein (Jiang et al., 2000). The mutant protein can compensate, but is not as good a substitute as the wild-type protein. 47 Noticeable effects of removing the C-terminal half of RNase E on the abundance of specific mRNAs and their proteins have been reported (Kido et al., 1996; Nanbu-Wakao era/., 2000; Nogueira et al., 2001; Leroy etal., 2002; Dr. A.J. Carpousis, personal communication). In particular, the rate of decay of the mukB mRNA, RNAI, and REP sequences is strongly reduced in strains lacking the C-terminus of RNase E (Dr. A.J. Carpousis, personal communication; Khemici & Carpousis, 2004). In addition, RNase E needs its C-terminal half in order to effect its own auto-regulation (Jiang etal., 2000; Diwa et al., 2002; Leroy et al., 2002). It has also been observed that RNase E mutants lacking the scaffolding domain adopt filamentous cell shapes, and fail to form septa and divide (Leroy et al., 2002). While it may be tempting to conclude that RNase E has a role in cell division (Goldblum & Apirion, 1981; Faubladier et al., 1990; Cam et al., 1996; Kido etal., 1996; Wachi etal., 1997), particularly in light of its similarity to contractile proteins (Casaregola et al., 1992; McDowall et al., 1993), it is more likely that RNase E processes an mRNA (mukB?) with a critical role in this process (Cam etal., 1996; Kido et al., 1996). Thus, these studies support the idea that the C-terminal half of RNase E plays an important role in mRNA decay. 1.6.11 RNA decay machines in other organisms RNase E-based degradosomes appear to be conserved only among enteric bacteria. However, the grouping of various enzymes of mRNA decay into a complex appears to be preserved in most organisms. 48 RNase E-based degradosomes The existence of an RNase E-based degradosome has been confirmed in Rhodobacter capsulatus and Streptomyces coelicolor (Jager et al., 2001; Lee & Cohen, 2003). In Rhodobacter, the complex contains RNase E, PNPase (not a major component), two DEAD-box RNA helicases of 65- and 74-kDa (one is a homologue of CsdA (Section 4.1.4; Dr. R.W. Simons, personal communication)), the transcription termination factor Rho, possibly enolase (a protein with similarity to E. coli enolase, but not Rhodobacter enolase), and a 36-kDa unidentified protein (Jager et al., 2001). It has been purified biochemically and immunologically, and has functional activity. RNase E is the scaffold of this complex, which is 116-240-kDa. The complex exhibits a certain degree of heterogeneity, as the proteins associated with RNase E in different glycerol gradient fractions change. Most notably, the two DEAD-box helicases are found in alternate versions of the complex. The association of two DEAD-box helicases with this complex has been putatively attributed to the high G-C content of the Rhodobacter genome, which may increase the stability of mRNA secondary structures in this organism. The gram-positive bacteria Streptomyces coelicolor has a 140-kDa homologue of RNase E (Hagege & Cohen, 1997; Lee & Cohen, 2003). It has 49 endonucleolytic activity with a specificity resembling E. coli RNase E, and can complement an E. coli mutant lacking a copy of its gene. Amino acids 563-973 have 58% similarity (36% identity) to the N-terminal catalytic domain of E. coli RNase E. While the N- and C-terminal regions do not bear direct sequence similarity to any region of the E. coli protein, it does contain discrete acidic, arginine- and proline-rich regions similar to those in the C-terminal half of E. coli RNase E, and can bind S. coelicolor PNPase. Given that Haemophilus influenzae is a close evolutionary relative of Escherichia coli, that this organism has homologues of PNPase, RhIB, and enolase, and that there is extensive similarity between the RNase E sequences of the two organisms (even in the C-terminal region), it is likely that a degradosome exists in Haemophilus (Fleischmann et al., 1995; Kido et al., 1996; Cohen & McDowall, 1997; Kaberdin etal., 1998; Coburn & Mackie, 1999; Lee & Cohen, 2003). Salmonella typhimurium and Salmonella enterica also contain an RNase E homologue with a long C-terminal region that has the potential to form a degradosome-like complex, although none has been reported so far. Some bacteria contain a "truncated" RNase E homologue with only the catalytic region (Kaberdin et al., 1998; Aravind & Koonin, 2001; Baginsky et al., 2001; Lee & Cohen, 2003). This may reflect the fact that most prokaryotes do not use rho-independent terminators as a means of ending transcription, and hence their mRNAs do not contain stable 3' terminal hairpins that require special processing 50 by the degradosome (Washio et al., 1998). Thus, degradosomes may only be present in those organisms where REP or other structured RNAs are present. The exosome The exosome is a complex of enzymes involved in the 3'->5' decay of mRNA, rRNA, snRNA and snoRNA in eukaryotes (Mitchell et al., 1997; Decker, 1998; van Hoof & Parker, 1999; Mitchell & Tollervey, 2000). At its core are six phosphorolytic and three hydrolytic exonucleases. Both a nuclear and cytoplasmic "version" exist, each containing a different DEvH-box (related to DEAD-box) helicase (Section 4.1.5; de la Cruz et al., 1998; Jacobs et al., 1998; Allmang etal., 2000; Bousquet-Antonelli et al., 2000; Mitchell & Tollervey, 2000; Araki et al., 2001; Hilleren et al., 2001). The structural organization of the exosome is somewhat reminiscent of PNPase. The six phosphorolytic exonucleases arrange into a ring-structure that mirrors the "trimer of dimer" organization of PNPase (Section, and three additional subunits each containing an S1 RNA-binding domain are positioned on the outer surface of the ring (van Hoof & Parker, 1999; Aloy etal., 2002; Raijmakers etal., 2002; Estevez et al., 2003). No endonuclease is known to associate with this complex. The mitochondrial degradosome The mitochondrial degradosome consists of the 3'->5' hydrolytic exonuclease Dsslp and the DExH-box (related to DEAD-box) helicase Suv3p (Min et al., 1993; Dmochowska et al., 1995; Margossian et al., 1996; 51 Dziembowski et al., 1998; Dziembowski & Stepien, 2001; Dziembowski et al., 2003). It is involved in mitochondrial mRNA degradation and some mitochondrial rRNA processing, and is essential for mitochondrial biogenesis (Dziembowski et al., 1998). Surprisingly, mutation in either of the proteins of this complex results in defects in 3'- but also 5'-end processing (Dziembowski et al., 2003). Perhaps this is due to the genetic interaction of dss1 and suv3 with pet127, which encodes a membrane-bound enzyme involved in the 5'-end processing of mitochondrial RNA (Wiesenberger & Fox, 1997; Wegierski et al., 1998). Defects in Suv3p can also be suppressed by overexpression of Mss116p, an RNA helicase (not of any DEAD-box-related family) (Minczuk etal., 2002). This is the first reported substitution of a helicase by one from another class. Proteins of both ribosomal subunits co-purify with this complex, suggesting that degradosome particles associate with ribosomes (Dziembowski etal., 2003). The chloroplast degradosome mRNA decay in chloroplast resembles the E. coli process, with the exception that mRNAs are stable for hours, not minutes (Barkan & Stern, 1998; Hayes etal., 1999; Schuster etal., 1999; Baginsky etal., 2001). The search for a complex of enzymes involved in mRNA decay yielded the RNP100 complex, containing PNPase, a polyadenylation activity, and p67 (a protein recognized by E. coli RNase E antibodies) (Hayes et al., 1996). Upon closer inspection, the poly(A) activity was attributed to PNPase (Li (Q.S.) etal., 1998; Yehudai-Resheff etal., 2001), and p67 was identified as GroEL, which is known to cross-react with 52 RNase E antibodies (Hajeer & Bernstein, 1993; Sohlberg et al., 1993; Alconada et al., 1994; Baginsky et al., 2001). thus, the RNP100 complex contains only PNPase. 1.7 Adaptation of mRNA decay Since mRNA decay is an important mechanism for controlling gene expression, and given the focal role of the degradosome in this process, this complex could participate in both global and specific responses to stresses such as heat, cold, starvation, etc... 1.7.1 Global responses A few conditions are known to cause global changes in RNA decay. For example, when E. coli are starved for essential nutrients, rRNAs are destabilized (Ben-Hamida & Schlessinger, 1966; Kaplan & Apirion, 1975; Davis et al., 1986). Meanwhile, anaerobiosis and a translational block both cause bulk mRNAs to be stabilized (Lindahl & Forchhammer, 1969; Pato et al., 1973; Georgellis et al., 1993; Sousa et al., 2001). The degradosome is a good target for effecting global changes to mRNA decay. There are three mechanisms for achieving such regulation: limiting the quantity of degradosomes available, modulating the activity of its enzyme constituents, or altering the composition of the degradosome. 53 Limiting the amount of RNase E is the mechanism used to stabilize mRNA following a translational block (Sousa et al., 2001). It is also used to stabilize mRNAs during a transient diauxic lag approximately midway through the exponential phase (Barlow et al., 1998), and during the 15 to 30 minutes period following a temperature upshift to 44°C in minimal media (Le Derout etal., 2002). There are several mechanisms that can be used to control the concentration of RNase E. One is to increase the amount of RNA substrate while maintaining the concentration of RNase E constant, thereby titrating the enzyme (Sousa et al., 2001). Another is to increase polyadenylation. Polyadenylation stabilizes the RNase E and PNPase transcripts, leading to increased enzyme concentrations (Mohanty & Kushner, 1999, 2000a, 2002). Another possibility is that a protease, such as the membrane-bound protease MrsC/HfIB which is believed to act on RNases, digests RNase E and PNPase under certain conditions to regulate their concentrations (Granger et al., 1998; Wang etal., 1998). The activity of degradosomal proteins can be modulated using a variety of mechanisms. The first is through phosphorylation. Gene 0.7 of bacteriophage T7 encodes a serine/threonine kinase capable of phosphorylating both RNase E and RhIB (Marchand et al., 2001). Phosphorylation of RNase E stabilizes transcripts synthesized by T7 RNA polymerase, without affecting the degradation of other RNAs. It is believed that phosphorylation impedes 5'-end-independent entry, which is the mechanism used by RNase E to degrade T7 transcripts. Allosteric control of the activity of degradosomal proteins could also be used. For 54 instance, in chloroplasts, light induces a protein to bind and inhibit the rate-limiting endonuclease, stabilizing all the mRNAs encoding photosynthetic proteins (Baginsky & Gruissem, 2002). It has been noted that the Streptomyces antibioticus PNPase is able to bind to both mRNA and pppGpp (Jones & Bibb, 1996). Since pppGpp is a signal for differentiation (Hoyt & Jones, 1999), its binding to PNPase could be involved in regulating mRNA decay during the lifecycle of this organism (Symmons etal., 2002). Although there are currently no known examples of proteins that associate/dissociate from the degradosome in response to environmental changes, the variability of the composition of this complex suggests that such a mechanism could exist. 1.7.2 Specific Responses There is already a long list of factors that affect the decay of specific mRNAs. In aerobic cultures, growth rates do not seem to affect the stability of most transcripts (Pato & von-Meyenbrug, 1970; Coffman era/., 1971; Gausing, 1974, 1977; Maaloe, 1979; Bremer & Dennis, 1996; Meyer & Schottel, 1991; Bernstein et al., 2002), yet the stability of some mRNAs are clearly affected (Nierlich, 1968, 1972, 1978; Nilsson etal., 1984; Lundberg etal., 1988; Melefors & von Gabain, 1988; Melin etal., 1990; Vytvytska etal., 1998; Emory & Belasco, 1990; Georgellis et al., 1992; Bernstein et al., 2002). During anaerobiosis, the stability of the bla and ompA mRNAs increases (Georgellis et al., 1993). In 55 Rhodobacter, the half-life of the pufBALMX mRNA, encoding components of the photosynthetic apparatus varies with oxygen tension (Klug, 1991). In Agrobacter, the groESL operon is cleaved endonucleolytically following heat shock to release monocistronic groEL mRNA (Segal & Ron, 1995). While heat shock does not appear to influence bulk mRNA stability in E. coli (Henry et al., 1992), it influences the lifespan of the mRNA encoding ATP synthase (Lagoni et al., 1993). To affect the degradation of all mRNAs, the RNA degradation machinery is altered (Section 1.7.2). To cause changes in the stability of a specific transcript, the recognition of this RNA by RNA decay enzymes is modified. This is usually achieved by binding of a protein to the RNA. For example, growth rate-dependent binding of the Hfq protein to the 5'-end of the ompA mRNA stimulates cleavage by RNase E (Vytvytska et al., 1998, 2000). Conversely, binding of GroEL to specific mRNA targets during nutrient limitation has been proposed to protect these RNAs against the endonucleolytic action of RNase E (Georgellis et al., 1995). Post-transcriptional regulation of bacterial carbohydrate metabolism is attributed to the protein CsrA, which binds to and destabilizes glgCAP and other mRNAs to repress their expression (Romeo, 1996, 1998). Proteins are not the only molecules whose binding changes the susceptibility of certain mRNAs to the degradosome: small RNAs can also perform this function. The small RNA DsrA can bind to both the start and stop codon of hns mRNA to decrease the mRNA half-life (Lease & Belfort, 2000). Binding of this RNA to rpoS mRNA during cold 56 shock has the opposite effect (Lease & Belfort, 2000). Thus, based on the information available to date, the factors that cause changes in the stability of specific RNAs target the RNA, not the degradosome. In support of this assertion, changes in growth rate, which affect the stability of only some transcripts, do not influence the cellular amounts of RNase E, PNPase, and RhIB (Vytvytska e t al., 1998). 2 GOALS The goal of this dissertation is to study how the activity of the degradosome is altered in response to changes in the environment. Two different mechanisms for conveying these changes to the degradation machinery are explored. The first is the possibility that enolase modulates the activity of RNase E, PNPase, and RhIB in response to changes in the cell's energetic environment. A possible mechanism for doing this is through changes in its conformation after binding to 2-phosphoglycerate or phosphoenolpyruvate. If the other degradosome proteins are sensitive to the structural configuration of enolase, then the availability of the two glycolytic intermediates could have an effect on the activity of the degradosome. Enolase phosphorylation is another possible mode of action. This chemical modification occurs in response to growth on certain carbon sources and is known to alter the glycolytic activity of enolase. 57 This alteration could potentially signal other degradosomal proteins of the shift in conditions. The second area explored is the effect of cold shock on the composition of the degradosome. Specifically, given that RNA secondary structures are stabilized by a drop in temperature, can the degradosome adapt by recruiting a second, possibly "stronger" helicase to help PNPase degrade stable 3' hairpins. A clear candidate helicase is CsdA, which is expressed under cold shock conditions, and has already been implicated in mRNA decay at 15°C. The variability of degradosome composition and size hints at the possibility that different forms of the degradosome could exist. If CsdA can be shown to combine with the degradosome under cold shock conditions, it will represent the first example of regulation of degradosome activity at the level of composition. 58 CHAPTER II MATERIALS AND METHODS 2.1 INTRODUCTION The majority of the work presented in this thesis was done using in vitro laboratory techniques. Most protocols are modifications of already established methods. 2.2 SOURCE of ENZYMES and CHEMICALS When available, enzymes were obtained from commercial sources such as Amersham Biosciences, Gibco-BRL, MBI Fermentas, New England Biolabs, and Promega. Most chemicals were purchased from Fisher, Sigma, and VWR (BDH). Chromatographic materials were obtained from Amersham Biosciences, and Bio-Rad. 2.3 BACTERIAL STRAINS AND PLASMIDS E. coli strain BL21 (DE3), obtained from Novagen, is the parent of all overexpressing strains used in this work. CF881 was a gift from Dr. M. Cashel (National Institutes of Health, Bethesda). This strain harbours a deficient copy of ribonuclease I, a major contaminant in ribonuclease purification. P90C is the wild type strain used by the laboratory of Dr. R.W. Simons (UCLA), a collaborator for the cold shock work. Many plasmids were derived from the pET series of bacterial vectors (Novagen). 59 2.3.1 Overexpressing strains E. coli cells capable of overexpressing RNase E, FLAG-RNase E, His-RNase E, PNPase, His-PNPase, RhIB and enolase were obtained from the appropriate sources (Tables 2.1 and 2.2). A His-CsdA overexpressing strain was created for this work, in collaboration with Rudolf K. Beran at the University of California in Los Angeles. Wild-type Rne Strain GM402 (BL21 (DE3)/ pGM102; Cormack et al., 1993) is capable of overexpressing wild-type Rne protein upon induction. pGM102 encodes the complete rne coding sequence under the control of the T7 lac promoter-operator of pET-11. FLAG-Rne cells Plasmids pflagLRC and pRE196, encoding the inducible FLAG epitope (N-Asp-Tyr-Lys-Asp-Asp-Asp-Asp-Lys-C) or FLAG-Rne constructs, respectively, were obtained from Dr. Sue Lin-Chao (Miczak et al., 1996) and transformed into BL21 (DE3) (Section 2.4.5). His-Rne cells Plasmid pRne16-15, a derivative of pET-16b containing the complete Rne coding sequence between the Pstl and Ndel restriction sites, was transformed into BL21 (DE3). This places the Rne coding sequence behind a histidine tag (10 sequential histidine residues), under the control of the T7/ac 60 Table 2.1 List of bacterial strains Strain Genotype Description Source/References BASIC STRAINS BL21 (DE3) E. coli B F" ompJ hsdSB (rB" mB") gal dcm MDE3) Encodes the T7 RNA polymerase under lac promoter, protease deficient Novagen, Studier & Moffatt, 1986; Studier era/., 1990 CF881 F" A/ac argA trp recB1009A (xthA-pnc) Arna RNase I deficient Dr. M. Cashel (National Institutes of Health, Bethesda) DH5cc™ F-, <j)80d/acZAM15 A(/acZYA-argfF)U169 deoR recM endA1 hsdRM (rk\ mk+) phoA supE44 r thiA gyrA96 re/A1 Mutations increase suitability for plasmid cloning procedure (more stable and greater quantity) Invitrogen Life Technologies, Inc. P90C A(lac-pro) thi ara Miller, 1992 OVEREXPRESSING STRAINS ^ GM402 BL21 (DE3) - pGM102 Rne overexpression Cormack etal., 1993 BL21 (DE3) -pflagLRC BL21 (DE3) - pflagLRC FLAG epitope construct Miczak etal., 1996 BL21 (DE3) -PRE196 BL21 (DE3)-pRE196 FLAG-Rne overexpression Miczak etal., 1996 RNE16-15 BL21 (DE3)-pRNE16-15 His-Rne overexpression J.L. Genereaux and G.A. Mackie, (U.B.C., unpublished) GC400 BL21 (DE3) - pGC400 Pnp overexpression Coburn & Mackie, 1998 BL21 (DE3) -pEPct18 BL21 (DE3) -pEPcx18 His-Pnp over-expression Py etal., 1994 GC300 BL21 (DE3) - pGC300 RhIB overexpression Coburn etal., 1999 BL21 (DE3) -pET-eno BL21 (DE3) - pET-eno Enolase over-expression Dr. A.J. Carpousis, (Laboratoire de microbiologie et genetique moleculaire Toulouse) BL21 (DE3) -pRS3486 BL21 (DE3) - pRS3486 His-CsdA over-expression Rudolf K. Beran, UCLA MUTANT STRAINS ENS134 pnp::Tn5 ENS134 (BL2KDE3), P,ac-T7 RNA polymerase, PJ7-lacZ), pnp::Tn5 pnp deletion strain Marc Dreyfus (Laboratoire de genetique moleculaire, Paris) RS9052 P90C, csdA::Km R csdA deletion strain Rudolf K. Beran, UCLA RS9215 BL21 (DE3), pnp:: Tn5 pnp deletion strain Rudolf K. Beran, UCLA RS9216 BL21 (DE3), csdA::Kan csdA deletion strain Rudolf K. Beran, UCLA 61 Table 2.2 List of plasmids Plasmid Comment Reference/Source PLASMIDS FOR RNA SYNTHESIS pCH77 Contains the intergenic spacer McLaren et al., 1991 region of the MalE-MalF mRNA under the control of the T7 promoter - derivative of pGEM™-2 pGM79 rpsT mRNA behind SP6 promoter Mackie, 1991 PLASMIDS FOR PROTEIN OVEREXPRESSION pGM102 Derivative of pET-11 containing the complete Rne gene, behind a P-T7 O-lac promoter Cormack etal., 1993 pflagLRC Ap H Miczak etal., 1996 PRE196 FLAG-me + Miczak etal., 1996 pRNE16-15 pET-16b containing the complete Rne gene (between the Pstl and Ndel restriction sites) behind a His tag J.L. Genereaux & G.A. Mackie (University of British Columbia, unpublished) pGC400 pET-11 containing the complete pnp gene Coburn & Mackie, 1998 pEPa18 His-pnp+, based upon pET-14b Py etal., 1994 PGC300 pET-11 with rhIB sequence Coburn etal., 1999 pET-eno pET-11a with complete enolase A.J. Carpousis (Laboratoire sequence between Ndel and de microbiologie et BamHI sites genetique moleculaire Toulouse) pRS3486 pET-15b containing the csdA Rudolf K. Beran, U.C.L.A. sequence between the Nde I and Hind III restriction sites 62 promoter. This construct was obtained from J.L. Genereaux and G.A. Mackie (University of British Columbia, unpublished). Wild-type Pnp cells GC400 containing the plasmid pGC400 in BL21 (DE3) (Coburn & Mackie, 1998). This plasmid is a derivative of pET-11, and encodes the entire Pnp sequence under the control of the T7 lac promoter-operator. His-Pnp cells Plasmid pEPal 8 (Py et al., 1994), a pET-14b-derived plasmid, was transformed into BL2KDE3) for overexpression of N-terminally His-tagged PNPase. Wild-type RhIB cells Strain GC300, harbouring a pET-11 plasmid with the complete RhIB sequence (Coburn et al., 1999) was used to overexpress RhIB. Wild-type enolase cells Plasmid pET-eno, a derivative of pET-11a encoding the entire enolase coding sequence, was transformed into BL21 (DE3) to create a strain capable of overexpressing enolase following induction by IPTG. This strain was a gift from Dr. A. J . Carpousis (Laboratoire de microbiologie et genetique moleculaire, Toulouse, France; Py etal., 1996). 63 His-CsdA cells Plasmid pECSRA201, encoding the csdA sequence behind the T7 promoter of pET-11a, was obtained from Dr. Pamela Jones (University of Georgia, Athens). The csdA gene was removed from this plasmid by digestion with Nde I and Hind III. The 2,300 bp fragment obtained by gel extraction was ligated into Nde I and Hind Ill-digested pET-15b using T4 ligase to generate pRS3486, which was transformed into BL21 (DE3). 2.3.2 pnp and csdA mutant strains ENS134 pnp::Tn5 cells Strain ENS134 (Lopez et al., 1994) containing an interrupted (null) pnp gene was obtained from Dr. Marc Dreyfus, Laboratoire de Genetique Moleculaire (CNRS UMR 8541), Paris, France. In addition, pnp and csdA knock-out strains were created for this work, in collaboration with Rudolf K. Beran (UCLA), as follows: RS9052 cells Strain P90C containing csdA::Km R was obtained from Rudolf K. Beran, UCLA. BL21 pnp::Tn5 cells Strain JC357 containing pnp::Tn5 was obtained from Dr. Claude Portier (Py et al., 1994). P1 phage was used to transduce pnp::Tn5 into BL21 (DE3) by selecting for kanamycin resistance. The absence of Pnp protein in this new strain, RS9215, was verified by Western blotting, and by growth deficiency at 15°C. 64 BL21 c s d A : : T n 5 cells Strain JC7623 csdA::Kan, containing an insertion of Tn5 into csdA, was obtained from Dr. Pamela Jones (Jones et al., 1996). P1 phage was used to transduce this mutation into BL21 (DE3), thereby creating strain RS9216. Disruption of the csdA gene was confirmed by DNA sequencing of the amplified csdA::Kan construct, and by a lack of signal on an anti-CsdA Western blot. 2.4 BACTERIAL CULTURES All bacterial culture work was done using sterile techniques. Unless otherwise noted, all strains were grown using the following standard methods. 