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The S1 domains of Escherichia coli mRNA degradation Edge, Robert Edward 2003

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The S1 domains of Escherichia coli mRNA degradation. by Robert Edward Edge Hon. B . S c , Simon Fraser University, 2000  A T H E S I S S U B M I T T E D IN PARTIAL F U L F I L L M E N T O F T H E R E Q U I R E M E N T S FOR T H E D E G R E E O F M A S T E R S O F S C I E N C E in THE FACULTY OF GRADUATE STUDIES Department of Biochemistry and Molecular Biology  We accept this thesis as conforming to the required standard  T H E UNIVERSITY O F BRITISH C O L U M B I A December, 2003 © Robert Edward Edge, 2003  Library Authorization  In presenting this thesis in partial fulfillment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission.  Robert Edward Edge  18/12/2003  Name of Author (please print)  Date (dd/mm/yyyy)  Title of Thesis:  Degree:  The S1 domains of Escherichia coli mRNA degradation  Master of Science  Department of  Biochemistry and Molecular Biology  The University of British Columbia Vancouver, B C Canada  Year:  2003  Abstract The S1 domain is a recurring theme in the enzymes of mRNA decay in Escherichia coli, occurring in polynucleotide phosphorylase (PNPase), ribonuclease II (RNase II), ribonuclease E (RNase E) and ribonuclease G (RNase G). This thesis focuses on the function of the S1 domain of P N P a s e and RNase E. In the former, the hypothesis examined was that the domain provides an R N A -substrate contact that imparts processive phosphorylysis to the enzyme. Results show that P N P a s e lacking this domain still possesses catalytic activity, although at an extremely reduced rate, but appears to retain processive phosphorylysis of R N A substrates. In addition, the isolated P N P a s e domain has a K for R N A that is very high, suggesting d  that this domain may not play a role in the interactions between P N P a s e and its substrate. The S1 domain of RNase E was studied as an isolated domain. N M R experiments conducted by Dr. Mario Schubert revealed the solution structure of this isolated domain as well as potential dimerization contacts and identified residues implicated in nucleic acid interaction. Crosslinking data confirmed the dimerization of this domain. C D spectroscopy showed that a classical temperature -sensitive mutation in this domain alters its structure. The work on the RNase E S1 domain also tested the hypothesis that the domain confers the preference of RNase E for monophosphorylated substrates. The data suggest that this domain indeed binds to R N A (Kd 2 \M to 5 ixM); however, it displays no increased affinity for differentially phosphorylated R N A s or detectable sequence specificity.  ii  Table of contents Page Abstract  ii  List of figures  vii  List of abbreviations  ix  1 INTRODUCTION  1  1.1 m R N A degradation in E.coli  1  1.1.2 Mechanisms and Models of m R N A decay  2  1.1.2 Enzymes of m R N A decay in E.coli  4  1.2 ob-folds  5  1.2.1 RNase E  7  1.2.2 P N P a s e  10  1.3 Focus of this study  15  2 MATERIALS AND METHODS  17  2.1 Source of Reagents  17  2.2 Bacterial Strains and culture techniques  17  2.3 Common Buffers  17  Table 1: Table of strains  18  2.4 P C R and site-directed mutagenesis  19  2.5 Other molecular biological methods  19  Table 2: Table of primers  20  2.6 Enzyme Purifications  20  2.6.1 P N P a s e  20  iii  2.6.2 S1 domain of RNase E  21  2.6.3 RNase E  23  2.7 Protein Analysis  23  2.7.2 DNA titration  23  2.7.3 Mass spectroscopy  24  2.7.4 UV quantification  24  2.7.5 C D spectroscopy  24  2.7.6 Sodium dodecylsulphate polyacrylamide gel electrophoresis ( S D S - P A G E )  25  2.7.7 Western Blotting  25  2.7.8 Glutaraldehyde crosslinking  25  2.8 R N A substrates  26  2.8.1 Template preparation  26  2.8.2 In-vitro transcription  26  2.8.3 Monophosphorylation of R N A  27  2.8.