2.4.1 Growth media Cell growth was carried out in either Luria-Bertani (LB) broth or agar, Terrific broth, or M9 minimal media, as noted. These were prepared as described by Sambrook et al. (1989): LB broth (1% (w/v) granulated peptone from casein (pancreatically digested; EM Biosciences), 0.5% (w/v) yeast extract (EM Biosciences), and 86 mM NaCI); L B a g a r (LB broth with 20 g/L of agar); Terrific broth (1.3% (w/v) granulated peptone from casein (pancreatically digested), 2.7% (w/v) yeast extract, 0.4% (v/v) glycerol, 0.017 M K H 2 P 0 4 , and 0.072 M K 2 HP0 4 ) ; M9 minimal media (50 mM Na 2 HP0 4 , 22 mM K H 2 P 0 4 , 2 mM NaCI, 19 mM NH4CI, supplemented after sterilization with 0.1 mM CaCl2, 1 mM 65 MgS0 4 , 0.001 % (w/v) vitamin B1, 0.2% (w/v) glucose, 0.004% (w/v) amino acids (all except methionine and cysteine)). 2.4.2 Rescue of strains from frozen stocks To prevent growth of contaminated cultures, a loopful of a -70°C stock was streaked onto LB agar plates containing appropriate antibiotics (in most cases 75 u.g/ml ampicillin) and grown overnight at 37°C. Cells from one colony were picked and grown with vigorous shaking at 37°C in 15 ml of LB broth containing a suitable antibiotic (in most cases 100 u.g/ml ampicillin) LB broth for 4 hours. This "starter culture" was used to inoculate 150 ml of appropriate media and the cell grown at 30°C in a water bath shaker at 225 rpm. 2.4.3 Induction of cultures BL21 (DE3) cells bearing the appropriate plasmid were grown with vigorous aeration in 150 ml of LB supplemented with selective antibiotic at 30°C to an OD6oo of 0.4. To induce the expression of protein, IPTG was added to a final concentration of 1 mM, followed by addition of 150 ml of fresh LB broth containing antibiotics. Expression of protein was allowed to proceed for an additional 4 hours at the same temperature. Overexpression was confirmed by sampling 1 ml of culture before and after induction, pelleting the cells in a microfuge at top speed for 5 minutes, resuspending the cells in 150 uJ 2X sample buffer (Section 2.6.4), and boiling 5 66 minutes. The overexpressed protein band was easily detectable on an SDS-PAGE gel (Section 2.6.4). 2.4.4 Recovery of cells by centrifugation Following growth or overexpression, cells were harvested by centrifugation using a Beckman JLA 10.5 rotor at 5,000 X g for 10 minutes. The pelleted cells were washed twice in buffered solutions (various, Section 2.5), and again recovered by centrifugation. 2.4.5 Bacterial transformation Competent cells Cells (250 ml) were grown in LB broth to an OD6oo of 0.9. The bacteria were harvested by centrifugation, resuspended in 50 ml of ice-cold 100 mM MgCI2, and gently mixed on ice. The cells were again recovered by centrifugation, re-suspended in 110 ml of 100 mM CaCI 2, and left on ice for 1 hr. Following this treatment, the cells were pelleted, and re-suspended in 12.5 ml of 85 mM CaCI 2 and 15% (v/v) glycerol. These competent cells were aliquoted in 0.5 ml portions, quickly frozen on an ethanol/dry-ice bath, and stored at -70°C until use. Transformation One aliquot of competent cells was thawed on ice. Isolated DNA (1-10 ng) was gently mixed with the cells, which were left on ice for 60 minutes. The cells were heat-shocked for 3 minutes at 42°C, and then cooled on ice for 5 minutes. LB broth (1 ml) was added to cells, which were incubated at 67 37°C for 1 hr. The cells were then plated on LB agar plates containing appropriate antibiotics, and grown overnight at 37°C. A few colonies were selected, streaked for single colonies on fresh LB-antibiotics plates, and grown overnight at 37°C. 2.5 ENZYMES and ANTIBODIES 2.5.1 Enzyme purification The degradosome, all individual degradosomal proteins, as well as CsdA were purified from E. coli cells. Overexpressing strains were used to facilitate and optimize purification, except for degradosomes. Established purification protocols were followed with only minor modifications in the case of RNase E, FLAG-RNase E, PNPase, and degradosomes. Novel ways of purifying His-PNPase, RhIB, and CsdA were devised for this work. SDS-PAGE gels illustrating the purification achieved through these protocols are shown in Figure 2.1. These proteins were used in all subsequent experiments. Table 2.3 describes some physical properties of these proteins. Degradosome purification Degradosomes were purified from CF881 cells as previously described (Carpousis ef al., 1994; Py et al., 1996; Coburn & Mackie 1996a; Coburn & Mackie, 1998; Carpousis et al., 2001), with some modifications. CF881 cells were grown in 500 mL of Terrific broth in 2 L flasks. Growth was carried out overnight at 37°C, or over a 3 day period at 15°C. Cells were harvested by 68 Figure 2.1 Purified Proteins A. 0.75 u.g purified RNase E, PNPase, RhIB and enolase were separated electrophoretically on a 7.5% SDS-PAGE gel and stained with Coomassie Blue dyes to show purity. B. 3.6 u.g degradosomes purified from cells grown at 37°C. C. Purified tagged proteins: 27.5 \ig total protein from the AS26 extract of cells overexpressing FLAG-RNase E, 1 u.g His-Pnp, 1 u.g His-CsdA. 69 Table 2.3 Physical properties of purified proteins Protein MW (kDa) amino acids oligomerization Pi RNase E 118 1061 tetramer 5.5 PNPase 78 711 trimer 5.1 RhIB 48 421 dimer / monomer 7.3 enolase 46 432 dimer 5.3 CsdA 73 646 9.0 70 centrifugation, and washed twice in cell wash buffer (10 mM Tris-HCI (pH 7.5), 10 mM magnesium-acetate, 10 mM KCI) (Misra & Apirion, 1979). Typically, 18 g of cells were recovered at the higher temperature, and 4 g at the lower one. Cells were lyzed by resuspending them in 15 mL lysozyme-EDTA buffer (50 mM Tris-HCI (pH 7.5), 100 mM NaCI, 5% (v/v) glycerol, 3 mM EDTA, 1 mM DTT, 1.5 mg/ml lysozyme, 1 mM PMSF, 2 u.g/ml aprotinin, 0.8 u.g/ml leupeptin, 0.8 u.g/ml pepstatin A) at room temperature, and subjecting them to the following treatment: stirring for 1 min, incubation on ice for 10 min, stirring 1 min, incubation on ice 20 min, stirring 1 min, incubation on ice 40 min. Subsequently, 7.5 ml of DNase-RNase-Triton buffer (50 mM Tris-HCI (pH 7.5), 100 mM NaCI, 5% (v/v) glycerol, 1 mM DTT, 30 mM magnesium acetate, 3% (v/v) Triton X-100, 20 u:g/ml DNase I, 1 mM PMSF, 2 u.g/ml aprotinin, 0.8 u.g/ml leupeptin, 0.8 u.g/ml pepstatin A, 25 u.g/ml RNase A) at room temperature was added to the suspension, followed by 1 min of stirring, and a 30 min incubation on ice. Ammonium chloride was slowly added to a final concentration of 1M, and was stirred in for 30 min on ice. The lysate was subjected to 30,000 X g centrifugation in a Beckman JA-20 rotor for 60 minutes to remove unbroken cells and debris. The supernatant was then subjected to centrifugation for 2 hrs at 200,000 X g in a Beckman Ti 75 rotor. The supernatant was collected and fractionated by the slow addition of 26% (w/v) ammonium sulfate on ice, with constant stirring over a 30 min period. 71 The precipitated proteins were recovered by centrifugation at 10,000 X g for 60 min in a JA-20 Beckman rotor. Due to the presence of detergents, some of the pellet floated to the surface and was collected as well. The pellet and "floating scum" were re-suspended in 40 ml buffer D (10 mM Tris-HCI (pH 7.5), 5% (v/v) glycerol, 0.5% (v/v) Genapol X-080, 1 mM EDTA, 0.1 mM DTT, 0.1 mM PMSF, 2 pg/ml aprotinin, 0.8 u.g/ml leupeptin, 0.8 u.g/ml pepstatin A) and 50 mM NaCI. This suspension was applied to an SP Sepharose Fast Flow (Amersham Pharmacia) cation exchange column (2.5 cm X 10.5 cm) previously equilibrated with buffer D and 50 mM NaCI. The column was washed with three column volumes of buffer D + 50 mM NaCI, at a flow rate of 1 ml/min. The bound proteins were eluted in buffer D containing 1% (v/v) Genapol X-080 with a gradient from 0.1 M NaCI to 1 M NaCI. The eluate was collected in 2 ml fractions, at a flow rate of 1 ml/min, over 50 fractions. 100 \x\ from each fraction was precipitated with acetone prior to loading on an SDS-PAGE gel (Sections 2.6.2 and 2.6.4). Coomassie Blue staining of the gel revealed that the proteins eluted in fractions 39 to 47 for the cells grown at 37°C, and 26 to 49 for the cells grown at 15°C. These fractions were pooled and concentrated using ammonium sulfate (Section 2.6.1). The precipitated proteins were resuspended in buffer D, 50 mM NaCI, and 15% (v/v) glycerol. This fraction was loaded on a Bio-Rad Biogel A 5M size exclusion column (1 cm X 50 cm) pre-equilibrated with buffer D and 50 mM NaCI. The proteins 72 were separated on the column using a flow rate of 0.5 ml/min. Fractions of 4 ml were collected. Degradosomes isolated from 37°C cells eluted from the column in fractions 10 to 14, and those from 15°C in fractions 8 to 11. The pooled fractions were concentrated using Millipore's Amicon Ultra centrifugal filter devices (MWCO: 10,000) to roughly 1 ml. The 37°C preparation yielded a 0.4 mg/ml sample, and the 15°C cells a 0.8 mg/ml degradosome preparation (as determined by the Bradford Assay, Section 2.6.3). Rne purification Rne purification Rne protein was purified as previously described (Cormack et al., 1993; Coburn et al., 1999; Mackie et al., 2001). Cultures of GM402 cells were grown at 30°C and induced with 1 mM IPTG for 3 hours. They were harvested by centrifugation, and washed twice with cell wash buffer (Section supplemented with 6 mM |3-mercaptoethanol. The cells were re-suspended in 4 ml/g in buffer RNE-A (50 mM Tris-HCI (pH 7.6), 10 mM MgCI2, 60 mM NH4CI, 0.5 mM EDTA), supplemented with 7.5% (v/v) glycerol, 0.1 mM DTT, and 0.2 mM PMSF. Bacteria were lyzed by two passages through a French pressure cell (Aminco) at 8,000 psi, after which additional DTT and PMSF (to twice their original concentrations) were added, as well as 2.3 u.g/ml of DNase I, and left on ice for 10 min. An S30 supernatant was obtained by centrifugation at 30,000 X g for 45 min in a JA-20 Beckman rotor. This 13 ml fraction was diluted to 40 ml with buffer RNE-A containing 7.5% (v/v) glycerol, 0.1 mM DTT, and 0.2 mM PMSF. The diluted extract was made to 26% (w/v) ammonium 73 sulfate. The salt was slowly stirred in over a 30 minute period in the cold room. The precipitated proteins were collected by centrifugation at 17,000 X g for 20 minutes in a Beckman JA-20 rotor. The pellet was re-suspended in cell wash buffer and 0.1 mM DTT, 7.5% (v/v) glycerol, 0.2 mM PMSF, and 0.2 ug/ml leupeptin. Rne was separated from the other proteins of the AS26 extract by electrophoresis. Using a preparative gel apparatus and 3 mm spacers, the extract was loaded in 2X sample buffer (Section 2.6.4) on a 5.5% (49:1) SDS-PAGE g e l (separating gel: 5.5% (g/ml) polyacrylamide (49:1 acrylamide: N, N'-methylenebis-acrylamide), 0.375 M Tris-HCI (pH 8.8), 5% (v/v) glycerol, 0.1% (v/v) SDS, 0.1% (v/v) TEMED and 0.04% (g/ml) ammonium persulfate; stacking gel: 4.5% (g/ml) polyacrylamide (49:1 acrylamide: N, N'-methylenebis-acrylamide), 0.13 M Tris-HCI (pH 6.8), 0.1% (v/v) SDS, polymerized by the addition of 0.1% (v/v) TEMED and 0.04% (g/ml) ammonium persulfate). The left and right side of the gel were cut off and stained using Coomassie Blue. The location of the Rne band on these gel slices was used to estimate the location of the Rne band in the unstained portion of the gel (the staining process causes swelling, which was taken into account in the calculations). The estimated location of the Rne band was cut out using a clean razor blade, cut into smaller pieces, and placed in a dialysis tubing with a minimal volume of Laemmli buffer (Section 2.6.4). The protein was electroeluted from the gel by placing the dialysis tubing in a 100 volt field in Laemmli buffer (Section 2.6.4) for 6 hrs, the 74 current running perpendicularly to. the length of the tubing. The liquid was extracted, and the proteins precipitated with 5X volume of acetone (Section 2.6.2). The precipitated proteins were dissolved with 200 u.1 of solubilization buffer (6 M guanidine-HCI, 50 mM Hepes-NaOH (pH 7.6), 100 mM NaCI, 1 mM EDTA, 1 mM DTT, and 0.05% (v/v) Tween-20), for 45 min at 30°C. The solution was then centrifuged in a microfuge at maximal speed for 10 min at 4°C to remove insoluble material. The concentration of guanine was reduced to 1.5 M by the addition of renaturation buffer (50 mM Hepes-NaOH (pH 7.6), 100 mM NaCI, 1 mM EDTA, 1 mM DTT, 0.05% (v/v) Tween-20, and 5% (v/v) glycerol). Following a 35 minute incubation at room temperature, the protein preparation was dialyzed at 4°C in 500 ml of the following solution: 25 mM Hepes-NaOH (pH 7.6), 100 mM NaCI, 0.1 mM EDTA, 0.1 mM DTT, and 5% (v/v) glycerol. After an initial 5 hour dialysis, the buffer was refreshed and continued overnight. Renatured Rne was then concentrated using Millipore UltraFree centrifugal filter devices (BioMax 5K NMWL membranes), and the protein concentration determined using the Bradford assay (Section 2.6.3). Aliquots were stored at -70°C until use. Partially Purified FLAG-Rne Cultures of BL21 (DE3) containing pRE196 were grown in ampicillin-containing LB broth and induced by the addition of 0.5 mM IPTG for 4 hours. Cells were collected by centrifugation, re-suspended in 75 buffer RNE-A supplemented with 7.5% (v/v) glycerol, 100 uM DTT, 2 ug/ml aprotinin, 0.8 u.g/ml leupeptin, and 0.8 u.g/ml pepstatin A, and lyzed by two passages through a French pressure cell (8,000 psi). The lysate was treated with 2.3 u.g/ml DNase I and 1 mM PMSF for 10 min on ice, and centrifuged at 30,000 X g for 45 min at 4°C. The supernatant was made to 26% (w/v) with ammonium sulphate, and the precipitated FLAG-Rne recovered by centrifugation at 17,000 X g for 20 min at 4°C. The pellet was resuspended in buffer RNE-B (50 mM Tris-HCI (pH 7.5), 10 mM MgCI2, 1.5 mM NaCI, 2.5 mM EDTA, 1 mM DTT, 7.5% (v/v) glycerol, 0.2 mM PMSF, 2 u.g/ml aprotinin, 0.8 u.g/ml leupeptin, and 0.8 u.g/ml pepstatin A). The resuspended protein fraction is referred to as an "AS26". The protein concentration was determined using the Bradford Assay (Section 2.6.3). Pnp purification Pnp purification. Pnp protein was purified by chromatographic techniques as previously described, with some modifications (Coburn & Mackie, 1998). GC400 was grown in 150 ml of LB broth containing ampicillin at 30°C, and induced for 4 hours by the addition of 1 mM IPTG. Cells were collected by centrifugation at 5,000 X g for 15 minutes in a JLA 10.5 Beckman rotor, and washed twice in buffer PNP-A (50 mM Hepes-NaOH (pH 7.5), 0.2 mM EDTA, 2 mM DTT, 0.1 mM PMSF, and 5% (v/v) glycerol). The bacteria were lysed by two passages through a French pressure cell at 8,000 psi. DNase I (2.3 u.g/ml), and the protease inhibitors aprotinin (2 u.g/ml), pepstatin A (0.8 u.g/ml), and leupeptin 76 (0.8 u.g/ml) were added to the lysate and left at room temperature for 15 minutes. Unbroken cells and debris were removed by centrifugation at 30,000 X g for 1 hour in a Beckman JA-20 rotor. The extract was loaded onto a heparin-agarose column (26 mm X 10 cm; Bio-Rad), at a flow rate of 2 ml/min in buffer PNP-B (25 mM Tris-HCI (pH 7.5), 5% (v/v) glycerol, 1 mM DTT, 0.1 mM EDTA) using an FPLC. The column was washed with this buffer at this rate, and the bound proteins eluted with a salt gradient from 0 to 1 M NaCI. Fractions of 2 ml were collected. As assessed by SDS-PAGE, PNPase eluted in fractions 12 to 26, with a substantial amount in the flowthrough. This portion was reloaded on the column, and the bound proteins combined with those of the first load. The original purification protocol called for passage through an Affi-Blue column. This was not performed in the current studies, as previous experience suggested this step is of only limited usefulness in separating PNPase from the remaining contaminants. To prepare the extract for the next step, the pooled heparin fractions were concentrated using ammonium sulfate (Section 2.6.1). The precipitated proteins were re-suspended in buffer PNP-B supplemented with 100 mM NaCI, loaded on a Biogel A 0.5M (100-200 mesh, MWCO: 500,000; 1 cm X 48 cm; Bio-Rad) size exclusion column and subjected to chromatography in the same buffer. Four milliliter (4 ml) fractions were collected. PNPase eluted in 77 fractions 11 to 22. These were concentrated using Millipore centrifugal filter devices (MWCO: 10K). As an undesirable amount of contaminants was still present at this point, an additional step was added. The concentrated PNPase solution was passed through a Resource Q column (FPLC) at 0.75 ml/min, washed in buffer PNP-B at 1 ml/min, and the bound proteins eluted with a NaCI gradient from 0 to 1 M over 100 ml. Fractions of 2 ml were collected, and PNPase eluted from the column roughly midway through the gradient, in fractions 21 through 25. The majority of contaminants eluted slightly ahead of PNPase. Fractions of interest were concentrated using Millipore centrifugal filter devices, producing 700 ul of a 3 mg/ml PNPase solution, as determined by the Bradford assay (Section 2.6.3). His-Pnp purification E. coli BL2KDE3) harbouring pEPa18 were grown with shaking in 300 ml LB broth with 50 ug/ml ampicillin at 30°C. After induction with 1 mM IPTG for 2 hrs, cells were collected by centrifugation, resuspended in 5 ml cell wash buffer supplemented with 1 mM PMSF, and lyzed using a French pressure cell (10,000 psi). The lysate was treated with 2.3 u.g/ml DNase I on ice for 10 minutes, and the extract centrifuged at 30,000 X g for 45 min at 4°C. The supernatant was passed over a TALON Metal Affinity column (Clontech). The column was washed with buffer PNP-C (50 mM Tris-HCI (pH 8.0), 500 mM NaCI) containing 10 mM imidazole, and the bound proteins eluted with buffer PNP-C supplemented with 50 mM imidazole. The fractions containing Pnp were 78 collected, concentrated using Millipore centrifugal filters to 1.2 ml, and subjected to size exclusion chromatography on a Biogel A 0.5M column (1.5 cm X 35 cm, Bio-Rad) in buffer PNP-D (25mM Tris-HCI (pH 7.7), 100 mM NaCI, 5% (v/v) glycerol, 1 mM DTT, 0.1 mM EDTA). Fractions containing Pnp were pooled and concentrated using Millipore centrifugal filter devices. Enolase Purification Enolase purification Enolase was purified by the method described by Kuhnel & Luisi (2001), with some modifications. Enolase was overexpressed from BL21 (DE3) cells containing the pET-eno plasmid as described in Section 2.4.3., recovered by centrifugation, and re-suspended in buffer RNE-A containing 7.5% glycerol, and 0.1 mM DTT. Lysis was accomplished by two passages through a French pressure cell at 8000 psi, following which DTT was added to 0.2 mM, PMSF to 0.2 mM, DNase I to 2.3 u.g/ml, and the extract was left on ice for 10 min. Contaminating proteins were removed by adding ammonium sulfate to 65% saturation, and stirring for 30 min at 4°C. The preparation was centrifuged at 10,000 X g for 25 min in a JS-13.1 rotor, and the supernatant collected. It was subjected to another round of ammonium sulfate precipitation, this time to 95% saturation. Enolase was found in the pellet after centrifuging at 10,000 X g for 25 min in a JS-13.1 rotor, and was resuspended in 0.5 ml of buffer ENO-A (20 mM Tris-HCI (pH 7.6), 1.9 M (NH 4) 2S04, 5 mM MgCI2, and 2 mM DTT). This extract was loaded on a 20 cm X 1 cm phenyl Sepharose fast flow (Amersham Pharmacia) column at a flow rate of 0.4 ml/min in buffer ENO-79 A. Following a 40 ml wash in this buffer, the protein was eluted in 3 ml fractions with an eluting gradient from buffer ENO-A to buffer ENO-B (20 mM Tris-HCI (pH 7.6), 5 mM MgCI2, and 2 mM DTT). Enolase eluted toward the end of the gradient. The fractions containing enolase were pooled, and Millipore centrifugal filter devices were used to simultaneously concentrate the proteins and change the buffer to buffer ENO-C (20 mM Tris-HCI (pH 8.0), 300 mM NaCI, 20 mM MgCI2, and 2 mM DTT). The protein sample was applied to a Sephacryl S-300 size exclusion column, and passed through the column at a flow rate of 0.4 ml/min in buffer ENO-C. The fractions containing enolase were pooled and concentrated on a Millipore centrifugal filter device. This extract was loaded on a Resource Q (Amersham Pharmacia) column using an FPLC apparatus. The sample was loaded at a flow rate of 1 ml/min in buffer ENO-D (50 mM Tris-HCI (pH 7.5), 10 mM MgCI2, 20 mM KCI, and 20 mM DTT), and collected in the flowthrough. The flowthrough was concentrated using a Millipore centrifugal filter device, and stored in aliquots at - 70°C until use. CsdA Purification His-CsdA The purification scheme for CsdA was devised and carried out by our collaborators, Rudolf K. Beran and Dr. Robert W. Simons at the University of California in Los Angeles. E. coli BL21 (DE3) harbouring pRS3486 were grown at 37°C in 250 ml LB broth to mid-log and induced for 3 hours with 1 mM IPTG. Cells were recovered by centrifugation, resuspended in buffer CSDA-A (20 mM Tris-HCI (pH 8.0), 500 mM NaCI, 5 mM MgCI2) supplemented with 1.7 80 mg/ml lysozyme, 17 u.g/ml DNase I and 10 mM imidazole, and left on ice for 30 minutes. They were then lysed by sonication with eight 10 second pulses. The extract was centrifuged at 12,000 X g for 30 minutes at 4°C. The supernatant was passed twice through a N i + + NTA agarose column (Qiagen) pre-equilibrated with buffer CSDA-A + 10 mM imidazole. The column was washed with 2.5 column volumes of buffer CSDA-A + 20 mM imidazole, and the bound proteins eluted in 1 ml of buffer CSDA-A + 1 M imidazole. The eluate was concentrated using Millipore centrifugal filter device to a final volume of 500 u.l, and supplemented with an equal volume of glycerol. This protein preparation was stored at -70°C. 2.5.2 Antibody sources Anti-RhIB and anti-CsdA polyclonal antibodies were developed for this work, as described below. Polyclonal antibodies against Rne and Pnp were available from our laboratory. Anti-enolase were a gift from Dr. A.J. Carpousis (Laboratoire de microbiologie et genetique moleculaire, Toulouse, France). Anti-FLAG® M2-agarose affinity gel and goat anti-rabbit lgG(H+L) affinity purified antibodies labeled with peroxidase were obtained from Sigma and Kirkegaard & Perry Laboratories, MD, respectively. Table 2.4 is a list of all antibodies used in this work. Anti-RhIB rabbit polyclonal serum RhIB (229 ug) isolated from GC300 cells was further purified on a 7.5% (36:1) SDS-PAGE gel by cutting out 81 Table 2.4 List of antibodies primary antibodies Comment ANTIBODIES DEVELOPED FOR THIS WORK Rabbit anti-RhIB polyclonal, from the laboratory of Dr. G.A. Mackie Rabbit anti-CsdA polyclonal, from the laboratory of Dr. R.W. Simons ANTIBODIES AGAINST DEGRADOSOMAL PROTEINS Rabbit anti-Rne polyclonal, from the laboratory of Dr. G.A. Mackie (Niguma, 1997) Rabbit anti-Pnp polyclonal, from the laboratory of Dr. G.A. Mackie (Niguma, 1997) Rabbit anti-eno polyclonal, from the laboratory of Dr. A.J. Carpousis COMMERCIALLY AVAILABLE ANTIBODIES Goat anti-rabbit IgG affinity purified, horse radish peroxidase conjugate, Kirkegaard & Perry Laboratories, MD Mouse antiFLAG® M2 monoclonal, Sigma 82 Comassie-stained (Commassie dissolved in 2% methanol) acrylamide gel slices with a clean razor blade. These were lyophilized for 24 hours and ground with phosphate buffered saline (PBS)-1 (0.14 M NaCI, 2.7 mM KCI, 10 mM Na2HP0 4, 1.4 mM KH2PO4, pH adjusted to 7.2, filter-sterilized). This preparation was emulsified with an equal volume of Freund's incomplete adjuvant (Sigma), and intramuscularly injected into New Zealand white rabbits to raise antibodies. A test bleed was performed before the first immunization and after each four subsequent boosts. The rabbit was ex-sanguinated, the blood allowed to clot and the serum stored frozen. Working portions were stored at -20°C in 50% (v/v) glycerol. Specificity was determined by Western blotting on whole E. coli cell extract, purified degradosomes, and purified RhIB and CsdA. Anti-CsdA rabbit polyclonal serum Purified His-CsdA protein (1 mg) was sent to Washington Biotechnology, Inc. (Baltimore, MD) to generate anti-CsdA rabbit polyclonal serum. Six weeks after the initial injection, a second 1 mg boost was used to increase the specificity of the antibodies. A specific antibody was obtained by week 13. 2.6 PROTEIN ANALYSIS The following techniques were used to prepare or analyze the contents of various protein samples. 83 2.6.1 Ammonium sulfate concentration of proteins This protocol was used to recover proteins from dilute solutions (for example, pooled column fractions) during purification (Schreier et al., 1977). Typically, 500 ml of a 50% (w/v) ammonium sulfate solution is prepared and placed in a graduated cylinder. The protein sample is placed in dialysis tubing (Spectra/Por® Membrane MWCO: 3,500) and immersed in the ammonium sulfate solution overnight at 4°C. The precipitated proteins are collected by centrifugation at 12,000 X g in a Beckman JA-20 rotor for 15 minutes. 2.6.2 Acetone precipitation of proteins This procedure was used to load sufficient material on a gel from a dilute protein sample. Proteins are precipitated overnight at -70°C by the addition of 5 volumes of 100% (v/v) acetone (Barritault et al., 1976), and are recovered by centrifugation at maximal speed in a microfuge for 10 min at 4°C. In addition to proteins, acetone precipitates salts, which are washed out of the pellet with 80% (v/v) acetone. The samples are dried by standing at room temperature, and taken-up in 10 u.l of 2X sample buffer. These samples are boiled 2 min prior to loading on an SDS-PAGE gel. 2.6.3 Protein quantification Protein concentrations were determined using a commercially available version of the Bradford assay (Bio-Rad Protein Assay; Bradford, 1976). BSA standard (1 mg/ml) were obtained from Pierce. Various dilutions of the protein 84 samples were mixed with a five-fold dilution of Bio-Rad Protein Assay reagent, and absorbance read at 595 nm. Using the values of the standard curve obtained for BSA and the absorbances, the amount of protein in each sample was calculated. 2.6.4 Protein / SDS-PAGE gel Proteins were separated using the sodium dodecylsulfate-polyacrylamide gel electrophoresis (SDS-PAGE) method (Weber & Osborn, 1969). Samples were prepared by mixing with an equal volume of 2X sample buffer (120 mM Tris-HCI (pH 6.8), 3% (v/v) SDS, 10% (v/v) glycerol, 50 mM DTT, 0.1% (w/v) bromophenol blue), and boiling for 4 minutes. Unless otherwise noted, the gels were 7.5% (36:1) polyacrylamide (separating gel: 7.5% (w/v) polyacrylamide (36:1 acrylamide: N, N'-methylenebis-acrylamide), 400 mM Tris-HCI (pH 8.8), 0.1% (v/v) SDS, 5% (v/v) glycerol, 0.03% (w/v) ammonium persulfate, 0.16% (v/v) TEMED; s t a c k i n g g e l : 5% polyacrylamide (36:1 acrylamide: bis-acrylamide), 73 mM Tris-HCI (pH 6.8), 0.1% (v/v) SDS, 0.04% (w/v) ammonium persulfate, 0.2% (v/v) TEMED), and were placed in an electric field in Laemmli buffer (25 mM Tris, 192 mM glycine, 0.1% (v/v) SDS; Laemmli, 1970). Broad range SDS-PAGE standards (Bio-Rad) were run alongside the samples to serve as molecular weight markers. The separated proteins were then visualized by incubating the gel with Coomassie stain (0.5 mg/ml Coomassie® Brilliant Blue R-250 (Bio-Rad), 45% (v/v) methanol, 10% (v/v) acetic acid) and the background 85 stain removed with destain/fix solution (5% (v/v) acetic acid, 5% (v/v) ethanol). Bands containing 0.2 to 20 u.g of protein were detectable. 2.6.5 Western blot Proteins separated by electrophoresis as described above were transferred to a Trans-Blot Transfer Medium nitrocellulose membrane (Bio-Rad) at 250 mA for 2.5 hours in transfer buffer (3 mM Na 2 C0 3 , 10 mM NaHC0 3 , 20% (v/v) methanol) on ice (Dunn, 1986). The blot was incubated with shaking for 1 hour in PTBN (80 mM Na 2 HP0 4 , 20 mM NaH 2 P0 4 , 100 mM NaCI, 0.1% (w/v) BSA, 1 mM sodium azide, 0.05% (v/v) Tween 20, 5% (w/v) casein) and the primary antibody diluted 1:10,000. Excess antibodies were washed away by three 5-minute washes in 15 ml of PBS-2 (80 mM Na 2 HP0 4 , 20 mM NaH 2 P0 4 (pH 7.5), 100 mM NaCI). The blot was then incubated with stirring in PBS-2 and a 1:5,000 dilution of the secondary antibody (goat anti-rabbit HRP-labelled) for 1 hour. As before, excess antibodies were removed by three 5-minute washes in PBS-2. A chemiluminescent peroxidase substrate (4 ml) (ECL Western Blotting Detection Reagent, Amersham-Pharmacia) was added to the blot for 1 minute, and the protein bands detected on X-OMAT™ Scientific Imaging Film (Kodak). 2.6.6 Identification of proteins by mass spectrometry To unambiguously identify proteins, matrix-assisted laser desorption ionization (MALDI) analysis was used. The protein sample was alkylated with iodoacetamide and digested with trypsin as previously described (Kinter & 86 Sherman, 2000; UVic-Genome BC Proteomics Centre, Sample Preparation for Mass Spectrometry Analysis). Protein samples (20 ug) were loaded on an SDS-PAGE gel, the proteins electrophoretically separated, and stained using Coomassie Blue® (Section 2.6.4). Each protein band was carefully excised from the gel using a sharp razor blade, and placed in separate Eppendorf tubes. The gel slices were incubated with a series of successive reagents to generate trypsin-digested alkylated peptides as described below. Reagent 1 50% (v/v) methanol, 5% acetic acid 2 50% (v/v) methanol, 5% acetic acid 3 acetonitrile speedvac 4 10 mM DTT 5 100 mM iodoacetamide 6 acetonitrile 7 50 mM ammonium bicarbonate 8 acetonitrile speedvac 9 20 ng/ul trypsin, 50 mM ammonium bicarbonate 10 50 mM ammonium bicarbonate 11 50 mM ammonium bicarbonate get supernatant 12 50% (v/v) acetonitrile, 5% (v/v) formic acid combine supernatant with previous 13 50% (v/v) acetonitrile, 5% (v/v) formic acid combine supernatant with previous speedvac to < 20 ul adjust volume to 20 ul with 1 % (v/v) acetic acid Volume Time of Temperature incubation 200 ul 3 days room temp. 200 ul 4 hrs room temp. 200 ul 5 min room temp. 30 ul 30 min room temp. 30 ul 30 min room temp. 200 ul 5 min room temp. 200 ul 10 min room temp. 200 ul 5 min room temp. 30 ul 10 min ice 5ul overnight 37°C 30 ul 10 min room temp. 30 ul 10 min room temp. 30 ul 10 min . room temp. 87 The samples were submitted for MALDI mass spectrometry at the UVic-Genome BC Proteomics Centre (University of Victoria, Victoria, BC), and the peptide mass fingerprint analyzed using the software MASCOT ( to query the NCBInr 200330124 database (National Center for Biotechnology Information). 2.7 RNA PREPARATION and PROTOCOLS The two types of RNA used in this work were prepared by in vitro transcription. Intact RNAs and their degradative intermediates were separated on the basis of size on a denaturing polyacrylamide gel. 2.7.1 Preparation of template DNA DH5a™ cells containing the appropriate plasmid (pCH77 or pGM79) were grown in 25 ml of LB broth with selective antibiotics overnight, and recovered by centrifugation as outlined in Section 2.4.4. The plasmid DNA was extracted by alkaline lysis as described in Sambrook et al. (1989). First, the pellet was resuspended in 1 ml of GET buffer (25 mM Tris-HCI (pH 8.0), 50 mM glucose, and 10 mM EDTA), and left at room temperature for 5 minutes. Two milliliters (2 ml) of cooled a l k a l i n e s o l u t i o n (0.2 N NaCI and 1 % SDS) was then gently mixed in with the cells, and left on ice for 10 min. Potassium acetate solution (1.5 ml) (3 M potassium acetate, 11.5% (v/v) acetic acid) was gently incorporated into the preparation, followed by another 10 minutes on ice. The lyzed cell preparation was centrifuged in a JA-20 rotor, 20 min at 19,000 X g. The supernatant was 88 then subjected to two rounds of extraction with phenol-chloroform-isoamyl alcohol (25:24:1). Centrifugation in a Heraeus centrifuge enabled clearer separation of the top (aqueous) layer from the bottom (phenol) phase. The aqueous layer was extracted, and precipitated in 2.5 volumes of 95% ethanol by leaving the mixture at room temperature for 10 min. The pellet recovered after centrifugation in a JS-13.1 rotor for 20 min at 16,000 X g was washed with 80% ethanol, and re-isolated by centrifugation. The pellet was then air-dried at room temperature and re-suspended in 400 uJ of TE buffer (10 mM Tris-HCI (pH 8.0), 1 mM EDTA). This suspension was treated with 2 pJ of 2 mg/ml RNase A at room temperature for 1 hour. The enzyme was then removed by phenol-chloroform-isoamyl alcohol extraction as described above, and precipitated with ethanol. The final pellet was resuspended in 400 uJ of diethyl pyrocarbonate (DEPC)-treated water, and its DNA concentration measured by reading the absorbance at 260 nm. 2.7.2 Linearization of plasmids for transcription Typically, 30 u.g of plasmid DNA was digested with 135 units of the appropriate enzyme, using the conditions and buffers supplied by the manufacturer of the enzyme. Following cleavage, 5 mM EDTA and 2 M ammonium acetate were added to the reaction mixture, and the DNA extracted twice with an equal volume of phenol-chloroform-isoamyl alcohol (25:24:1). n(1)-butanol was added 1 ml at a time to dehydrate the sample (Cathala & Brunei, 1989). This was achieved by vortexing the mixture, spinning it in a microfuge to 89 separate the aqueous and organic layer, removing the remaining butanol, and redoing all of these steps until a white prepicitate is obtained. This precipitated sample was washed with 100 ul of water to dissolve some of the salts, and the butanol precipitation steps repeated. The DNA was resuspended in TE buffer to make a 1 ug/ul solution, and kept at 4°C until use. Confirmation of the cleavage was obtained by loading 1 ul of this preparation on a 0.8% agarose gel (0.8 % (w/v) agarose, 100 ml TAE buffer), and separating the DNA by electrophoresis at 75 volts for roughly 30 min in TAE buffer (40 mM Tris-HCI, pH 8.0, 20 mM sodium acetate and 1 mM EDTA). The DNA bands were visualized by staining in a solution of 0.5 ug/ml ethidium bromide in TAE buffer for 10 minutes at room temperature, and exposure to UV light. 2.7.3 In vitro synthesis of RNA transcripts The in vitro transcription of RNAs was performed as previously described, with slight modifications (Cormack & Mackie, 1992; Mackie & Genereaux, 1993). MalEF RNA The intergenic spacer region of the MalE-MalF mRNA (MalEF RNA) was synthesized in vitro over the course of 2 hrs at 30°C. Plasmid pCH77 (1.2 ng/ul) (McLaren e t al., 1991), linearized with EcoRI, served as template. Transcription was carried out using 1 U/ul of T7 RNA polymerase, in the presence of 0.5 uCi/ul [a 3 2P]-CTP (3000Ci/mmol), 0.1 mM CTP, 0.5 mM each of ATP, GTP, and UTP, 10 mM DTT, 1.5 U/ul of RNAguard (porcine), and 1X Promega transcription buffer (40 mM Tris-HCI (pH 7.5), 6 mM MgCl2, 2 mM 90 spermidine, and 10 mM NaCI). This reaction generates a 375 nucleotide radio-labeled transcript. The amount of MalEF synthesized was determined by calculating the percentage of radio-labeled nucleotide incorporated into the uniformly labeled RNA. To this end, the total radioactivity of the sample was measured by spotting a known volume of the reaction onto a glass filter (Whatman GF-C) and counting in a Beckman LS6000 IC scintillation counter for Cerenkov radiation. Separately, the same volume of reaction was precipitated in 5% (v/v) trichloroacetic acid on ice for 15 min, then collected by filtration on a glass fibre filtration, and counted. The ratio of radioactivities was used to calculate the yields. rpsT mRNA The rpsT mRNA, encoding ribosomal protein S20, was synthesized using 0.4 U/jJ of SP6 RNA polymerase, 45 ng/u.l of pGM79 digested with Dral as template (Mackie & Genereaux, 1993), 0.5 mM each ATP, GTP, UTP, and CTP, 10 mM DTT, 1X Promega transcription buffer, and 0.6 U/u.1 RNAguard (porcine) at 30°C for 1.5 hr. This reaction generates a 372 nucleotide unlabeled transcript. An identical reaction was carried out simultaneously, but with the addition of radiolabeled [a 3 2P]-CTP, to be able to estimate the amount of RNA transcribed (as above for MalEF). The transcripts produced were purified from enzymes and nucleotides by two phenol-chloroform-isoamyl alcohol (25:24:1) extractions, made 2 M in 91 ammonium acetate, and precipitated with 2.5 volumes of 95% (v/v) ethanol overnight at -20°C. The precipitated material was recovered by centrifugation, and the pellet washed with 80% (v/v) cold ethanol, bench dried, re-suspended in DEPC-treated distilled water, and stored at -20°C until use. 2.7.4 Electrophoretic analysis of RNA Four microliter (4 ul) samples were quenched with 12 ul of FA loading d y e s (90% deionized formamide [formamide was deionized by mixing for 2 hrs with Amberlite Monobed Resin (Bio-Rad), and then filtered], 22 mM Tris, 22 mM boric acid, 0.5 mM EDTA, 0.1% (v/v) SDS, 0.1% (w/v) xylene cyanol FF, and 0.1% (w/v) bromophenol blue), boiled for 2 minutes and set on ice. Five microliters (5 ul) were loaded on a 6% denaturing polyacrylamide gel (6% (w/v) polyacrylamide (29:1 acrylamide: N, N'-methylenebis-acrylamide), 90 mM Tris, 90 mM boric acid, 2 mM EDTA disodium salt, 0.5% (w/v) urea, polymerized with 0.033% (w/v) ammonium persulfate and 0.115% (v/v) TEMED), and the RNAs were separated electrophoretically in TBE buffer (90 mM Tris, 90 mM boric acid, 2 mM EDTA disodium salt). The nucleotides were fixed (Section 2.6.4), and dried under vacuum with heat. The dried gels were exposed to a Phosphorlmager screen and the radiolabeled RNA visualized using Phosphorlmager imaging technology. 92 2.8 TRANSMISSION ELECTRON MICROSCOPY Purified degradosomes (5 ul) were incubated with a similar volume of a ten-fold dilution of anti-Rne or anti-Pnp antibodies on ice for one hour. Half of this mixture was deposited onto a N i + + grid 200, and the excess liquid was absorbed with filter paper 1 minute later. Five microliters (5 ul) of a 50-fold dilution of goat anti-rabbit immunogold conjugate EM antibodies (BB international) was added to the N i + + grid and wicked off 1 minute later. The grid was washed by immersion into water, dried, and the proteins negatively stained using 5 ul of a 2% (w/v) uranyl acetate solution for 5 seconds. The samples were placed in a Hitachi H7600 transmission electron microscope. 2.9 IMMUNOPURIFICATION of DEGRADOSOMES from CELL EXTRACTS 2.9.1 Using antibodies to specific proteins Antibody-conjugated agarose beads Protein A agarose beads (300 ul) (Gibco BRL) were incubated with 4.5 ml of CLB buffer (20 mM Na 2 HP0 4 , 5 mM NaH 2 P0 4 , 0.2 mM NaCI, and 0.5 mM EDTA) and 15 ul of polyclonal antibodies directed against a specific degradosomal protein for 1 hr at 4°C with constant shaking. The beads were washed three times in 5 ml of CLB buffer, and the antibodies cross-linked to the beads for 1 hr at 4°C by the addition of 1% glutaraldehyde. The beads were washed three times in 5 ml of CLB buffer and then twice in 5 ml of IP buffer (50 mM Tris-HCI (pH 7.5), 100 mM NaCI, 1% nonidet-P40, 0.1% SDS, 0.5% sodium desoxycholate acid, 1 mM DTT, 1 mM 93 PMSF, 1 mM EDTA, 2 u.g/ml aprotinin, 0.8 u.g/ml leupeptin, and 0.8 u.g/ml pepstatin). They were then re-suspended in 570 u.l of IP buffer. Co-immunoprecipitation Cell pellets obtained from 10 ml cultures grown in the presence of [35S]-methionine were incubated with 250 u.l lysozyme-EDTA solution (50 mM Tris-HCI (pH 7.5), 100 mM NaCI, 3 mM 0.2 M EDTA, 5% glycerol, 1 mM DTT, 1.5 mg/ml lysozyme, 1 mM PMSF, 2 pg/ml aprotinin, 0.8 u,g/ml leupeptin, and 0.8 u.g/ml pepstatin). The incubation proceeded according to the following regiment: 10 min on ice, stirred for 1 min, 20 min on ice, stir for 1 min, and 40 min on ice. DNase-RNase-Triton buffer (166 u.l) (50 mM Tris-HCI (pH 7.5), 100 mM NaCI, 30 mM magnesium acetate, 5% glycerol, 1 mM DTT, 3% Triton X-100, 1 mM PMSF, 2 u.g/ml aprotinin, 0.8 u.g/ml leupeptin, 0.8 [ag/ml pepstatin, 20 u.g/ml DNase I, and 2 mU/ul RNase A) was then added to the mixture, and left on ice for 30 min. Ammonium chloride was added to 1 M concentration, and stirred in on ice for 30 additional minutes. The extract was spun in a cold room microfuge at full speed for 1 hr. The supernatant was collected and incubated with 190 u.l of beads prepared as described above. The cell extract was incubated with the beads for 1 hr at 4°C with shaking. The beads were washed 5 times with 500 uJ IP buffer, and the bound proteins extracted by incubation in 30 uJ of 2X sample buffer (Section 2.6.4) lacking DTT and dyes for 10 min at 55°C. The eluted proteins were collected and separated on an SDS-PAGE gel (Section 2.6.4). The proteins were visualized using Phosphorlmaging technology. 94 2.9.2 Using FLAG-Rne and anti-FLAG antibodies E. coli BL21 (DE3) strains containing pflagLRC or pRE196, and therefore expressing the FLAG epitope or N-terminally FLAG-tagged Rne, respectively, were grown in M9 minimal media supplemented with all the amino acids except methionine and cysteine, in the presence of 150 ug/ml ampicillin at 37°C to an OD6oo = 0.4. A 5 ml aliquot was pulse-labeled with 50 uCi of [35S]-methionine (NEN, 1000 Ci/mmol) and 1 mM IPTG for 15 minutes at 37°C, then washed by adding 5 ml of ice-cold minimal media containing 0.5% (w/v) sodium azide and 0.3% (w/v) methionine to stop incorporation of the label. The cells were quick frozen in an ethanol/dry-ice bath. The remainder of the culture was grown at 15°C for 2 hours, at which point 5 ml was removed and pulse-labeled for 30 minutes at 15°C as described above. Following cell isolation, a crude degradosome preparation was carried out. Labeled cells were lyzed in 250 ul of lysozyme-EDTA buffer (Section 2.9.1), and mixed on ice periodically over the course of an hour. DNase-RNase-Triton buffer (125 ul) (Section 2.9.1) was then added and left on ice for an additional 30 minutes. One molar (1 M) NH4CI was then stirred in on ice over the course of 30 minutes. Cell debris were pelleted by centrifugation for 15 minutes at 16,000 X g. The supernatant was precipitated with (NH4)2S04 to 40% of saturation, recovered by centrifugation for 10 minutes at 16,000 X g, and suspended in 500 ul of TBS buffer (50 mM Tris-HCI (pH 7.4), 150 mM NaCI). Immunoprecipitation with 40 ul of anti-FLAG M2-agarose affinity gel (Sigma) was performed for 2 95 hours at 4°C with gentle agitation. The beads were washed 3 times in 500 u.1 of TBS buffer, and then incubated for 30 min at 4°C with 100 uJ of a 150 ug/ml FLAG-epitope (Sigma) solution to elute bound proteins. Fifteen microliters (15 uJ) of this immunoprecipitate was resolved on a 7.5% SDS-PAGE gel (Section 2.6.4). The dried gel was exposed to Phosphorlmager screen for identification of the bands. 2.9.3 Using His-Rne and a TALON Column Rne16-15 cells were grown in 150 ml of LB broth containing antibiotics at 37°C until mid-log phase (OD60o = 0.4). The culture was then divided equally into 2 flasks. One culture was induced with 1 mM IPTG, 75 ml of fresh antibiotic-containing LB was added, and the culture grown at 30°C for 2 hrs. Chloramphenicol was added to a concentration of 160 jag/ml prior to harvesting to prevent translation of cold shock proteins during cell pelleting. The other culture was supplemented with 75 ml cold antibiotic-containing LB and incubated with vigorous shaking at 15°C for 1 hr. Following this adaptation period, the culture was induced with 1 mM IPTG, grown for another 2 hrs at 15°C, and harvested. Chloramphenicol (160 u.g/ml) was added to these cells as well. Cells were recovered by centrifugation as described in Section 2.4.4, washed in cell wash buffer (50 mM Tris-HCI (pH 7.5), 100 mM NaCI, and 5% glycerol, 160 u.g/ml chlorampheniocl), and re-suspended in 1.25 ml of IP buffer (Section 2.9.1) supplemented with 160 u.g/ml chloramphenicol. The cells were lyzed by adding 3 mg of lysozyme and then passing them twice through a French pressure cell at 96 8,000 psi. DNase I was added to 20 ug/ml, and the cell debris removed by centrifugation at top speed in a microfuge for 10 min at 4°C. The supernatant was passed through a column filled with 3 ml of a slurry of TALON metal affinity resin (Clontech). The column was washed with 20 ml TALON wash buffer (50 mM Tris-HCI (pH 8.0), 500 mM NaCI), and then with TALON wash buffer supplemented with 5 mM imidazole. The bound proteins were eluted with 8 ml of TALON wash buffer supplemented with 50 mM imidazole. The proteins recovered were separated on an SDS-PAGE gel (Section 2.6.4) and visualized by Western blotting (Section 2.6.5). 2.10 CO-IMMUNOPRECIPITATION EXPERIMENTS USING PURIFIED PROTEINS 2.10.1 Proteins without tag Antibody-conjugated beads were prepared as described in Section 2.9.1. One and a half micrograms (1.5 ug; amount subject to change - see Results Section for detail) of the appropriate proteins were mixed in the presence of pnp assay buffer (Section 2.11.2) supplemented with 0.05% Genapol X-080, and incubated at 30°C for 20 min. Ten microliters (10 ul) of this mixture was added to 190 ul of antibody-conjugated beads prepared as described above, and left at 4°C with constant shaking for 1 hr. The beads were washed and the bound proteins eluted as described in Section 2.9.1. The eluted proteins were separated on an SDS-PAGE gel (Section 2.6.4). Western blotting was used to visualize the proteins (Section 2.6.5). 97 2.10.2 Using FLAG-RNase E In vitro reconstitutions were performed by mixing partially purified FLAG-Rne (100 ng of total protein from an AS26 fraction, Section, 5-20 ug of purified His-Pnp, and 5 u.g purified enolase or 20 u,g of purified His-CsdA in a 200 u,l volume composed of buffer RNE-A, 7.5% (v/v) glycerol, 50 u.M DTT, 2 u.g/ml aprotinin, 0.8 u.g/ml leupeptin, and 0.8 u.g/ml pepstatin A at 30°C. To pull down FLAG-Rne, 40 uJ of anti-FLAG M2-agarose affinity gel was added to the mixture, and incubated with gentle agitation at 4°C for 2 hours. The beads were recovered by gentle centrifugation (3,000 rpm in a microfuge) and then washed 3 times in 500 u.l of TBS buffer, and incubated for 30 min at 4°C with 100 u.l of a 150 u.g/ml FLAG-epitope solution to elute bound proteins. Ten microliters (10 uJ) of this solution was loaded on a 7.5% SDS-PAGE gel, and blotted to a nitrocellulose membrane. This blot was probed using anti-Rne, anti-Pnp, and anti-enolase or anti-CsdA rabbit polyclonal antisera (Section 2.6.5). 2.11 RECONSTITUTION ASSAYS In order to examine the activity of degradosomes reconstituted from purified proteins, in vitro assays were performed. By selecting appropriate substrate RNA and buffer components, it is possible to monitor the activity of only RNase E, PNPase, or RhIB in the degradosome. 98 2.11.1 RNase E assay Assays were assembled in a 40 ul volume as described previously (Mackie, 1991; Mackie & Genereaux, 1993), with slight modifications. In vitro-transcribed radiolabeled rpsT mRNA (20 nM) (Section 2.7.3) was refolded by incubating the transcripts in Rne assay buffer (25 mM Hepes-NaOH (pH 7.6), 5 mM MgCI2, 5% glycerol, 60 mM KCI, 100 mM NH4CI, 0.1 mM EDTA, 0.1 mM DTT, 0.1 mg/ml BSA, 0.05% (v/v) Genapol X-080) for 2 min at 50°C, 10 min at 37°C, and on ice for 5 min. Enzymes (1 ug/ml purified Rne, ±1 ug/ml Pnp, and ±1 ug/ml RhIB or His-CsdA) were added at the "0" time point. Due to the lack of phosphate and ATP in the reaction buffer, Rne is the only enzyme capable of activity on rpsT. The reaction mixture was incubated at 30°C, and 4 ul aliquots removed at 0, 1, 2, 5, 10, 20, 30, 45, and 60 min. These aliquots were plunged in 12 (al of FA loading dyes (Section 2.7.2), and kept on ice until the end of the experiment. They were then boiled for 2 min and electrophoretically separated on a 6% denaturing polyacrylamide gel (Section 2.7.2). The radiolabeled RNA was visualized by Phosphorlmaging. 2.11.2 PNPase and RhIB assay Assays were assembled in a 40 ul volume as described previously (Py et al., 1994) with slight modifications. The MalEF RNA (1.6 nM) was mixed with 1 ug/ml purified Rne protein, 2 ug/ml Pnp, and 1 ug/ml RhIB or His-CsdA in p n p assay buffer (20 mM Tris-HCI (pH 7.5), 1.5 mM DTT, 1 mM MgCI2, 20 mM KCI, 10 mM sodium phosphate, 0.1 mg/ml BSA, 0.05% (v/v) Genapol X-080), in the 99 presence or absence of 3 mM ATP. The enzymes were pre-incubated together for 15 minutes at 15°C or 30°C to regenerate a "minimal degradosome" (Coburn et al., 1999; Mackie et al., 2001) prior to addition of RNA. The reaction was incubated at 15°C or 30°C. Samples of 4 u.l were removed from the assay at 0, 1, 2, 5, 10, 20, 30, 45, and 60 min after mixing of RNA and enzymes, and quenched with three volumes of FA loading dyes. Portions of these samples were electrophoretically separated on a 6% polyacrylamide gel containing 8 M urea (Section 2.7.4), and visualized using Phosphorlmager technology. 2.12 ATPASE ASSAYS The ATP hydrolytic activity of DEAD-box helicases was assayed by measuring the release of [ 3 2P]-Pi from [y 3 2P]-ATP. Reactions were assembled in a 10 ul volume, and contained 10.3 nM of helicase (750 ng/ml of CsdA or 4.8 u.g/ml of degradosome), pnp assay buffer (Section 2.11.2), 0.1 mM ATP, 2.5 u.Ci [y 3 2P]-ATP (3000 Ci/mM), 20 nM in vitro transcribed rpsT mRNA (Section 2.7.2, Mackie & Genereaux, 1993). Some assays also contained 150 pg DNA, and up to 7.5 (^g/ml of Rne protein. Assays were assembled on ice, and the reactions started by the addition of helicase at the 0 min time point. The mixture was incubated at 15°C or 30°C. At the 0, 15, 30, 45, and 60 min time points, a 1 u.l aliquot was removed and spotted on a poly(ethylenimine) (PEI)-cellulose thin layer chromatography (TLC) 100 plate. This plate was developed in 0.375 KH2PO4 (pH 3.5). The plate was then dried and exposed to Phosphorlmager to quantify the phosphate released. 2.13 E N O L A S E A S S A Y Enolase dehydrates 2-phoshoglycerate to produce phosphoenolpyruvate. The latter chemical absorbs at 240 nm. This property is exploited to measure the glycolytic activity of enolase (Spring & Wold, 1971; Dannely et al., 1989). In this spectrophotometry assay, 1 ml of enolase assay buffer (50 mM Tris-HCI (pH 8.1), 100 mM KCI, 1 mM 2-phosphoglycerate, 1 mM MgS0 4 , 10 uM EDTA) is placed in a spectrophotometer cuvette. The reading is set to 0, and the timer is started when a set amount of enzyme is added. Spectrometric readings are taken at a wavelength of 240 nm every 10 sec for 9.5 min, and then more sparsely until 45 min. 2.14 COMPUTATIONAL METHODS The RhIB (GenBank accession P24229) and CsdA (GenBank accession P23304) sequences were aligned using ClustalW through the European Bioinformatics Institute WWW service (; Higgins et al., 1994). The results were viewed using the Multiple Sequence Alignment Editor and Shading utility GeneDoc Version 2.6.001 (Nicholas & Nicholas, 1997). Determination of sequence identity and similarity was done using the National Center for Biotechnology Information (NCBI) pairwise BLAST program BLAST 2 101 Sequences, available on the world wide web (; Tatusova & Madden, 1999). 102 CHAPTER III ENOLASE & THE RNA DEGRADOSOME 3.1 BACKGROUND Enolase has consistently been isolated as part of the degradosome using a variety of purification methods (Py et al., 1996; Miczak et al., 1996; Coburn & Mackie, 1998). It interacts with residues 739-845 in the C-terminal tail of RNase E, as determined by the yeast two-hybrid method and immunoprecipitations (Vanzo etal., 1998; Kuhnel & Luisi, 2001). A potential homolog is also present in the degradosome of Fthodobacter (Jager et al., 2001). These findings suggest that the association between enolase and RNase E is specific, and therefore biologically significant. What role could enolase have in the degradosome? There are currently three hypotheses. The first, prompted by the discovery that yeast enolase has the ability to bind polynucleotides (Al-Giery & Brewer, 1992) and some anecdotal reports that E. coli enolase possesses this property as well (Kuhnel & Luisi, 2001), proposes that enolase serves as an accessory RNA binding protein to assist the degradosome in substrate recognition and/or retention (Kuhnel & Luisi, 2001). To date, no enolase-RNA interaction has been demonstrated using E. coli components in vitro (Py et al., 1996; Kuhnel & Luisi, 2001). Nonetheless, the idea is appealing and may require the serendipitous discovery of the appropriate RNA. The second hypothesis, which may be related to the first, suggests that enolase is involved in 5'-end-independent substrate recognition by RNase E (Section; Marchand et al., 2001; Leroy et al., 103 2002; G.A. Mackie, personal communication). Enolase may promote substrate recognition directly, or else be required to fold a region of RNase E into a conformation that allows RNase E to contact a substrate. This is a relatively new idea that awaits further experimental confirmation. The last hypothesis, which is the basis for the experiments described in this chapter, postulates that enolase is involved in sensing the cell's metabolic state and modulates the activity of the degradosome in response to changes in the environment (Kuhnel & Luisi, 2001). 3.2 RESULTS The experiments described in this chapter sought to answer three main questions: • Can the presence of enolase modulate the activity of RNase E, PNPase, or RhIB? • Does enolase respond to specific signals to affect the activity of RNase E, PNPase, or RhIB? • Does enolase physically associate with RNase E? 3.2.1 Enolase Purification and Activity Enolase was prepared as described by Kuhnel & Luisi (2001), with some modifications (Section The purity of the enzyme is shown on a gel in Figure 2.1. The glycolytic activity of enolase was determined by incubating varying amounts of the enzyme with 1 mM 2-phosphoglycerate in enolase assay buffer (Section 2.13). The conversion of 2-phosphoglycerate to 104 phosphoenolpyruvate was monitored spectrophotometrically at 240 nm. This assay is based on the finding that phosphoenolpyruvate absorbs at 240 nm, but 2-phosphoglycerate does not (Winstead & Wold, 1966; Spring & Wold, 1971). The initial (linear) rates of conversion using varying amounts of enzyme are shown in Figure 3.1, panel A. 1 unit of enolase activity is defined as the amount of enzyme required to cause a change in A 2 4o of 0.1/min (Spring & Wold, 1971), and for this enolase preparation corresponds to roughly 0.5 ug (or 2,000 units/mg). A reaction using this amount of enzyme was allowed to proceed to equilibrium, and is shown in panel B of Figure 3.1. 3.2.2 RhIB Purification Purified RhIB is needed for the in vitro experiments described below. The previously established RhIB purification protocol (Coburn et al., 1999) did not yield pure RhIB protein in this experimenter's hands; therefore, a novel purification protocol was devised. Cultures of GC300 were grown in LB broth containing 100 ug/ml ampicillin at 30°C, and induced for 3 hours with 0.5 mM IPTG. The cells were harvested by centrifugation, washed, and suspended in buffer RHLB-A (50 mM Hepes-NaOH (pH 7.5), 0.1 mM EDTA, 2 mM DTT, 0.1 mM PMSF, and 5% (v/v) glycerol). Cell lysis was carried out by two passages through a French pressure cell at 8,000 psi. DNase I (2.3 ug/ml) was added prior to centrifugation at 30,000 X g for 45 min in a Beckman JA-20 rotor to yield an S30. This supernatant was loaded at a rate of 0.5 ml/min onto a Resource S column (Amersham Biosciences) using an FPLC. Most proteins did not bind to 105 0 100 200 300 400 500 time (sec) time (sec) Figure 3.1 Assay of enolase activity The conversion of 2-phosphoglycerate to phosphoenolpyruvate by purified enolase was monitored by measuring the absorbance at 240 nm as described in Section 2.13. A. Initial rates of reaction catalyzed by varying amounts of enolase. (•) 0.35 ng/ml, (•) 0.5 u.g/ml, (A) 0.75 ,ug/ml, (•) 5 ug/ml. B. 0.5 u,g/ml of purified enolase was used to characterize a reaction over a 45 min period. 106 this cation exchanger and were recovered in the flowthrough. The column was washed at a rate of 1 ml/min with buffer RhIB-B (25 mM Tris-HCI (pH 8.0), 5% (v/v) glycerol, 0.1 mM EDTA, and 1 mM DTT) with 50 mM NaCI until the A 2 6o of the wash returned to baseline levels, and the bound proteins were eluted with a salt gradient from 50 to 500 mM NaCI at a flow rate of 1 ml/min over 100 ml. Two milliliter (2 ml) fractions were recovered, with RhIB eluting in fractions 12 through 19. Figure 3.2 illustrates how this one step successfully separated RhIB from most other proteins of the S30 fraction. Fractions containing crude RhIB (e.g. fractions 13-18 in Figure 3.2) were pooled and concentrated with a Millipore centrifugal filter device (MWCO: 10K) before loading onto a size exclusion column. A Superdex 75 column (Amersham Biosciences; 3,000 to 80,000 MR fractionation range for a globular protein) was used with an FPLC. Proteins were eluted at a flow rate of 0.75 ml/min in buffer RhIB-B with 50 mM NaCI, and collected in 2 ml fractions. Fractions 8, 9, and 10 were pooled and applied to a HiTrap® Blue 1 ml column (Amersham Biosiences). The bound proteins were eluted using 30 ml of a gradient consisting of buffer RhIB-B with 50 mM NaCI that progressively changed to buffer RhIB-B with 3 M KCI. RhIB eluted in fractions 10, 11, and 12 of the 2.5 ml fractions, and was concentrated using Millipore centrifugal filter devices (MWCO: 10K) to 0.5 ml. By Bradford assay (Section 2.6.3) the protein concentration was 107 Figure 3.2 RhIB purification on Resource S column The S30 fraction from induced GC300 was passed over a Resource S column as described in Section 3.2.2. 10 of the flowthrough and from alternative fractions 2 through 26 were separated on a 7.5% S D S - P A G E gel (Section 2.6.4). 108 established at 3 mg/ml. A gel showing the purity of the final protein purification is shown in Figure 2.1. 3.2.3 Addition of enolase to in vitro degradosome assays Purified RNase E, PNPase, and RhIB are known to assemble into a functional degradosome in vitro (Coburn etal., 1999). Enolase has thus far been omitted from these reconstituted complexes. To ascertain whether the presence of enolase can influence the activity of each of the degradosome protein, 1 ug/ml purified Rne, 1.5 ug/ml Pnp, and 1 ug/ml RhIB were incubated together in either rne or pnp assay buffer (Section 2.11) in the presence or absence of 1 ug/ml enolase. The activity of these reconstituted complexes was then assayed on 20 nM of in v/Yro-transcribed radiolabeled RNA substrate (MalEF RNA or rpsT mRNA) (Section 2.11). A schematic diagram of the enzymatic reactions and intermediates created in each assay is shown in Figure 3.3. As a control, purified enolase was incubated with MalEF RNA for 60 minutes in pnp assay buffer at 30°C. Figure 3.4, panel A shows that both the buffer solution and the enolase preparation are free of contaminating nuclease activity. The effect of adding enolase on the activity of RNase E was assayed on the rpsT mRNA, a well-characterized substrate for RNase E. The presence of enolase did not influence either the disappearance of intact rpsTtranscript or the accumulation of intermediates (results not shown). PNPase was then added to 109 Rne Rne rpsT mRNA B C MalEF RNA PNPase PNPase MalEF RNA * Intermediate (some stalling) * Intermediate (some stalling) REP-Stabil ized RNA (RSR) (stable intermediate) REP-Stabil ized RNA (RSR) (some stalling while RhIB works) Hairpin is destabilized and degraded Entire RNA degraded Figure 3.3 Diagram of the reactions and intermediates created in reconstitution assays A. RNase E cleavage of rpsT substrate. The full-length rpsT mRNA incubated with RNase E and PNPase is resistant to PNPase attack, due to the lack of phosphate in the reaction buffer (Section 2.11.1). RNase E makes two major endonucleolytic cleavages to create three stable intermediates. B. PNPase activity against MalEF RNA. The intercistronic region of the MalEF RNA is incubated with RNase E and PNPase . MalEF is a poor substrate for RNase E. As the buffers contain phosphate (Section 2.11.2), PNPase can attack the 3'-end of the substrate and remove nucleotides exonucleolytically. The smaller hairpin denoted * stalls PNPase for a short period. However, PNPase overcomes this stem, only to be blocked at the larger R E P stem-loop. PNPase cannot overcome this stem on its own, leading to the accumulation of a stable intermediate called R S R . C. RhIB activity against MalEF RNA. This assay resembles the PNPase assay but also contains the helicase RhIB and ATP. As before, PNPase stalls at the R S R intermediate, but RhIB uses the energy derived from the hydrolysis of A T P to unwind the stem-loop, creating single-stranded RNA which is a substrate for PNPase . The RSR accumulates for a short while until RhIB separates the strands and PNPase is able to degrade the entire RNA. 1 1 0 A B 20 40 60 time (minutes) 80 20 40 60 time (minutes) 20 40 60 time (minutes) 20 40 60 time (minutes) 20 40 60 time (minutes) Figure 3.4 Effect of enolase on the activity of degradosome proteins in vitro The activity of 1 u.g/ml purified Rne, 1.5 u,g/ml Pnp, and 1 (ag/ml RhIB on 20 nM in w'fro-transcribed radiolabeled RNA was assayed in the presence and absence of 1 uc|/ml purified enolase (Section 2.11). Each assay was performed at least twice, but only one representative assay from each condition is shown in this figure. A. Control reaction in which MalEF RNA was incubated with pnp assay buffers in the absence (O), or presence (•) of enolase. B. Effects of enolase on the activity of RNase E assayed on r p s T mRNA. (X) - PNPase; (*) - RNase E; (O) - RNase E, and PNPase; (•) - RNase E, PNPase, and enolase. C & D. Effect of enolase on the activity of PNPase assayed on the MalEF RNA. The disappearance of intact MalEF RNA is plotted in panel C, while the appearance and accumulation of RSR intermediate is shown in D. (O) - RNase E and PNPase; (•) - RNase E, PNPase, and enolase. E & F. Effect of adding enolase on the activity of RhIB assayed on the MalEF RNA. The disappearance of intact MalEF RNA is plotted in panel E, while the appearance and accumulation of RSR intermediate is shown in F. (O) - RNase E, PNPase, RhIB, and ATP; (•) - RNase E, PNPase, RhIB, ATP and enolase. 111 this assay to test whether enolase might require PNPase to exert an effect on RNase E. Due to the absence of phosphate in the rne assay buffer (Section 2.11.1), PNPase is unable to attack free 3'-ends. As shown in panel B of Figure 3.4, PNPase alone (X) could not degrade the rpsT transcript. As anticipated, RNase E did cleave the rpsT mRNA (*), and this process was not stimulated by the presence of PNPase ( O ) . Finally, the addition of enolase to RNase E and PNPase (•) did not further stimulate or inhibit the disappearance of intact rpsT substrate. To test the effects of enolase on PNPase activity, the MalEF RNA was used. This transcript is resistant to RNase E activity, and is therefore well suited to studies of the activity of PNPase in the degradosome (Section 2.11.2). The results of assays monitoring PNPase activity in the presence and absence of enolase are shown in Figure 3.4. The disappearance of intact substrate is shown in panel C, while the simultaneous accumulation of RSR intermediate is plotted in panel D. The degradation of intact MalEF and the appearance of RSR intermediate by RNase E and PNPase ( O ) appears to be no different from the decay and accumulation of these RNAs by RNase E, PNPase, and enolase (•). Finally, this same assay was repeated in the presence of RhIB and ATP to determine whether the activity of the helicase is affected by the presence of enolase. In this assay, the RSR intermediate initially accumulates, but then disappears as RhIB is presumed to unwind the RSR hairpin, enabling PNPase to 112 digest through the stem-loop. The disappearance of intact MalEF was unaffected by the presence (•) or absence (O) of enolase (Figure 3.4, panel E). Similarly, enolase had no effect on the accumulation and subsequent disappearance of the RSR intermediate (Figure 3.4, panel F). 3.2.4 Effects of enolase ligands on degradosome activity If enolase is the sensor of the metabolic state of the cell, it may need to interact with or be modified by regulatory molecules in order to exert an effect on other degradosomal proteins. Thus, the next step was to test the activity of the degradosome in the presence of enolase ligands. The glycolytic substrate (2-phosphoglycerate) and product (phosphoenolpyruvate) of enolase are known to cause a shift in the conformation of the enzyme upon binding (Lebioda et al., 1989; Duquerroy etal., 1995; Kuhnel & Luisi, 2001). Since the ratio of these two ligands could vary with the metabolic state of the cell, binding of one of these to enolase could specifically alter the activity of the degradosome. Assays were performed to monitor the activity of both RNase E and PNPase in the presence of 1 mM 2-phosphoglycerate. Five jag/ml intact purified degradosomes (Section provided the source of enzymes (this corresponds to 3 nM degradosome, assuming a molecular weight of 1,500 kDa, and implies the presence of excess ligand). Panels A through D of Figure 3.5 show the effects of adding 2-phosphoglycerate on the activity of RNase E (panels A and B) and PNPase (panels C and D). Purified enolase did not show a 113 r UJ H < CC LU O > _l (5 O X 0. CO o I E I C\J RNase E Assay PNPase Assay r LU H < > DC > UJ O X D_ CO o X 0. RNase E Assay PNPase Assay v . Q O CO LU Q X O =3 o ^ PNPase Assay 50 T 200 < CC c 150 , ' c ! o n j ' « E 100 | CD < DC 50 -0 -20 40 60 80 Time (minutes) 0 20 40 60 80 Time (minutes) Figure 3.5 Effect of enolase ligands on degradosome activity 1 mM 2-phosphoglycerate was added to 5 jxg/ml degradosome in assays designed to monitor RNase E (panels A and B), or PNPase (panels C and D) activity. Using a similar protocol, the effects of 1 mM phosphoenolpyruvate were also tested in panels E & F for RNase E activity, and panels G & H for PNPase activity. Finally, I and J show the results of an assay where 1.5 mM sodium fluoride was added to the reaction mixture. All graphs on the left represent the amount of intact RNA remaining over time, while those on the right show the appearance and accumulation of stable intermediates. (X) are controls where degradosomes were omitted, but the tested ligand was present; (O) represents the assay performed in the absence of ligand, and (•) represents the results of an assay performed in its presence. 114 significant amount of nuclease activity (X), as expected. The disappearance of the rpsT substrate, used to monitor RNase E activity, was not affected by the presence (•) or absence ( O ) of 2-phosphoglycerate (panel A). However, the accumulation of the 3'-terminal 147-nucleotide Rne decay intermediate reached slightly more elevated levels in the presence of the enolase ligand (panel B). The activity of PNPase was examined using the MalEF RNA. The disappearance of intact MalEF was unaffected by the presence (•) or absence ( O ) of 2-phosphoglycerate (panel C). However, the RSR intermediate accumulated to slightly increased levels in the presence of 2-phosphoglycerate (panel D). Panels E through H of Figure 3.5 show the effects of adding 1 mM phosphoenolpyruvate on the activities of RNase E (panels E and F) or PNPase (panels G and H). The addition of this compound had no observable effect on the disappearance of intact substrate by either RNase E or PNPase. However, its presence is correlated with a slight reduction in the accumulation of intermediates as a result of the activity of either enzyme. Sodium fluoride (NaF) is a known inhibitor of the glycolytic activity of enolase. In the presence of phosphate, F" forms fluorophosphate and associates with magnesium ions in the enolase active site, inhibiting the enzyme (Guminska & Sterkowicz, 1976; Maurer & Nowak, 1981; Lebioda etal., 1993). Accordingly, 5 ug/ml intact purified degradosomes were assayed in the presence (•) and 115 absence ( O ) of 1.5 mM NaF. Since the buffer used to study RNase E activity does not contain phosphate, which is essential for the inhibition reaction, only the activities of PNPase and RhIB on the MalEF RNA were assayed. The results are shown in panels I and J of Figure 3.5. The addition of NaF did not affect the activity of either PNPase or RhIB. 3.2.5 Physical Association of RNase E and enolase in vitro Degradosomes reconstituted in vitro from purified proteins in the presence of enolase were used in the assays described in Section 3.2.3 to assess whether enolase can modify the function of other degradosome proteins. These assays assume that purified enolase assembles with RNase E into a degradosome in vitro, which is not unreasonable given the ease with which purified RNase E, PNPase, and RhIB reconstitute into a functional degradosome (Coburn et al., 1999). To confirm that re-association takes place, 1.5 ug of purified RNase E and enolase were incubated together in pnp assay buffer at 30°C for 15 minutes (Section 2.10.1), and then immunopurified using either anti-Rne or anti-enolase antibodies. Each protein was also incubated alone with each antibody to determine the specificity of the interaction. Figure 3.6 shows that the anti-Rne antibody interacts with RNase E (lane 1), but not with enolase (lane 2). This antibody pulled-down RNase E and a small quantity of enolase when these two proteins were present in the assay (lane 3). Conversely, the anti-enolase antibody did not cross-react with purified RNase E (lane 4), but did immunoprecipitate enolase (lane 5). However, enolase was the only protein 116 RNase E antibodies enolase antibodies si o 0) Rne enolase Rne Rne enolase Rne enolase enolase Antibody used in IP Proteins Included anti-Rne - — w anti-enolase v 5 Figure 3.6 In vitro association of enolase with RNase E Anti-Rne- and anti-enolase-conjugated beads were incubated with either 1.5 ug purified RNase E, 1.5 ug purified enolase, or a mixture of both proteins in pnp assay buffer for 2 hrs (Section 2.10.1). The proteins retained on the beads were separated by SDS-PAGE electrophoresis and identified by Western blot. The top portion of the figure represents the result of a Western blot probed with anti-Rne antibodies, while the bottom section shows the result of probing the blot with anti-enolase antibodies. 117 brought down by this antibody when it was incubated with a mixture of the two proteins (lane 6). The presence of purified PNPase, RhIB, DnaK, or ATP in the assay could not strengthen the interaction between RNase E and enolase (results not shown). 3.2.6 Physical Association of RNase E and enolase in vivo Since a direct assay hinted at a weak interaction between purified RNase E and enolase, the presence of enolase in degradosomes purified by different means from cell extracts was re-examined. To confirm previous results, a degradosome preparation was isolated by the biochemical method described in Section A band of the size expected for enolase co-purified with this complex (Figure 3.7, panel A),, and its identity was confirmed by mass spectrometry (Section 2.6.6). The presence of enolase in the degradosome preparation shown in panel A of Figure.3.7 could be due to the fortuitous isolation of both degradosomes' and enolase by the protocol employed. Degradosomes were therefore purified using alternative methods. Anti-Rne antibodies were used to pull-down radiolabeled proteins from an E. coli extract (Section 2.9.1). A protein with a molecular weight matching that of enolase was detected in the immunoprecipitate (Figure 3.7, panel B). Enolase also co-purified with His-RNase E on a TALON column (Section 2.9.3). In this case, the protein's identity was confirmed by Western blot (Figure 3.7, panel C). Finally, cells capable of overexpressing FLAG-RNase E 1 1 8 B kDa 205—TI ^U—RNase E 121-70-\ 52-PNPase -RhIB -enolase D FLAG-FLAG Rne >, K I— FLAG -RNase E -PNPase -RhIB -enolase anti-Rne anti-Pnp anti-enolase Figure 3.7 Degradosomes purified from cell extracts by different methods all contain enolase A. Biochemical purification of degradosomes (Section Lane 1 contains size markers, lane 2, purified degradosomes. B. Degradosomes immunoprecipitated using anti-Rne antibodies (Section 2.9.1). C. Western blots of degradosomes purified by binding His-RNase E to a TALON column (Section 2.9.3). D. Degradosomes in a FLAG-Rne overexpressing strain were "pulled-down" using anti-FLAG beads (Section 2.9.2). The FLAG-Rne construct was overexpressed in wild-type and pnp::Tn5 backgrounds while labeling with P5S]-methionine for 15 minutes once cells reached log phase. The identity of the proteins co-purifying with FLAG-Rne in the wild-type background was confirmed by Western blotting using appropriate antibodies (E). 119 were grown to log phase, induced, and then pulse-labeled with [35S]-methionine for 15 minutes. The cell extract was then immunoprecipitated using anti-FLAG beads. One of the proteins co-eluting with FLAG-RNase E cross-reacts with anti-enolase antibodies (Figure 3.7, panel E), and co-migrates with enolase, as judged by its molecular weight (Figure 3.7, panel D). Despite the finding that PNPase cannot improve the association of RNase E and enolase in in vitro co-immunoprecipitation experiments, it seemed appropriate to test whether PNPase might be involved in recruiting enolase to the degradosome. Enolase and PNPase have historically co-purified together (Portier, 1975; Carpousis etal., 1994; Py etal., 1996), and indeed seem to do so in the PNPase purification protocol described in Section (Figure 2.1). To test this assumption, the FLAG-RNase E pull-down was repeated in a pnp::Tn5 background, which lacks PNPase. Although the loading on the gel shown in panel D of Figure 3.7 between wild-type and pnp::Tn5 cells is uneven, the presence of enolase is observable in both immunoprecipitations. However, quantification of the amount of enolase, relative to the amount of RNase E in each lane, shows that only a third of the amount of enolase associates with RNase E in the absence of PNPase. This experiment was repeated in another context (Figure 4.6 panel C) and although the loading is different, the results are identical6. 6 Interestingly, deletion of PNPase resulted in a marked increase of RhIB (or the band whose size matches RhIB - its identity was never confirmed by Western blotting) in the immunopurified degradosome, relative to RNase E (6-fold in Figure 3.7, panel D, and 16-fold in Figure 4.6 panel C). Perhaps this is due to the fact that PNPase may normally regulate the amount of rhIB mRNA. 120 If PNPase promotes enolase binding to RNase E, then adding this protein to pnp mutant extracts should restore enolase in the degradosome. FLAG-tagged RNase E was partially purified (Section from cells with a pnp gene deletion (ENS134 pnpr.TnS). One hundred micrograms (100 u.g) total protein from this AS26 extract was immunoprecipitated using anti-FLAG beads, and the bound proteins identified by Western blot. Lane 1 of Figure 3.8 shows the presence of RNase E, RhIB, and enolase in the immunoprecipitate. The ability of PNPase to stimulate the incorporation of enolase into the degradosome was then tested in lanes 2 to 6. First, 5 u.g purified His-PNPase and enolase were incubated with anti-FLAG beads to confirm that these proteins do not cross-react with the support or the antibodies. His-PNPase showed no sign of interaction (lane 2), while purified enolase was slightly retained by the beads (lane 3). The addition of 5 ug purified His-PNPase to 100 ug of the AS26 extract from pnp deleted cells resulted in the incorporation of His-PNPase to the degradosome, as shown by the presence of this protein in the IP (lane 4). The addition of 5 u.g purified enolase to 100 ug of the AS26 extract did not increase the amount of enolase brought down by the anti-FLAG antibodies (lane 5), and this quantity could not be noticeably increased by the addition of 5 u.g purified His-PNPase (lane 6). 121 + + + _l_ _|_ i FLAG-Rne extract (Apnp) + + His-PNPase - + + enolase anti-Rne anti-Pnp anti-RhIB anti-eno Figure 3.8 Immunoprecipitation of the degradosome from Apnp cells ENS134 pnp::Tn5 cells overexpressing FLAG-RNase E from the plasmid pRE196 were lyzed and an AS26 extract prepared (Section 100 Lig total protein from this extract was passed over an anti-FLAG column and the bound fraction eluted with an excess of F L A G peptide. These proteins were electrophoretically separated and identified by Western blots. Each "row" of the figure was probed with the antibody labeled on the left. Lane 1 shows the proteins co-eluting with FLAG-Rne in the AS26 extract. Lane 2 is a control for the interaction of 5 up, of purified His-Pnp with anti-FLAG beads. Lane 3 is also a control for the interaction of 5 ^g of purified enolase with anti-F L A G beads. Lane 4 is the result of an immunoprecipitation after incubating 100 jxg of the AS26 extract with 5 ^g purified His-PNPase. Lane 5 is the result of an immunoprecipitation after incubating 100 jug of the AS26 extract with 5 pig purified enolase. Lane 6 shows the proteins remaining on the column after incubating 100 ^ g of the AS26 extract with 5 ug purified His-PNPase and 5 iig purified enolase. 122 3.3 DISCUSSION 3.3.1 Does enolase influence degradosome activity? Since the degradosome has always been reconstituted from purified components in the absence of enolase, it seemed appropriate, as a first step, to simply include enolase in the reconstitution and assay the activity of each protein. The addition of enolase failed to influence the activities of RNase E, PNPase, and RhIB. The intent of these experiments was to study possible changes in the activity of the degradosome in response to the inclusion of enolase in the complex. However, it is apparent from Figure 3.6 that purified enolase does not assemble with RNase E as readily as PNPase and RhIB do in vitro. This implies that the reconstitution assays described above may only test the effects of enolase present in trans on degradosome activity. This condition may be relevant to the cell, as only 5-10% of cellular enolase is estimated to be associated with the degradosome (Py etal., 1996). If the role of enolase is to adapt the degradosome to changing cellular conditions, it may require a signal to carry out these effects. Enolase ligands known to cause the enzyme to undergo structural changes were therefore included in assays. The addition of 2-phosphoglycerate or phosphoenolpyruvate produced no observable differences in the disappearance of intact substrates by either RNase E or PNPase present in the degradosome. However, the rapidity with which the reactions proceeded could hide a subtle effect. A more sensitive and specific measure of the activity of a nuclease is the accumulation of 123 intermediates generated specifically by that enzyme. The presence of 2-phosphoglycerate is correlated with an increased accumulation of intermediates in both RNase E and PNPase assays, while incorporation of phosphoenolpyruvate in assays caused the opposite effect. The assays were only performed once, so it is dangerous to ascribe too much significance to the small differences observed, but the opposite effects of the two ligands suggest that these compounds can affect the activity of the degradosome. The presence of 2-phosphoglycerate may stimulate the activity of both RNase E and PNPase, and phosphoenolpyruvate may have the opposite effect. From these results, it is impossible to determine whether such effects are mediated through enolase, but the ability of this protein to interact with these ligands suggests it may be involved. The addition of 1.5 mM NaF, which can bind the active site of enolase and inhibit its activity, to 3 nM degradosomes did not noticeably affect the activity of either PNPase or RhIB. Based on the fluoride inhibition index7, under the conditions of the PNPase assay, only one active enolase molecule is expected for every 30,000 inhibited enzymes, suggesting a very efficient inhibition. The effects of adding NaF should have been tested on assays where 2-phosphoglycerate and phosphoenolpyruvate were also present to determine whether NaF can counteract the effect of these ligands. 7 The fluoride inhibition index = inhibited enzymes * [Mg+2] * [HP04] * [FT / active enzyme, and for E. coli enolase has a value of 0.5 X 10"12 M (Spring & Wold, 1971). 124 Although not presented in the results section, some preliminary experiments with enolase phosphorylation were undertaken, but no effects of this modification on degradosome activity were detected. In summary, neither enolase alone, nor enolase ligands, nor potential enolase modifications affect the activity of the E. coli RNA degradosome under the conditions tested. 3.3.2 Does enolase bind RNase E? Purified RNase E, PNPase, and RhIB associate into a complex when incubated together in an appropriate buffer (Coburn et al., 1999). Since enolase is reproducibly isolated with these proteins in the cell (Figure 3.7; Py et al., 1996; Miczak et al., 1996; Coburn & Mackie 1996a), it seemed reasonable to expect that purified enolase would also reconstitute into a degradosome in vitro. Surprisingly, purified RNase E only brought down a small amount of enolase, and enolase did not co-precipitate any RNase E (Figure 3.6). This suggests only a modest association between RNase E and enolase in vitro. A third protein could mediate the interaction between RNase E and enolase in vivo. Given the history of association between enolase and PNPase (Portier, 1975; Carpousis et al., 1994; Py et al., 1996), a prime candidate for this role is PNPase. However, adding PNPase to in vitro assays did not stimulate the formation of a complex between RNase E and enolase (results not shown). In addition, enolase is present in the degradosome of pnp mutant cells (Figure 3.7, panel D and Figure 3.8), and the amount found in this complex cannot be detectably increased by 125 the addition of extraneous PNPase. These results show that enolase associate with RNase E in the absence of PNPase. There are some discrepancies between the immunoprecipitation experiments shown in Figure 3.7, panel D and Figure 3.8. The first demonstrates that although enolase can interact with RNase E in the absence of PNPase, the latter protein stimulates the interaction. The second experiment shows that PNPase is not required for optimal interaction between RNase E and enolase. The first assay is probably a more sensitive measure of the composition of the degradosome due to its use of radiolabel, instead of excess protein, to detect the immunoprecipitated proteins (i.e. it is not as prone to saturation). Assuming that the results shown in Figure 3.7, panel D, are a more accurate description of the effects of a p n p deletion on degradosome composition, then there appears to be a "gradient" of RNase E-enolase interactions. In the absence of any other proteins, RNase E and enolase can interact, albeit poorly, as shown in the in vitro assays (Figure 3.6). PNPase can facilitate the interaction between RNase E and enolase, perhaps by binding to the C-terminal tail of RNase E and maintaining this and adjacent regions in a proper conformation. PNPase helps enolase bind RNase E, but is not sufficient for optimal binding. A second factor, perhaps a chaperone, is required to achieve an RNase E-enolase association that is comparable to the in vivo situation. Perhaps this chaperone is needed to fold either of these proteins into a conformation amenable to interaction. 126 A reciprocal experiment consisting of deleting the portion of RNase E involved in PNPase binding and determining whether enolase still associates with the complex, has already been done (Vanzo et al., 1998; Kuhnel & Luisi, 2001). In these experiments, PNPase is still present in the cell but is not associated with the degradosome. The results show that enolase interacts with RNase E in the absence of PNPase in the degradosome. These experiments were performed with overexpressed proteins, which may favour an interaction between RNase E and enolase. The amount of protein present in the complex was not quantified for comparison with the amount found using full-length RNase E, but a qualitative analysis of the evidence suggests that there is relatively less enolase in the degradosome in the absence of PNPase. This confirms the results presented above. 3.3.3 Possible role of enolase in "internal entry" Based on the experiments described in this chapter, it appears that if enolase is involved in modulating the activity of other degradosome proteins, it does not do so in any obvious way. Its presence is not enough to cause a change in the activity of these proteins, nor is its activity sufficient. Glycolytic ligands may cause a mild effect on the activity of RNase E and PNPase, although these results need to be repeated and studied further before conclusions can be made. Preliminary studies of the effects of enolase phosphorylation also failed to detect changes in the activity of the degradosome. 127 All these data suggest that enolase may not be involved in transducing a signal from the cell to link metabolism to RNA decay. The region of RNase E involved in binding enolase seems important for the degradation of circularized RNA in vivo (G.A. Mackie, personal communication). These RNAs lack a 5'-end, are 6-fold more stable than their corresponding linear RNAs, yet can be degraded slowly by RNase E (Mackie, 1998, 2000; Baker & Mackie, 2003). Whether it is the presence of enolase or the region of RNase E involved in the interaction with enolase that is required for this function is currently the focus of experimentations. If enolase facilitates the degradation of circularized RNA, a re-investigation of its RNA-binding ability may be in order. After all, the region of enolase that binds the phosphate of 2-phosphoglycerate could have the ability to interact with the phosphate backbone of RNA (Kuhnel & Luisi, 2001). Conversely, if the region of RNase E that binds enolase is involved in processing circular RNA, it will be interesting to find out whether enolase is required to fold RNase E into the appropriate conformation. These data are supported by the finding that the 693-845 region of RNase E, which overlaps with the enolase binding site, is important for RNA binding, and that deletion of this region decreased the activity of RNase E toward substrates whose degradation is known to be 5'-end-independent (Leroy et al., 2002). These studies also did not differentiate whether enolase or the region of RNase E binding enolase is necessary for these effects. These data are not the only ones supporting the existence of an "internal entry mechanism" in the C-terminal 128 region of RNase E (Jiang et al., 2000; Marchand et al., 2001), but they are the first ones to pinpoint the region involved. 129 CHAPTER IV CHARACTERIZATION of a "COLD SHOCK" DEGRADOSOME 4.1 BACKGROUND E. coli is an enterobacterium. Its lifestyle entails periodic expulsion from the thermally regulated intestinal environment and sudden exposure to cold (Lahti et al., 2003; Burton et al., 1987). Like all other organisms, it is capable of mounting a response to this stress and adapt to the new conditions (Herendeen et al., 1979). There is no precise temperature below which a response is initiated; rather, the temperature difference determines the magnitude of the response. In the laboratory, studies of cold shock have been standardized as a downshift from 37°C to 15°C. Understanding the cold shock response will undoubtedly lead to new techniques to avoid food spoilage (Ansay et al., 1999; Bollman et al., 2001; Janes et al., 2002), help develop cold-inducible systems for the expression of heat- and proteolysis-sensitive enzymes (Vasina et al., 1998; Mujacic et al., 1999), help combat plant bacterial pathogens whose virulence is triggered by cold (Konkel & Tilly, 2000; Smirnova et al., 2001; Felix & Boiler, 2003), protect crops from the cold (Sun et al., 1995), and provide insights into stress adaptation and changes in gene regulatory networks (Weber & Marahiel, 2003). 130 4.1.1 Overview of effects of cold shock Temperature is one of the most influential parameters for life. It determines the rate at which all chemical reactions take place, and directly influences the structure of cellular components (Eriksson et al., 2002). The effects of increases in temperature on the cell have been studied extensively. The heat shock response is under the central control of the RNA polymerase alternative sigma factor a 3 2, which directs the expression of a set of chaperones and proteases that help refold or remove heat-denatured proteins (reviewed in Bukau, 1993; Arsene et al., 2000). Cold shock does not rely on a similar central regulator, although a set of proteins is expressed. To adapt to growth at temperatures above 8°C following a temperature downshift, E. coli induces a set of 26 cold shock proteins (Jones et al., 1987; Gualerzi et al., 2003; Table 4.1). Below this temperature, E. coli cannot grow, although the expression of 69 proteins has been reported in the few hours following a shift to 4°C (Das & Goldstein, 1968; Friedman etal., 1969, 1971; Broeze etal., 1978; Perrot etal., 2000). One of the problems encountered by cold shocked cells is the rigidification of membranes, which can affect all membrane-coupled processes such as transport, energy generation, and cell division (Janoff etal., 1979; Eriksson etal., 2002; Denich et al., 2003). To compensate for the loss of fluidity, cells must incorporate fatty acids of lower melting point (desaturated fatty acids, or with 131 Protein Description Reference CsdA DEAD-box RNA helicase Jones etal., 1996 CspA Major cold shock protein; RNA chaperone, Trancriptional activator and antiterminator Goldstein etal., 1990; LaTeana etal., 1991 j CspB CspA homologue Jones & Inouye, 1994 CspE CspA homologue Phadtare & Inouye, 1999 CspG CspA homologue Nakashima etal., 1996 Cspl CspA homologue Wang etal., 1999 DnaA Initiation of DNA replication Atlung & Hansen, 1999 GyrA Topoisomerase Jones et al., 1992 H-NS Histone-like nucleoid protein La Taena etal., 1991 Hsc66 DnaK homologue Lelivelt & Kawula, 1995 HscB DnaJ homologue Lelivelt & Kawula, 1995 HU(3 Histone-like protein Giangrossi etal., 2002 IF1 Translation initiation factor Calogero etal., 1987 IF2 Translation initiation factor Jones & Inouye, 1994 IF3 Translation initiation factor Gualerzi etal., 2003 NusA Termination and anti-termination of transcription Jones & Inouye, 1994 OtsA Trehalose phosphate synthase Kandror etal., 2002 OtsB Trehalose phosphatase Kandror etal., 2002 PNPase 3'->5' exonuclease Jones & Inouye, 1994 ProX Subunit of ProVXW transporter Polissi etal., 2003 Pyruvate dehydrogenase E1 Converts pyruvate to acetyl-CoA and C 0 2 Jones & Inouye, 1994 Pyruvate dehydrogenase E2 Converts pyruvate to acetyl-CoA and C 0 2 Jones & Inouye, 1994 RbfA Ribosome assembly Jones & Inouye, 1996 RecA Homologous recombination Jones & Inouye, 1994 RhIE Putative DEAD-box helicase Polissi etal., 2003 RNase R Hydrolytic 3' 5' exonuclease Cairrao etal., 2003 RpoE a E extracytoplasmic stress response regulator Polissi etal., 2003 RseA Negative regulator of RpoE Polissi ef al., 2003 " | Trigger Factor Protein chaperone Kandror & Goldberg, 1997 | Ves Unknown Yamada etal., 2002 | PY Translation inhibitor Agafonov ef al., 2001 f Table 4.1 List of all currently known E. coli proteins expressed in response to cold shock. Adapted from Gualerzi etal. (2003). 132 more branching or shorter chain lengths) into their membranes, a process called the homeoviscous adaptation (Sinensky, 1974; reviewed in Suutari & Laakso, 1992; Sajbidor, 1997). A reduction in thermal energy also affects proteins by impeding their folding and preventing them from achieving a proper conformation. Energy barriers for protein folding become difficult to overcome, and folding may occur too slowly to yield an active protein. Even peptides that fold to their native structure may not be functional if their conformational flexibility is reduced. With time, they may even denature due to the reduction in the stabilizing influence of hydrophobic interactions as temperature decreases (as temperature decreases, there is less energy available to remove water from around hydrophobic groups in contact with solvent) (Salotra et al., 1995). All of these problems can be partly remedied with the help of protein chaperones, which are induced by cold shock (Table 4.1; trigger factor, IF2 and Hsc66) (Lelivelt & Kawula, 1995; Kandror & Goldberg, 1997; Caldas et al., 2000). In fact, the overexpression of the chaperones DnaK and GroEL makes E. coli freeze-tolerant (Chow & Tung, 1998). While these proteins are not normally expressed during cold shock in this bacterium, they are naturally induced by cold in Leuconostoc esenteroides (Salotra et al., 1995). In addition to chaperones, low molecular weight compounds such as polyols, polyamines, sugars (most notably the disaccharide trehalose), and amino acid derivatives are accumulated in the cytosol during cold shock to serve as cryoprotectants in the event of freezing (Kandror etal., 2002). Increased concentrations of these solutes results 133 in increased viscosity and lowered freezing point, thereby protecting proteins from the cold (Weber & Marahiel, 2003). Twenty of the 31 proteins known to be induced by cold shock (Table 4.1; Gualerzi et al., 2003) are involved in interactions with, or modifications of nucleic acids. This probably reflects the fact that DNA and RNA conformations are greatly affected by lowered temperatures. Their secondary structures are stabilized, making replication, transcription, translation, ribosome assembly, and degradation more difficult than under previous conditions. Among these are five members of the "cold shock protein family" (CspA, CspB, CspE, CspG, Cspl), a conserved group of small proteins whose roles include transcription activation, RNA chaperone, translation factor, and transcription antitermination (Goldstein et al., 1990; La Teana et al., 1991; Jones & Inouye, 1994; Nakashima etal., 1996; Jiang etal., 1997; Phadtare & Inouye, 1999; Wang etal., 1999; Bae etal., 2000; Giuliodori et al., 2004). CspA is the major protein induced by cold shock. Its production accounts for 13% of protein expression during the first two hours of cold shock (Goldstein et al., 1990). Also induced are the cx-subunit of DNA gyrase (not surprising given the increase in DNA negative supercoils after cold shock (Grau et al., 1994; Jones et al., 1992b; Goldstein & Drlica, 1984; Mizushima et al., 1997)), HU(3 (one of the two subunits of the nucleoid-associated protein HU), the nucleoid-associated DNA-binding protein H-NS, the recombination factor RecA, the replication protein DnaA, the translation initiation factors IF1, IF2 and IF3, the ribosome inhibitor pY, the transcription 134 terminator/antiterminator NusA, the RNA helicases RhIE, CsdA and RbfA, and the exonucleases RNase R and PNPase. 4.1.2 Stages of cold shock The cellular response to a sudden drop in temperature occurs in two stages. Immediately upon cold shock, translation is stopped, and most mRNAs are stabilized (Friedman et al., 1971; Wice & Kennell, 1974; Broeze et al., 1978; Goldenberg etal., 1996; Beran & Simons, 2001; Mathy etal., 2001; Zangrossi et al., 2000; Yamanaka & Inouye, 2001). This latter fact is known to be crucial in the expression of a set of cold shock proteins which are induced despite the general block in protein synthesis (Brandi et al., 1996; Goldenberg etal., 1996; Fang et al., 1997). The Shine-Dalgarno and translation initiation sites are insufficient for translation initiation in the cold, but the mRNAs for some of the cold shock proteins bypass this problem by encoding a sequence called the "downstream box" in their coding region that interacts with ribosomes and allows translation in the cold (Mitta ef al., 1997; Fang et al., 1998; Xia et al., 2002). These proteins accumulate, and are presumed to adapt the cell (and particularly the ribosome) for growth at low temperature. This is called the acclimatization or acclimation phase, and usually lasts about 2 hours following a temperature downshift. After this time, the cell resumes translation of all proteins, albeit at a slower rate (Goldstein ef al., 1990). This recovery or cold-adapted phase can last an indeterminate amount of time (Ng ef al., 1962; Jones et al., 1987; Herendeen et al., 1979; Ingraham & Marr, 1996). The mRNAs encoding the cold 135 shock proteins that helped acclimatize the cell for growth in the cold are specifically targeted for degradation, and the levels of these proteins reach new lower steady states (Goldenberg et al., 1996, Zangrossi et al., 2000; Beran & Simons, 2001; Yamanaka & Inouye, 2001; Figure 4.1). If this selective degradation of cold shock protein mRNA does not occur, neither does growth resumption. Presumably, cold shock mRNAs trap ribosomes and prevent the translation of other mRNAs (Jiang etal., 1996a; Neuhaus etal., 2000). 4.1.3 mRNA decay during cold shock PNPase is induced upon exposure to cold in E.coli, Yersinia enterocolitica and Photorhabdus sp. (Jones etal., 1987, 1996; Clarke & Dowds, 1994; Goverde et al., 1998; Zangrossi et al., 2000; Beran & Simons, 2001; Mathy et al., 2001; Polissi et al., 2003). It reaches its maximal expression level, which is 2- to 8-fold greater than at 37°C, approximately 3-5 hrs after temperature downshift (Herendeen et al., 1979; Jones et al., 1987; Zangrossi et al., 2000; Beran & Simons, 2001; Cairrao etal., 2003). The protein remains active in the cold (Raue & Cashel, 1974; Zangrossi et al., 2000; Beran & Simons, 2001; Polissi et al., 2003) and is important for growth at low temperatures (Luttinger et al., 1996; Wang & Bechhofer, 1996; Goverde et al., 1998; Beran & Simons, 2001). It has been suggested that the role of PNPase at cold shock is linked to its S1 domain, since the overexpression of this region suppresses the cold-sensitive phenotype of a cspA, cspB, cspE, and cspG quadruple mutant (Xia et al., 2001). However, the S1 domain bears strong structural similarity to the "cold shock domain" of 136 Log Accl imatizat ion Recovery P h a s e Phase P h a s e (37°) (0-2 hrs of cold shock) (2+ hrs cold shock) Figure 4.1 Rate of bulk and cold shock protein synthesis following temperature downshift The rate of bulk protein synthesis remains steady during logarithmic growth at 37°C (-). However, following a temperature downshift to 15°C, protein synthesis stops. This inhibition lasts for approximately 2 hrs, then growth resumes. This pattern of protein expression is not observed for cold shock proteins (---). Cold shock proteins are poorly expressed at 37°C. However, they are dramatically induced by exposure to the cold. The rate of their synthesis peaks during the acclimation phase and then drops to a new steady state level for as long as the cell remains at 15°C. Adapted from Thieringer etal. (1998). Copyright© 1998 Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc. Figure reproduced by permission of John Wiley & Sons, Inc. 137 these proteins (Bycroft et al., 1997; Graumann & Marahiel, 1998), and it is likely that this compensatory role is an artifact of high copy suppression. Rather, the exonucleolytic activity of PNPase is likely the essential function at cold shock. Mutants with a defective pnp gene respond to cold shock with the initial induction of cold shock proteins, as do wild-type cells (Yamanaka & Inouye, 2001). However, these proteins are not repressed after the acclimation phase, and there is no growth resumption (Yamanaka & Inouye, 2001). It is believed that PNPase is required for the degradation of mRNAs encoding cold shock proteins 2 hours after a temperature downshift (Neuhaus et al., 2000; Beran & Simons, 2001; Yamanaka & Inouye, 2001; Polissi et al., 2003). Two exceptions are the mRNAs encoding CsdA and RbfA, which seem immune to its activity. In addition, mRNAs encoding non cold shock proteins (e.g. enolase mRNA) are unaffected by the presence or absence of PNPase in the hours following cold shock (Yamanaka & Inouye, 2001). Thus, PNPase is required for the transient expression of most cold shock proteins. PNPase's ability to specifically degrade cold shock mRNAs is increased by the activity of PAPI and CsdA (Yamanaka & Inouye, 2001). However, PNPase can carry out this function in the absence of degradosome formation (Yamanaka & Inouye, 2001). Repression of cold shock protein expression in the recovery phase and resumption of cell growth are indistinguishable between a wild-type strain and one where the scaffolding region of RNase E is missing (smbB105) (Yamanaka & Inouye, 2001). While degradosome formation might be 138 dispensable for PNPase's role at cold shock, the activity of RNase E is required for viability at 15°C. Cells missing the distal part of the catalytic domain, the ARRBD, and the scaffold of RNase E (rneA610) are not viable, while rneA508 lacking the ARRBD and platform is viable at 15°C. Both strains can grow at 37° (Beran et al., 2003). Some changes clearly occur in the mRNA decay apparatus during cold shock. During acclimation, the apparatus becomes unable to degrade most mRNAs in the cell. Then after cold adaptation, it specially targets cold shock transcripts. The cellular concentration of RNase E is maintained following cold shock and remains unchanged throughout cold adaptation (Cairrao et al., 2003). While cold shock does not appear to modulate the concentration of degradosomes, the degradation complex could be adapted to the cold shock conditions by other means. Temperature has been demonstrated to influence the structure of RNAs, and cold shock in particular stabilizes RNA structures (Storz, 1999). These structures could conceivably impede the progress of PNPase, particularly if RhIB is unable to dissociate the two strands of stabilized hairpin structures. CsdA, an alternative DEAD-box helicase, is expressed at cold shock. This protein may replace RhIB in the degradosome, or else interact directly with PNPase to form a novel complex capable of overcoming cold-stabilized RNA structures. 139 4.1.4 CsdA The gene encoding cold shock DEAD-box protein A (CsdA) is located on a portion of the E. coli chromosome rich in cold shock proteins (linked genes include nusA, infB [encoding IF2B], pnp, and rbfA) (Toone et al., 1991). Mutant strains exhibit a reduced growth rate and filamentous growth at low temperature (lost & Dreyfus, personal communication; Jones et al., 1996; Yamanaka & Inouye, 2001). CsdA (a.k.a. DeaD/mssB/RhID) is a non-essential protein induced immediately upon cold shock, even prior to the expression of CspA (Kalman etal., 1991; Toone et al., 1991; Yamanaka etal., 1994b; Jones et al., 1996; Goldenberg et al., 1997). Its expression peaks 2 hrs after a temperature downshift, and then reaches a new lower steady state level. As mentioned earlier, it is a member of the DEAD-box family of RNA helicases and has been shown to possess RNA unwinding activity in vitro (Jones etal., 1996). Studies of CsdA's role in the cell can be grouped into two categories. One group reports that CsdA is associated with ribosomes (Jones et al., 1996) and is involved in either ribosome biogenesis or translation, while the other observes an effect of this protein on mRNA decay. Whether these two observations are different effects of one function or reflect two separate roles of CsdA in the cell remains to be established. Evidence that CsdA is linked to ribosomes and translation comes from several lines of evidence. Jones and colleagues (1996) recovered CsdA exclusively in the ribosome fraction following ultracentrifugation. This association 140 could be disrupted with 1 M salt, suggesting that ionic interactions mediated the interaction. However, more recently, Moll and colleagues (2002) were unable to reproduce these findings. Nonetheless, CsdA overexpression can suppress a temperature sensitive mutation in rpsB (encoding ribosomal protein S2) (Toone etal., 1991). The function of S2 is normally to recruit the ribosomal protein S1 to the ribosome (Laughrea & Moore, 1978). S1 has an RNA unwinding ability, recruits mRNA to the 30S ribosomal subunit (possibly by destabilizing mRNA secondary structures that impede such a process), and can influence the affinity of ribosomes for different mRNA translation initiation sequences (Szer et al., 1976; Steitz et al., 1977; Subramanian, 1983; de Smit and van Duin, 1994). While it might be tempting to conclude that CsdA can replace the RNA unwinding activity of S1 in the ribosome, CsdA appears to rescue rspB mutations by recruiting ribosomal protein S1 to the ribosome, even in the absence of S2 (Moll et al., 2002). CsdA may promote unfolding/refolding of 16S rRNA and allow mutant S2, and as a consequence S1 to associate with the ribosome (Moll et al., 2002). Thus, in the cold, 16S rRNA may be trapped in conformations that do not allow for the incorporation of S2 and S1, leading to translational defects. CsdA's role could be to remodel the 16S structure, thereby allowing the assembly of S2 and S1 into the ribosome, and restoring ribosome function. Indeed, upon cold shock, leaderless mRNAs (lacking 5' UTR and Shine-Dalgarno sequences), whose translation is not dependent upon the presence of either S1 or S2 in the ribosome, are preferentially translated by the cell (Grill et al., 2002). Lu and colleagues (1999) have found that CsdA is necessary for the in vitro translation 141 of structured mRNAs at 37°C. It is not required in the translation of mRNAs that lack this feature. This is not surprising if CsdA recruits S1 to the ribosome, given that S1 can remodel mRNAs to make them more amenable to translation. It is difficult to reconcile the seemingly opposite effects of CsdA on mRNA stability. One the one hand, CsdA and PAPI help PNPase repress the expression of cold shock mRNA after the acclimation phase to allow the resumption of growth (Yamanaka & Inouye, 2001). CsdA interacts with PAPI, which is capable of associating with RNase E (Raynal & Carpousis, 1999). These results suggest that CsdA facilitates degradation. However, overexpression of CsdA at 37°C leads to the stabilization of the cspA mRNA at that temperature (Brandi et al., 1999). In addition, mRNAs induced from a T7 RNA polymerase promoter are stabilized by the overexpression of a number of DEAD-box helicases, including CsdA (lost & Dreyfus, 1994). Such transcripts are usually very unstable, since the T7 RNA polymerase outpaces ribosomes and leads to the creation of "naked" (i.e. unprotected by ribosomes) mRNAs. Overexpression of CsdA does not affect the expression of proteins from mRNAs transcribed by the E. coli RNA polymerase, except for those that lack a functional Shine-Dalgarno sequence. Given the possible involvement of CsdA with the ribosome, it is tempting to suggest that CsdA facilitates translation of transcripts where such a process is slow, thereby protecting the mRNA. However, removal of the ribosome binding site on T7 transcripts did not prevent the stabilizing effect of CsdA (it should be noted, however, that their translation was not monitored, 142 and CsdA may be able to bypass the requirement for a ribosome binding site). Removal of the H-R-l-G-R-X-X-R sequence of CsdA (motif VI in Figure 4.2), implicated in NTP binding and hydrolysis or alternatively in RNA interaction in DEAD-box proteins8 (Hall & Matson, 1999; Caruthers & McKay, 2002), resulted in loss of the stabilizing effect of CsdA. CsdA has also been isolated as a multicopy suppressor of the cold-sensitive phenotype of the smbA2 mutation (Yamanaka et al., 1994b). The SmbA protein is a member of the aspartokinase family, involved in cell proliferation. Mutations in this protein cause abnormal protein expression, growth defects, cold-sensitivity and abnormalities in cell division. Like RNase E, overexpression of this protein can suppress mukB mutations (Yamanaka et al., 1994a; Kido et al., 1996). CsdA may therefore be able to carry out yet another function that restores growth in the cold when SmbA is absent. Alternatively, smbA maps in close proximity to the rpsB gene. Mutations in smbA may cause a polar effect on S2 expression. The rescue of smbA mutant by CsdA overexpression may, therefore, be due to suppression of a defect in the expression of S2 protein. 4.1.5 DEAD-Box Helicases Helicases are present in nearly all isolated RNA degradation complexes (Section 1.6.11), highlighting their importance in RNA processing and decay. By 8 At the time of publication of the article by lost & Dreyfus (1994), only the RNA-binding activity of this region was recognized. 143 analogy to the AAA ATPase of the proteosome which unfolds proteins and delivers them to the degradative enzymes, helicases could unwind RNA and deliver them to ribonucleases (van Hoof & Parker, 1999). Helicases can be divided into 4 groups (3 superfamilies, and 1 family) based on sequence analysis (Gorbalenya & Koonin, 1993; Hall & Matson, 1999). Superfamily 2 (SF2) is the largest group, and includes DNA helicases, RNA helicases, and DNA- or RNA-stimulated ATPases. All of the conserved motifs of the proteins belonging to this superfamily appear to be required for activity, and the three dimensional structures of these helicases are well conserved. The DEAD, DEAH, and DExH proteins are members of SF2. They share a core of 10 conserved sequence motifs. Their name reflects divergence of the amino acids present in motif II (also known as Walker B motif) (e.g. DEAD-box contain the sequence "aspartate (D) - glutamate (E) - alanine (A) - aspartate (D)" (Figure 4.2)). They are found in organisms from all kingdoms, and play important roles in RNA metabolism and processing, such as transcription, splicing, export, degradation, ribosome assembly, translation, and editing (Schmid & Under, 1992). Some members have also exhibited the ability to disrupt RNA-protein interactions (Jankowsky et al., 2001), and at least one DExH-box protein (NPH-II) is a demonstrated processive and directional helicase (Jankowsky etal., 2000). RhIB and CsdA are members of the DEAD-box protein family. There are at least five DEAD-box proteins in E. coli (Kalman et al., 1991). RNA helicase-Jike protein A (RhIA), also called SrmB, is a suppressor of 144 ribosomal maturation defects of rplX (L24) mutants (Nishi et al., 1988), and is involved in an early step of the assembly of 50S ribosomal subunits (Charollais et al., 2003). Deletion of its gene causes cells to grow at a slower rate at low temperature (Charollais et al., 2003). SrmB possesses RNA-dependent ATPase activity, which can be stimulated over 20-fold by the presence of PAPI (Nishi et al., 1988; Raynal & Carpousis, 1999). A physical association between SrmB and PAPI has been demonstrated by far-Western analysis (Raynal & Carpousis, 1999). SrmB has the ability to stimulate the polymerization activity of PAPI approximately 1.5-fold (Raynal & Carpousis, 1999). RhIB, the second DEAD-box helicase, has been described extensively in Section 1.6.4. It is a component of the RNA degradosome, and has been shown to have both ATP-dependent RNA helicase and RNA-dependent ATPase activity in vitro. RhIC, also known as DbpA, is an RNA helicase acting on 23S rRNA substrate (Iggo etal., 1990; Diges & Uhlenbeck, 2001; Henn et al., 2001). Interestingly, it adopts different conformations when it binds ATP, ADP, or RNA (Henn etal., 2002). RhIE has an ATPase activity that can be stimulated 2.5-fold by the incorporation of PAPI in the assay (Raynal & Carpousis, 1999). It has also been shown by far-Western analysis to interact with PAPI (Raynal & Carpousis, 1999). RhIE knock-outs are viable in E. coli (Ohmori, 1994), and it has recently been reported that RhIE is induced by cold shock (Polissi et al., 2003). Finally, CsdA is the only known cold shock-inducible member of this family in E. coli, although DEAD-box proteins are induced by cold in many organisms, from cyanobacteria to archae (Chamot etal., 1999; Lim etal., 2000; Yu & Owttrim, 2000; El-Fahmawi & Owttrim, 2003). An 145 Figure 4.2 Alignment of CsdA and RhIB amino acid sequences The amino acid sequences of CsdA and RhIB were aligned using ClustalW. Identical residues are highlighted with a black background, whereas conserved substitutions are shown over a grey backdrop. The two sequences are 38% identical (54% similar). Shaded boxes identify motifs common to all DEAD-box proteins, along with their putative functions. The conserved sequences corresponding to these motifs are shown under the alignment (Under et al., 1989; Gorbalenya & Koonin, 1993; de la Cruz et al., 1999; Hall & Matson, 1999; Jankowsky & Jankowsky, 2000; Tanner & Under, 2001; Tanner et al., 2003). 1 4 6 CsdA RhIB MOTIF 1 (Walker A Motif: P-loop) Binding t r i - P 0 4 moitie of ATP Q Motif Regulates ATP binding and hydrolysis 20 40 —AEF-HTT: SKTHLT THAff lLGf lKAPIL^f f lNDLff iYEKPS^^EClSHLBN K § S g F A j J H P K V V ^ 2 E K K I F H N C T ^ ® L A I ' a L T B A EKPS HNC1 GF PT IQ A TGTGKT AF 59 65 CsdA RhIB MOTIF la Nucleic acid interaction through s u g a r - P 0 4 backbone S:.?:,BQN LBPELKABGIHVL T F H Y § L S H P A I A | R K V N Q | R A | I M 1 00 AEAMTDFSKHM IHADAEPLAEAT ILAPTRELA Q" MOTIF lb 1 20 L^KLGLAB8JDGB8 GG 1 20 1 29 CsdA RhIB MOTIF Ic 1 4 0 fi] H\ ^ P G R L MOTIF II (Walker B Motiti Binds Mg + 2 to bind ATP 160 GTLDHSKLSGL IQNHINBGAIQ IEBLRMI VLDEAD M 180 JEfflVETIMAQlH JKflRWLFRRMg EGH PA - T A LNM 183 1 94 MOTIF III Transmission of conformational change upon ATP hydrolysis —um 2 20 CsdA : E M M P E A I 3RITRRF| RhIB : ( J j L • 1 gELAFEQj FSAT MOTIF IV ssRNA binding 240 rfepDHsQSYWTWGMRBNEAHvRFLHAHD SJGHR|KEELFYPSNEE|MRLJQTLI|E§W 260 IDFS PI 280 CsdA : j&/P^NATLSvAEAHERI^YNSAAHN^MKi.i.LHEQ'TfflE RhIB : ^ A N ^ H R C E g l W G H g A A D f H R V G L g T ^ V a j K K g L R i g D I F T 300 RLKDl EFT: MOTIF V Nucleic acid binding 320 VERI IPAV LVATDVAARGLD 2 4 8 2 5 9 31 3 3 2 4 CsdA RhIB MOTIF VI NTP binding/hydrolysis 5TE3 360 AjJLFVEN SfflHSISLAC 380 Y HRIGRTGR G R S R R L L R N I E R T M K H T I B E V & E S Y A-JNL|§AI I L P N A E L L T Y| 378 374 CsdA RhIB CsdA RhIB CsdA RhIB 400 RRLEKFAAKVQQQLESSDLDQYRAL: HSIP 460 JEFSE SLiaLTS 440 |TAEGEEL|LETLAAALLKMAQGERTLIVPP|G : 443 ALM --T|L : 392 LHIQIT II V ^ Y N g 1-48( RH G^DSEDRJ p . E l ! 500 520 MQLYRIEVGRDDGVEVRHIVGAIA 508 420 540 560 580 NEGDISSRYIGNIKLFASHSTIELPKGMPGEVLQHFTRTRILNKPMNMQLLGDAQPHTGGERRGG : 573 600 620 640 C s d A : GRGFGGERREGGRNFSGERREGGRGDGRRFSGERREGRAPRRDDSTGRRRFGGDA RhIB : 628 1 4 7 alignment of RhIB and CsdA, along with the conserved DEAD-box motifs is shown in Figure 4.2. 4.2 RESULTS The experiments described in this chapter sought to answer three questions: • Are degradosomes assembled in cold-adapted cells? • Is CsdA part of a cold shock degradosome? • Can CsdA functionally replace RhIB in the degradosome? 4.2.1 Time course of CsdA induction during cold shock The literature reports the rapid induction and accumulation of CsdA (up to 10-fold) upon cold shock (Jones etal., 1996; Goldenberg et al., 1997; Thieringer et al., 1998). However, some investigators find CsdA in 37°C cultures (Rudolf K. Beran, personal communication). To verify that CsdA is indeed induced by a temperature downshift, BL21 (DE3) cultures were grown at 37°C. An aliquot was removed and the cells immediately lyzed in 2X sample buffer, avoiding freezing or cold room manipulations. The remainder of the culture was placed at 15°C and grown at this temperature for an additional 240 minutes. Aliquots were periodically removed and prepared as for the 37°C sample. The. amount of cells taken at each time point was normalized by monitoring the ODeoo of the culture. A typical time course of CsdA accumulation is shown in Figure 4.3. Cell extracts were electrophoretically separated on a gel and the corresponding Western blot 148 37°C 15°C 1 min 5 min 15 min 30 min 1 hr 2 hrs 4 hrs Figure 4.3 Time course of CsdA induction during cold shock 15 ml of BL2KDE3) cells were grown at 37°C in LB broth to mid-log phase (OD 6 0 0 = 0.4). Cells present in 1 ml of culture were harvested by centrifugation, and re-suspended in 150 ul of 2X sample buffer. Meanwhile the remainder of the culture was shifted to 15°C, and growth monitored using OD 6 0 0 . Aliquots containing an amount of cells comparable to the initial 1 ml aliquot were removed at the indicated time points, and subjected to the same treatment as the 37°C aliquot. 10 ul of these preparations were loaded on an SDS-PAGE gel. The picture shown is a Western blot of this gel, probed with anti-CsdA antibodies. 1 4 9 probed with anti-CsdA antibodies. CsdA is indeed present in 37°C cells. However, it is rapidly induced upon cold shock, and appears to reach a new steady state level within one hour of the temperature downshift. The CsdA band 4 hrs after cold shock is at least three times more intense than at 37°C. These results are in general agreement with previous reports (Jones et al., 1996; Goldenberg etal., 1997; Thieringer et al., 1998). 4.2.2 Existence and composition of the cold shock degradosome The immediate stabilization of bulk mRNA upon cold shock could be due to the inhibition of the degradosome, its disassembly, or to the proteolytic degradation of degradosome proteins. Are degradosomes present and/or assembled in cold shocked cells? If so, are these different from those found at 37°C? The re-initiation of mRNA degradation two hours after cold shock, with modified specificity, raises the possibility that the mRNA decay apparatus is altered during cold adaptation. Biochemical purification of cold shock degradosomes Cultures of CF881 E. coli were grown overnight at 37°C or for three days at 15°C in LB broth. Cell from both cultures were lyzed and subjected to identical protocols for the biochemical purification of degradosomes (Section This protocol involves obtaining a cytosolic extract, precipitating the large complexes contained therein with ammonium sulfate, subjecting the precipitated proteins to ion-exchange and size exclusion chromatography. The proteins thus 150 purified from the two cell cultures were separated on an SDS-PAGE gel, and visualized by staining with Coomassie Blue (panel A of Figure 4.4). Degradosomes were purified from cells grown at either temperature, confirming the existence of degradosomes in cold-adapted cells. The protein profile from the 37°C preparation is identical to published profiles (Figure 4.4, panel A, lane 2; Py et al., 1996; Coburn & Mackie, 1998). However, complexes isolated from cold-adapted cells contain additional bands (Figure 4.4, panel A, lane 3). Each of the bands from the cold shock degradosome (Figure 4.4, panel A, lane 3) was carefully excised and sent for identification by mass spectrometry at the University of Victoria's Genome BC Proteomics Centre. All bands were unambiguously identified9 (Table 4.2), and are labeled in panel A of Figure 4.4. In addition to the expected 37°C degradosome proteins (RNase E, PNPase, RhIB, and enolase), the cold shock complex co-purifies with CsdA, the a and |3 and/or |3' subunits of RNA polymerase, and unexpectedly, the E1 and E2 subunits of the pyruvate dehydrogenase complex. Both degradosome preparations were reacted with anti-Rne and anti-Pnp antibodies, followed by a secondary incubation with anti-rabbit gold-conjugated antibodies. The complexes formed were negatively stained with uranyl acetate and viewed by transmission electron microscopy (Section 2.8). No complexes 9 With the possible exception of the (3 or |3' subunit of RNA polymerase. These two peptides could not be distinguished, so the identity of this band is either the (3 subunit, or the |3' subunit, or a combination of both. 151 A kDa 4 & _ J RNase E PNPase -RNA pol (3 and/or (3' Pyruvate "dehydrogenase (E1) Pyruvate "dehydrogenase (E2) CsdA RhIB -enolase -RNA pol a 100 nm Figure 4 . 4 Degradosomes isolated from cells grown at 3 7 ° C or 1 5 ° C A. Degradosomes were purified from E. coli cells using the series of biochemical steps outlined in Section Bacteria were either grown overnight at 37°C or for 3 days at 15°C. 3.6 ug of the proteins obtained at the end of the purification protocol were loaded in each lane and separated by SDS-PAGE. Each protein band was unambiguously identified by mass spectrometry (Section 2.6.6). Lane 1) protein standards, lane 2) 37°C degradosomes, lane 3) 15°C degradosomes. B. Transmission electron micrograph of one of the bodies found in the cold shock degradosome preparation. These structures were absent in the protein sample isolated from 37°C-grown cells. 152 Table 4.2 Identification of cold shock degradosome proteins by mass spectrometry Protein RNase E RNA polymerase |3' RNA polymerase (3 Pyruvate dehydrogenase (E1) PNPase Pyruvate dehydrogenase (E2) CsdA RhIB Enolase RNA polymerase a Number of peptides positively identified by mass spectrometry 27 32 14 32 17 16 21 14 12 8 153 could be seen in the 37°C preparation. However, the 15°C preparation contained bodies shown in panel B of Figure 4.4. These bodies are reminiscent of the structure and size (- 45 nm) of pyruvate dehydrogenase complexes (Reed et al., 1964; Cajacob et al., 1985; Wagenknecht et al., 1990, 1991). No gold particles were found in the vicinity of these bodies, suggesting that pyruvate dehydrogenase complexes and degradosomes are not physically associated. Immunopurification of cold shock degradosomes To confirm the presence of new and additional components in cold shock degradosomes, immunological "pull-downs" were performed on cold shocked cells. This technique should not co-purify the RNA polymerase and pyruvate dehydrogenase complexes if they do not physically associate with the degradosome. Cells capable of overexpressing FLAG-tagged RNase E were grown at 37°C to mid-log phase, and then placed at 15°C for 2 hrs. FLAG-RNase E 1 0 was induced immediately upon temperature downshift. The cells were lyzed and crude extract incubated with commercially available monoclonal anti-FLAG beads (Sigma). This immunological procedure should retain FLAG-RNase E on the beads, along with any associated protein. Since the expression of FLAG-RNase E was initiated at cold shock, any protein found to interact with FLAG-RNase E will have done so at 15°C. Note, however, that due to the general stability of proteins in E. coli (Larrabee et al., 1980), there is no way to ascertain whether these proteins were made before or after the shift. The bound 1 0 The presence of the FLAG tag is known to have little effect on the activity of RNase E, as a plasmid encoding FLAG-Rne can rescue an rne mutant (Miczak etal., 1996). 154 proteins were eluted using excess FLAG epitope. The results of this experiment are shown in Figure 4.5, panel A. Lane 2 shows the proteins isolated with FLAG-RNase E. For comparison, lane 1 shows the biochemically purified degradosome from Section All the main degradosome components (RNase E, PNPase, RhIB, enolase) are present in both samples. The RNA polymerase (3 and/or (3' subunit(s) cannot be detected in lane 2, but the two pyruvate dehydrogenase subunits are observed in both lanes 1 and 2. CsdA cannot be seen in lane 2 of Figure 4.5A. However, Western blotting of this sample clearly shows the presence of CsdA (Figure 4.5, panel B, lane 1). Similar experiments were done with cells maintained at 37°C or heat-shocked at 42°C. The results of these experiments are shown as Western blots in Figure 4.5, panel B, lanes 2 and 3. Interestingly, RhIB appears to be most abundant in 15°C degradosomes, less abundant in 30°C degradosomes, and least abundant in 42°C degradosomes. To determine whether degradosomes could be assembled after cold shock entirely from proteins synthesized in the cold, degradosomes were immunologically purified from cells pulse-labeled with [35S]-methionine during cold shock. Cells were grown at 37°C to early-mid log phase, and then divided into 3 cultures. One was grown at 37°C for an additional 15 min with [35S]-methionine, while the other two were shifted to 15°C. One of these was cultured for 2 hrs at 15°C and then radiolabeled for 30 min. The other was diluted to OD6oo of 0.1, and the cells grown at 15°C for 2 generations (approximately 16 155 Rn RNA pol |3 or |3' C s d A — RhIB— enolase-E o tf) o •a ro CD -a O o UO -a o CD E o CO o 1=1 ro O J CD •a O 0 LO pur LU LU CD CD CO CD hemically ase ers Nas NPa hIB nola sdA hemically cc CL cc CD CJ hemically cc ro -o "O -a "O "0 hemically 6 E CD CD CD CD CD CJ Q < 5 i— i— i— ZJ ZJ ZS Z3 Z3 m ti- 5 CL D_ CL CL Q. anti-RhIB anti-enolase 2 3 4 5 6 7 8 Figure 4.5 Co-lmmunopurification from unlabeled cell extracts A. Cell extracts from FLAG-Rne expressing cells grown at 15°C were subjected to immunoprecipitation using a monoclonal F L A G antibody (Section 2.9.2). Lane 1 shows the proteins isolated in a biochemical purification of cold shock degradosomes, lane 2 is the result of the F L A G pull-down at 15°C, lane 3 contains size markers (200, 116, 97, 66, 45 kDa), and lanes 5, 6, 7, and 8 are purified Rne, Pnp, RhIB, enolase, and CsdA, respectively, shown for size. Proteins were detected by staining with Coomassie Blue. B. Cell extracts from FLAG-Rne expressing cells grown at 15°C, 30°C, and 42°C were subjected to immunoprecipitation using a monoclonal F L A G antibody (Section 2.9.2). A Western blot of the immunopurification is shown. Lane 1 shows the results of the immunopurification of the 15°C cell extract (the same that was shown by Coomassie stain in lane 2 of panel B). Lane 2 is the immunoprecipitation of the 30°C cells, and lane 3 shows the proteins from the 42°C cells that bound the F L A G column. 156 hrs) before the addition of radiolabel in the last 30 min of growth. Cell extracts were prepared and passed over beads conjugated to RNase E or PNPase antibodies (Section 2.9.1) to capture these proteins as well as any other associated with them. The beads were washed several times, and the bound proteins eluted with SDS and heat. These proteins were separated by electrophoresis and detected by Phosphorlmaging. Figure 4.6 shows the results of such immunopurification experiments. In panel A, extracts from a wild-type P90C strain as well as its isogenic csdA derivative were passed through a column packed with anti-Rne beads. The eluted proteins were identified by their position on an SDS-PAGE gel relative to size markers, including purified RNase E, PNPase, and CsdA proteins in adjacent lanes as references. An anti-Pnp immunoprecipitation was also performed, and yielded essentially identical results (not shown). RNase E and PNPase were unambiguously identified in each lane. The band whose electromobility and size match that of purified CsdA co-purified with RNase E and PNPase only in cold-shocked cultures (lanes 1, 2, and 3). More CsdA was present in the co-immunoprecipitate 2 hrs following a temperature downshift (lane 2), although it is still present in the immunoprecipitation from cells grown at 15°C for two generations (lane 3). This band is absent in the csdA deletion strain (lanes 4, 5, and 6), confirming its identity. The reciprocal immunoprecipitation experiment using anti-CsdA beads only purified CsdA (Figure 4.6, panel B). 157 P90C RS9052 37°C 15°C 15°C 37°C 15°C 15°C 2 hrs 16 hrs 2 hrs 16 hrs B P90C RS9052 37°C 15°C 15°C 37°C15°C 15°C 2 hrs 16 hrs 2 hrs 16 hrs — C s d A r Genotype Overexpressed wild-type F L A G -peptide F L A G Rne F L A G pnp::Tn5 csdA:\Tn5 F L A G - F L A G -Rne F L A G Rne Growth temperature 37°C15°C 37°C15°C 37°C 15°C 37°C15°C 37°C15°C 37°C15°C 205- RNase E 1 2 3 4 5 6 7 8 9 10 11 12 Figure 4.6 Co-lmmunoprecipitation from radiolabeled cell extracts A. Wild-type P90C and RS9052 (containing a csdA deletion) were grown at either 37°C, 15°C for 2 hours, or 15°C for 2 generations (approximately 16 hrs). [3 5S]-methionine was incorporated into the growth medium 30 minutes prior to harvesting. The cells were lyzed and incubated with agarose beads conjugated to polyclonal Rne antibodies (Section 2.9.1). The column-bound fraction was released with 2X sample buffer lacking dyes and DTT, and separated on a 7.5% S D S - P A G E gel. The proteins present on this gel were detected by Phosphorlmager technology. B. Experiment as described in A), but the beads used for immunoprecipitation were conjugated to CsdA polyclonal antibodies. C. Wild-type BL21, BL21 pnp::Tn5, or BL21 csdA::Tn5 cells were grown at either 37°C or 15°C, while expressing either a control F L A G epitope or the FLAG-Rne construct. Following pulse-labeling of proteins with [3 5S]-methionine, Rne was immunoprecipitated using anti-FLAG M2 affinity gel (Sigma). Proteins recovered from the IP were separated electrophoretically and exposed to a Phosphorlmager screen. Pnp and CsdA were identified on the basis of their molecular weight, by separating purified Pnp and CsdA proteins alongside these co-IP samples, by the absence of bands in the respective mutants, and by Western blot (not shown). 158 In order to confirm these results, an alternative method of immunologically purifying degradosomes from radiolabeled cells was employed. In this protocol, cells capable of overexpressing FLAG-RNase E are grown at 37°C to log phase, and then shifted to 15°C for 2 hrs or maintained at 37°C for this period. The FLAG-RNase E construct is only induced in the last 30 min of growth, as is radiolabel incorporation. FLAG-RNase E was pulled-down using anti-FLAG affinity resin. An immunoprecipitation was performed using cells that overexpressed the FLAG epitope to control for proteins that bind the column material and/or the FLAG epitope. This experiment was performed using BL21 as the wild-type strain, as well as its isogenic derivatives in pnp or csdA. The results of these immunoprecipitation experiments are shown in Figure 4.6, panel C. As expected, Rne was recovered in lanes 3, 4, 7, 8, 11, and 12. Likewise, Pnp was recovered in lanes 3, 4, 11, and 12. CsdA co-purifies with RNase E in both the wild-type (lane 4) and pnp mutant (lane 8) cold shock strains, but not in the csdA mutant (lane 12). The high background in lanes 3 and 4 probably reflect the fact that there was more protein synthesized in the wild-type than in the mutant strains. In summary, CsdA appears to co-purify with the degradosome at 15°C. Two subunits of the pyruvate dehydrogenase complex may also be associated with the degradosome at that temperature. 159 Co-immunoprecipitation of purified RNase E, PNPase, and CsdA Purified RNase E, PNPase, and RhIB can be reconstituted into a "minimal degradosome" if incubated together in appropriate buffers in vitro (Coburn et al., 1999). Partially purified FLAG-RNase E (an AS26 extract, see Section was prepared from cells overexpressing this construct. 100 Lig total proteins from this extract were mixed with 20 tag of purified His-PNPase and His-CsdA in pnp assay buffer (Section 2.11.2), and incubated at 15°C for 30 min. The formation of complexes between these proteins was monitored by immunoprecipitating FLAG-RNase E using anti-FLAG affinity resin. The bound proteins were visualized by Western blotting (Figure 4.7). Lane 1 confirms the presence of FLAG-RNase E in the AS26 extract. It also shows that PNPase in this extract forms a complex with RNase E, which is not unexpected. Lanes 2 and 3 are controls to verify that neither His-PNPase nor His-CsdA, respectively, can interact with either the column material or the antibodies. Lane 4 shows the result of mixing all three proteins. His-PNPase and His-CsdA co-purify with FLAG-RNase E. This experiment was repeated with lesser amounts of purified proteins. The co-purification of 5 ixg His-CsdA with 100 Lig crude FLAG-RNase E could still be detected, but Vg His-CsdA was insufficient for detection by the technique described above. 4.2.3 Functional interaction between CsdA and PNPase The MalEF RNA contains a stable stem-loop structure at its 3'-end that blocks the activity of PNPase (Newbury et al., 1987a, 1987b; McLaren et al., 160 anti-Rne anti-Pnp anti-CsdA FLAG-Rne (AS26) HIS-Pnp (purified) HIS-CsdA (purified) Figure 4.7 Physical interaction of Rne, Pnp, and CsdA in vitro Partly purified (AS26 fraction) FLAG-Rne and purified His-Pnp and His-CsdA were incubated together, and co-precipitated using anti-FLAG M2 affinity gel (Sigma; Section 2.10.2). The image above is a composite of three Western blots, each probed with the antibody listed on the left. Each lane represents one immunoprecipitation experiment. The proteins mixed together for the IP are indicated above each lane. Lanes 1-3 are controls where each protein was mixed individually with the anti-FLAG beads to test for reactivity. Lane 4 represents the actual co-IP experiment. 161 1991). PNPase can degrade a minor structure (the * hairpin) on its own, but stalls at the base of the more stable repetitive extragenic palindrome (REP)-encoded hairpin (Figure 3.3, panel B). To overcome it, PNPase requires the assistance of RhIB and ATP (Py etal., 1996; Coburn etal., 1999). RhIB uses the energy of ATP hydrolysis to unwind the double-stranded portion of the hairpin, and passes single-stranded RNA to PNPase (Figure 3.3, panel C; Figure 1.3). PNPase and RhIB must both be bound to RNase E in order to successfully reconstitute degradative activity (Coburn etal., 1999). Given the similarity between RhIB and CsdA, CsdA might also assist PNPase in the degradation of structured 3'-ends. Purified RNase E (1 p;g/ml), PNPase (2 Lig/ml), and 1 ixg/ml of one of the helicases (RhIB or His-CsdA) were incubated in pnp assay buffer (Section 2.11.2) with 1.6 nM in w'fro-transcribed radiolabeled MalEF RNA (Section 2.7.3) in the presence or absence of 3 mM ATP. Assays were performed with RhIB as a positive control, and with CsdA to assess whether CsdA can functionally interact with PNPase. These assays were carried out at 30°C and 15°C. Aliquots were removed from the reaction at predetermined times during a 1 hr assay. The intermediates contained therein were separated from each other on a denaturing polyacrylamide gel (Section 2.7.4) and visualized by Phosphorlmaging. Panel A of Figure 4.8 shows gels from representative assays. The accumulation of the BEP-stabilized BNA (RSR) intermediate, which is an indication of helicase (in)activity, was monitored by densitometry. The accumulation of RSR in the 30°C and 15°C assays is plotted 162 30SC 15SC without ATP without ATP min 0 1 2 5 10 20 30 45 60 0 1 2 5 10 20 30 45 60 mm «np 4nm 'jMWl'" 9 > f l l i l i l ••^ Sff VHP r^tftfer *SIP^  t^iitfc j^ jj^ ^ j J J J - ^ W p ^ '•H!^  with ATP with ATP 0 1 2 5 10 20 30 45 60 0 1 2 5 10 20 30 45 60 HP €(l SI V ***** • MalEF RSR 40 Time (minutes) 60 80 20 40 Time (minutes) 60 80 Figure 4.8 Functional equivalence of RhIB and CsdA in vitro A. Purified Rne, Pnp, and CsdA were added to in wfro-transcribed, radiolabeled MalEF RNA in the presence of phosphate, with or without ATP. Assays were performed at 15°C and 30°C. Aliquots were removed at 0, 1, 2, 5, 10, 20, 30, 45, and 60 minutes after the addition of enzymes. RNA intermediates were separated on a 6% denaturing polyacrylamide gel. Bands were visualized using Phosphorlmager technology. B. and C. Quantification of the band corresponding to the RSR intermediate over a 60 minute period, shown as a percentage of the signal of the intact MalEF at the 0 time point. The graph shown in B. is from the 30°C assay (error bars represent standard errors of the mean), while that in C. is from the 15°C assay. Circles represent data collected while using CsdA in the reconstitution assay, in the presence (•) or absence (o) of ATP. Squares show the data collected using RhIB in the assay, in the presence (•) or absence (•) of ATP. 163 in Figure 4.8, panels B and C, respectively. In the absence of ATP, the RSR intermediate accumulates regardless of the helicase used in the assay at both temperatures. The absence of helicase gave identical results (not shown). At 30°C in the presence of ATP, the RSR intermediate first accumulates and then disappears when either helicase is present (Figure 4.8, panel B). At 15°C in the presence of. ATP, the RSR intermediate slowly accumulates to levels substantially lower (and the * intermediate to higher levels) than those seen in the absence of ATP (Figure 4.7, panel C). Due to the slowed kinetics at 15°C, the assay would need to be monitored for longer periods in order to permit an equivalent amount of enzyme turnover to the 30°C assay. Nonetheless, overall, the results of these assays were identical whether RhIB or CsdA was included in the assay. 4.2.4 ATPase activity of CsdA The finding that CsdA possesses an ATP-dependent helicase activity prompted an investigation of its potential RNA-dependent ATPase activity. Purified His-CsdA (750 ng/ml) was incubated with [7 3 2P]-ATP in pnp assay buffer, in the presence and absence of 20 nM in w'fro-transcribed unlabeled rpsT mRNA (Section 2.12). Aliquots were removed over a 1 hr period and spotted onto a PEI-cellulose TLC plate. Cleaved terminal (7) phosphate was separated from [y32P]-ATP with 0.375 M K H 2 P 0 4 (pH 3.5). The plate was exposed to a Phosphorlmager screen and the spots quantified. An example of the raw data is shown in Figure 4.9, panels A (CsdA + no RNA) and B (CsdA + RNA). 164 Figure 4.9 ATPase activity of C s d A and degradosomes Purified His-CsdA (750 ng/ml) was incubated with [ Y 3 2 P ] - A T P in pnp assay buffer at 30°C or 15°C in the presence or absence of 20 nM unlabeled in w'fro-transcribed rpsT mRNA and up to 7.5 Lig/ml purified Rne (Section 2.12). Aliquots were removed at 0, 15, 30, 45, and 60 minutes, and spotted on a PEI-cel lulose TLC plate. ATP was resolved from single phosphate groups by developing the TLC plate in 0.375M K H 2 P 0 4 (pH 3.5). The raw data is shown in the absence (A) and presence (B) of R N A in the assay at 30°C. C. The assay was repeated with varying amounts of R N A (0-50-fold molar excess of RNA relative to CsdA). Released Pi at 60 minutes was quantified using Phosphorlmager technology, and is plotted as a function of RNA content. A "best fit curve" corresponding to the equation "% Pi released at 60 min = 20.27 * (1-e~ 1 5- 2 5 * ([RNA]:[CSCJA]))" j s shown as well. D. The amount of Pi released through the course of 30°C assays is plotted. The data represent the average value of at least 5 experiments, and the error bars show the standard error of the means. Assay conditions were as follows: (•) CsdA, (•) CsdA and unlabeled in w'fro-transcribed rpsT mRNA, (B) CsdA, unlabeled in w'fro-transcribed rpsT mRNA and Rne, (o) degradosomes, (•) degradosomes and mRNA. The amount of RhIB in the degradosomes used in these assays approximate the molar amount of CsdA. E. Assays performed as in D, but at 15 °C. 165 Quantified data are shown in panels C, D, and E of Figure 4.9. Panel D shows the ATPase assay carried out at 30°C, while panel E shows an assay performed at 15°C. The activity seems identical at both temperatures. In the absence of RNA (•), no ATPase activity is detectable over the 1 hr time course at either temperature. The addition of RNA (•) stimulates the hydrolysis of ATP over 50-fold. RNA on its own (no CsdA present) had no effect (result not shown). The addition of Rne in addition to RNA (a) did not further stimulate the ATPase activity present in the assay at either temperature. Neither PNPase nor DNA exerted any stimulatory effect on CsdA activity (results not shown). The RNA dependence of CsdA's ATPase activity was further explored by varying the amount of nucleotide present in the assay and measuring the amount of phosphate released at 60 min. Stimulation of activity is observed starting at around 5 molecules of RNA per 100 CsdA and plateaus around a 1:1 ratio (Figure 4.9, panel C). It has been reported that RhIB has an ATPase activity that can be markedly enhanced by the addition of RNase E (Vanzo et al., 1998). However, the RhIB protein purified as described in Section 3.2.2 had no detectable activity, nor could it gain more by addition of RNase E (results not shown). To compare the ATPase activities of RhIB and CsdA under the current assay conditions, whole degradosomes were used as a source of RhIB. There is no accurate stoichiometry for the degradosome, but it can be assumed that roughly 10% of its 166 mass is due to the presence of RhIB1 1. Using this estimate as a guideline to calculate the amount of RhIB added, similar molar concentrations of CsdA and RhIB were assayed. Using this assumption, RhIB was found to have only a modest ATPase activity compared to CsdA. In fact, CsdA is 10.4-fold more active than RhIB. The results of the 30°C assay are shown in Figure 4.9, panel D. RhIB's very weak ATPase activity ( O ) is also stimulated by the presence of RNA (•). 4.3 DISCUSSION 4.3.1 Evidence for CsdA in cold shock degradosomes When these studies were initiated, little was known about mRNA degradation at 15°C. Bulk mRNA was known to be stabilized by cold (Wice & Kennell, 1974, Goldenberg et al., 1996), and PNPase was a recognized cold shock protein whose absence made cells cold sensitive (Jones et al., 1987, 1996; Clarke & Dowds, 1994; Luttinger et al., 1996; Wang & Bechhofer, 1996; Goverde et al., 1998). No one had ever reported the existence of degradosomes at 15°C, and the stabilization of bulk mRNA allowed for the possibility that there might be none. Purification of degradosomes from cells grown at 15°C showed they contain RNase E, PNPase, RhIB, enolase, the RNA polymerase subunits a, (3 and/or (3', the pyruvate dehydrogenase complex subunits E1 and E2, and CsdA. 1 1 This assumes that the relative composition of the degradosome is a dimer of RNase E, two 167 Except for CsdA, all other novel proteins co-purifying with the degradosome at 15°C are part of large complexes. Two of the steps in the degradosome purification procedure (ammonium sulfate precipitation and size exclusion chromatography) rely on the large size of the degradosome to separate this complex from other proteins in the extract. It is possible, therefore, that the RNA polymerase complex (Mr 390,000) and the pyruvate dehydrogenase complex (M r > 4,500,000) are not associated with the degradosome but merely co-purify with it because of their large size. Indeed, neither the (3 nor |3' subunit of the RNA polymerase complex (Figure 4.4, panel A, lane 2 - the a subunit of RNA polymerase is too small to be seen on this gel) immunologically co-purified with FLAG-tagged RNase E. These results show that RNA polymerase does not tightly associate with degradosomes. In contrast, CsdA and the E1 and E2 subunits of the pyruvate dehydrogenase complex co-precipitated with FLAG-RNase E (although in lesser amounts than in the biochemical preparation). The electron micrograph of the cold shock degradosome preparation found no evidence of a tight association between pyruvate dehydrogenase and degradosomes. The high level of expression of subunits E1 and E2 during cold shock (Jones & Inouye, 1994; Gualerzi et al., 2003), the stoichiometric composition of pyruvate dehydrogenase (24 copies of subunit E1, 24 copies of subunit E2, and 12 copies of subunit E3 (Eley et al., 1972; Reed, 1974; Angelides et al., 1979)), and the reduced quantity of E1 and E2 in trimers of PNPase, two dimers of enolase, and a dimer of RhIB. 168 immunologically purified degradosomes compared to those isolated biochemically suggest that this complex associates non-specifically with the degradosome due to its abundance after cold shock. Perhaps the rigidification of membranes at cold shock causes the degradosome and pyruvate dehydrogenase to co-localize on membranes, and thus co-purify. This does not imply any functional relationship between the two complexes. Pulse-labeling and immunoprecipitation experiments further confirmed that CsdA is present in degradosomes formed in the cold. It is more abundant in the complex at the end of the acclimation phase than during the recovery period. This suggests that the composition of the 15°C degradosome is dynamically adjusted throughout cold shock. Pull-downs using anti-CsdA beads only recovered CsdA. This likely reflects an excess of CsdA over degradosomes, as is the case for PNPase and enolase (Liou etal., 2001). Surprisingly, while there appears to be slightly more CsdA. in the anti-CsdA immunoprecipiate from cold-shocked cells, CsdA is found in significant amount in the immunoprecipitate of 37°C cells. CsdA is known to be expressed at 37°C, but is significantly induced upon cold shock (Section 4.2.1; Jones et al., 1996; Brandi et al., 1999). The amount of CsdA found in the 10 ml of culture used in these immunoprecipitation experiments (Section 2.9.1) may exceed the binding capacity of the anti-CsdA column, leading to an under-representation of the amount of CsdA present in the immunoprecipitate of cold-shocked cells. 169 CsdA co-purifies with RNase E in the absence of PNPase. This argues for a direct interaction between RNase E and CsdA, and reinforces the similarity between RhIB and CsdA (Vanzo et al., 1998; Coburn et al., 1999). In the absence of PNPase, RhIB also co-purifies with RNase E, which suggests that if the reported RhIB-PNPase association exists, it is not critical for the association of the helicase with the complex (Section; Vanzo et al., 1998; Coburn et al., 1999; Ow et al., 2000; Liou et al., 2002). It is impossible to determine from these experiments whether RhIB and CsdA compete for a binding site on RNase E. The presence of RhIB and CsdA in the IP does not imply that degradosomes contain both proteins. It could reflect the presence of two alternate forms of the degradosome in the cell, one containing RhIB and one with CsdA. Oddly, the absence of CsdA in the IP from csdA mutant extract is correlated with an absence of RhIB in degradosomes. Perhaps CsdA stimulates recruitment of RhIB to the degradosomes during cold shock, implying that RhIB and CsdA do not compete for a similar binding site on RNase E. Thus far, all proteins known to interact with RNase E bind the C-terminal region of this protein. Given that CsdA may be involved in recruiting the S1 protein to the ribosome, it may have the ability to recognize and bind the S1 domain of RNase E. This would imply that CsdA can bind the N-terminus of RNase E. It would also imply that, although not detected here, an association between PNPase (which also contains an S1 domain) and CsdA is possible. Regardless of whether CsdA uses the S1 domain to bind to the degradosome, its presence in the 170 degradosome may explain the presence of the S1 protein in this complex (Section; Feng etal., 2001). In vitro reconstitution of a "minimal cold shock degradosome" containing RNase E, PNPase, and CsdA confirms that these proteins can assemble into a complex at 15°C. The amount of purified CsdA required to form this complex is slightly larger than for other degradosomal proteins. This may be due to the histidine tag present on CsdA, but more likely suggests that the association between RNase E and CsdA is not as strong as the other degradosome interactions. All of these results confirm the presence of CsdA in cold shock degradosomes. This is perhaps not surprising, given that, as mentioned in Section, the RNase E-based degradosome of Rhodobacter contains two helicases, one of which is the homologue of CsdA (R.W. Simons, personal communication; Jager et al., 2001). There is no doubt that DEAD-box helicases are important components of all RNA decay machines (Section 1.6.11). Here, evidence is presented that a decay machine is remodeled in response to stress through changes in its helicase composition. Perhaps helicases are the "gears" that can be shifted to adapt the activity of the RNA degradation machine to changing conditions. 171 4.3.2 Reconstitution of CsdA-containing "minimal degradosomes" mRNA degradation becomes inefficient at cold shock. This may be caused by RhIB's inability to overcome cold-stabilized RNA secondary structures. CsdA is a cold shock protein with similarity to RhIB that appears in the degradosome after a temperature downshift. It may possess a more active helicase activity that can functionally replace RhIB in the degradosome and adapt the complex to the cold environment. To investigate this possibility, the degradation of the MalEF RNA was reconstituted in vitro in the presence of either helicase at 30°C and 15°C. The conclusion from these assays is that the activity of RhIB and CsdA are indistinguishable. This would suggest that CsdA can functionally replace RhIB in the degradosome, and that it can interact with PNPase to degrade 3' stem-loops. One surprising finding is that the activity of RhIB and CsdA are identical at both temperatures. Given the hypothesis that RhIB is a "weaker" helicase, this enzyme was not expected to overcome the REP hairpin at 15°C with as much ease as CsdA. The structure of the MalEF RNA is stabilized by the cold temperature, as indicated by the appearance of more * intermediate in the 15°C assays. PNPase and CsdA are known to be involved in the degradation of cold mRNA in the early stages of the recovery phase (Yamanaka & Inouye, 2001). Perhaps a better substrate for this assay would be the mRNA of a cold shock protein (e.g. cspA mRNA), which may adopt a particular configuration that requires the activity of CsdA. 172 Surprisingly, the activity of CsdA, like that of RhIB, is ATP-dependent. CsdA had already been shown to have the capacity to unwind double-stranded RNA in vitro in the absence of ATP (Jones et al., 1996). The assay used to reach this conclusion was based on the separation of two strands of RNA bound together by 29 complementary base pairs. The addition of CsdA generated increasing amounts of single-stranded RNA. One problem with this assay is that large amounts of protein can non-specifically coat RNA and drive strand dissociation by trapping products. The RNA:helicase ratio must therefore be monitored carefully. The amount of RNA incorporated in the assay performed by Jones and coworkers is not disclosed in their 1996 paper. However, their assay is based on the one described by Flores-Rozas & Hurwitz (1993), in which 50 fmoles of RNA are used. Jones and coworkers used 0.14 nmoles to 2.7 nmoles of CsdA in their assay, and only observed dissociations starting at 0.68 nmoles of CsdA. If these concentrations were used in the assay described above, then there is a very large excess (13,600-fold) of CsdA relative to RNA in the assay. CsdA's apparent helicase activity in the absence of ATP was likely non-specific. It has also been previously reported that purified CsdA does not catalyze the hydrolysis of ATP, even in the presence of polynucleotides (Lu et al., 1999). Assays described in Section 4.2.4 clearly show that CsdA is an RNA-dependent ATPase. CsdA's ATPase activity is dependent upon the composition of the buffer in which it is incubated (CsdA had much reduced ATPase activity when 173 incubated in rne assay buffer compared to pnp assay buffer (data not shown)). Perhaps Lu and coworkers did not find appropriate conditions for the assay. CsdA is 10-fold more active than RhIB, even at 30°C. This fits very well with the hypothesis that CsdA is a more active helicase than RhIB, and that it is capable of overcoming stable stem-loops which inhibit RhIB. Unlike RhIB which may require an interaction with RNase E in order to fold its RNA binding domain and have ATPase activity (Section; Py etal., 1996; Vanzo etal., 1998), CsdA is apparently able to bind RNA and hydrolyze ATP in the absence of RNase E. This does not imply that RNase E is not required for its role in mRNA degradation: binding to RNase E may optimally position PNPase and CsdA for the degradation of stable stem-loop structures (Figure 1.3). 4.3.3 mRNA decay and cold shock The mRNA decay apparatus that operates at 37°C is inhibited at cold shock (Wice & Kennell, 1974, Goldenberg et al., 1996). This contributes to the induction of cold shock proteins that adapt the cell for growth in the cold (Fang et al., 1997). Following cold adaptation, the mRNA encoding these proteins must be degraded to prevent the dangerous accumulation of cold shock mRNA and allow growth resumption. As though the cell had foresight, two of the proteins that are induced by cold shock (PNPase and CsdA) are involved in removing cold shock mRNAs at the end of the acclimation phase (Yamanaka & Inouye, 2001; Polissi etal., 2003). 174 The hypothesis that directed the experiments described in this chapter is that CsdA replaces RhIB in the cold shock degradosome because it has a more active helicase activity, which is preferred under cold conditions. CsdA was shown to physically associate with the degradosome, and functionally interact with PNPase to degrade RNAs with stable 3' stem-loops. Given the apparent lack of difference in the degradation of MalEF using degadosomes containing RhIB and CsdA, the functional significance of cold shock degradosome formation remains to be determined. It may be that a difference exists, but simply was not uncovered because substrates that would be naturally encountered in the cold (i.e. cold shock mRNA) were not tested. Another question that needs to be answered is the advantage that is gained by the cell when CsdA joins the degradosome. Previous studies have shown that in the absence of degradosome formation, cold shock mRNA degradation is not affected (Yamanaka & Inouye, 2001). Perhaps the advantage conferred by degradosome assembly is as subtle at 15°C as it is at 37°C and requires a more detailed analysis than was carried out thus far. Given that RNase E initiates the degradation of most mRNAs in E. coli, and that bulk mRNA is stabilized upon cold shock, it appears that this enzyme becomes inefficient in the cold (see also Afonyushkin et a l . , 2003). The recent isolation of RraA, an inhibitor of RNase E, may offer some insight into this process (Lee et al., 2003). However, the rapidity with which mRNAs are stabilized upon cold shock (a dramatic response is detectable 10 sec following 175 cold shock; Beran & Simons, 2001), suggests an alternative method of preventing RNase E function. Given that RNase E is a membrane-bound enzyme, and that membranes are immediately rigidified in the cold, the activity of RNase E may be affected by changes in membrane fluidity. Alternatively, the immediate stabilization of mRNA secondary structures may affect the activity of RNase E, by shielding cleavage sites in novel folds. Whatever the mechanism, RNase E is clearly involved in the induction by stabilization of the cspA mRNA in the cold (Fang et al., 1997). RNase E activity is presumably resumed upon acclimation, as the half-life of the cspA mRNA is reduced (Goldenberg et al., 1996). Could CsdA be involved in reactivating RNase E, either by direct protein interaction, or by unwinding RNA structures? A study of the effect of heat shock on mRNA stability has failed to detect any effect of temperature upshift on RNA decay (Henry et al., 1992). However, under certain conditions, namely growth on minimal medium lacking casamino acids, an upshift to 44°C resulted in the reduction of mRNA cleavages by RNase E, and the stabilization of both rpsO and rpsT transcripts (9S RNA was not affected) (Le Derout et al., 2002). Furthermore, the amount of Rne protein decreased 1.5-fold 15 min after upshift, and 1.9-fold 30 min after heat shock. These effects were not observed in the presence of casamino acids (i.e. in a richer medium). Another study found an alteration in the processing of the rne transcript following a temperature upshift and the addition of casamino acids to cells grown in minimal media (Woo & Lin-Chao, 1997). In this chapter, 176 immunopurification of FLAG-tagged RNase E from cells grown at 42°C did show a lesser quantity of RhIB in the heat shock complex. During heat shock, RNA secondary structures will be destabilized, and PNPase will no longer require as much assistance from helicases. This result therefore fits the interpretation developed in this chapter that the presence of helicases is modulated in the degradosome in response to temperature stress. 4.3.4 A model for expression and function of CsdA during cold shock In the first few minutes of cold shock, degradosomes are sequestered to the membrane and rendered inefficient. This causes the stabilization of bulk mRNAs and allows the induction of cold shock proteins. Among these are CsdA and PNPase, which accumulate. Two hours later, the cell has adapted to cold conditions. Membrane fluidity is restored, and growth resumes. New degradosome components are synthesized. The excess CsdA present in the cell at this time favours its association with RNase E and assembly of a cold shock degradosome. This complex may confer some advantage to the cell, such as facilitating the degradation of cold shock mRNAs at low temperature. There is ample evidence that CsdA has a role in translation or ribosome biogenesis during cold adaptation (Toone et al., 1991; Jones et al., 1996; Lu et al., 1999; Moll et al., 2002). These degradosome-independent functions are not 177 excluded by the one suggested above. Rather, CsdA is proposed to be a multifunctional protein with many roles in adapting the cell to cold conditions (Figure 4.10). The abundance of CsdA in cold-adapted cells supports the possibility that CsdA has many roles in the cell. What the data presented in this chapter show is that CsdA is found in the degradosome at 15°C, and that CsdA-containing degradosomes can fully replace the activity of RhIB-containing degradosomes in in vitro assays. 178 UPON COLD SHOCK Sequestration of existing degradosomes in membranes TCsdA, PNPase 2 hrs later Pool of CsdA, PNPase R N a s e E, RhIB, eno lase Ribosome Translation mRNA CsdA-degradosomes assembly decay + RhIB-degradosomes v J \ J Degradosome-independent Degradosome-dependent functions of CsdA functions of CsdA Figure 4.10 Model of CsdA activity during cold shock U p o n co ld shock , ex ist ing d e g r a d o s o m e s are immedia te ly seques te red in rigidif ied m e m b r a n e s , caus i ng the stabi l izat ion of bulk m R N A and a l lowing co ld shock proteins (such as C s d A and P N P a s e ) to be e x p r e s s e d . T w o hours later, C s d A and P N P a s e have accumu la ted to substant ia l levels in the ce l l . D u e to the a b u n d a n c e of C s d A in the cel l at co ld shock , C s d A is a p roposed mult i functional e n z y m e with roles in r ibosome a s s e m b l y , t ranslat ion, and m R N A degradat ion (both a s a componen t of the d e g r a d o s o m e and act ing independent ly of the comp lex ) . D e g r a d o s o m e - b o u n d C s d A m a y be or iented in s u c h a w a y that it p romotes the coord ina ted activity of this e n z y m e with P N P a s e , and faci l i tates the degradat ion of co ld-s tab i l i zed structures. 179 CHAPTER V CONCLUSIONS 5.1 The role of enolase in the degradosome Enolase is somewhat different from other components of the degradosome. On the one hand, enolase cannot associate with RNase E as readily as the other proteins, and may even require the assistance of PNPase and other factors to form a complex with RNase E. On the other hand, enolase does not appear to carry out a function in mRNA decay that is as easily identifiable as those of RNase E, PNPase, and RhIB. Though not exhaustive, the studies undertaken here to uncover a role for enolase in regulating the activity of the degradosome showed very little sign that enolase performs such a role. It seems more likely that enolase is involved in RNA binding during 5'-end-independent entry of the degradosome on substrates that lack a free 5'-end. Experiments that are currently underway should soon clarify this possibility. 5.2 Cold shock and the RNA degradosome While mRNA decay is stalled in the first 2 hrs after a temperature downshift, degradosomes are still present in the cell, and new degradosomes assemble with novel proteins not found in the 37°C complex. The most prominent addition is CsdA, a cold shock DEAD-box helicase. The association between RNase E and CsdA is robust and can be recapitulated by purified proteins in vitro. CsdA can replace RhIB in an assay which recreates the 180 degradation of a model RNA substrate in vitro. While CsdA can functionally interact with PNPase to degrade stable hairpins, it does not appear to be a more efficient helicase than RhIB on the MalEF substrate, even at 15°C. Changes in degradosome composition during cold shock must adapt the degradosome such that it can perform particular tasks under those conditions. Cold shock mRNAs are specifically targeted for degradation 2 hrs following cold shock, and may contain structures that require the presence of CsdA in the degradosome. The activity of RhIB may be insufficient for this purpose. This possibility was not tested by use of a cold shock substrate in in vitro reconstitutions, but appears likely given that both PNPase and CsdA have recently been shown to be involved in the degradation of such mRNAs (Yamanaka & Inouye, 2001). Despite earlier reports, CsdA is both an ATP-dependent RNA helicase and an RNA-dependent ATPase. One elusive piece of data is that CsdA, unlike RhIB, appears to have ATPase activity, and by inference helicase activity, in the absence of RNase E. Perhaps the only purpose of the RNase E-CsdA interaction is to bring PNPase and CsdA in close proximity, but given the excess of these two proteins over RNase E, it seems a direct interaction might have been more productive. Perhaps this is precisely why such a direct interaction never occurred: to leave free CsdA to carry out its other roles in ribosome biogenesis/translation. This would assume that RNase E is required to place CsdA and PNPase in a correct and productive orientation to each other, but the dependence of the degradation on RNase E could not be tested. 181 5.3 Perspectives on adaptation Two global mechanisms of adapting the mRNA decay machinery to new conditions were investigated. One examined the role of enolase in the degradosome, since this protein is strategically placed to both sense the metabolic environment of the cell and regulate the activity of RNase E, PNPase, and RhIB. The other searched for changes in the composition of the degradosome under cold shock conditions. Enolase could not be shown to affect the activity of the degradosome, but studies of cold shock successfully identified that a helicase is added to the decay machinery in response to new conditions. There are other mechanisms that could be used to globally regulate the activity of the degradosome. Marchand and colleagues (2001) have found that RNase E and RhIB are phosphorylated after T7 infection, and that the activity (and specificity) of the degradosome is changed in response to this signal. Similarly, Lee and coworkers (2003) have recently identified a protein that can inhibit the activity of RNase E and cause a global change in the stability of bulk mRNA. Whether this protein is expressed in response to specific signals or conditions remains to be established. While the possible methods of globally regulating RNA decay described in Section 1.7.1 focused on changing the activity of RNase E, which is responsible for the initiation of bulk mRNA decay in E. coli, the studies of cold shock adaptation and phosphorylation show that other 182 degradosome proteins could be the target of signals that change the activity of the degradosome. The basic mechanism by which mRNA is degraded in Escherichia coli is now sufficiently understood that the methods employed by the cell to regulate it can be investigated. Given the importance of mRNA in modulating gene expression, it seems likely that these studies will provide important insights into the complex organization of gene regulatory networks and fundamental mechanisms for adaptation. 183 REFERENCES Afonyushkin, T., Moll, I., Blasi, U., and V.R. Kaberdin (2003). 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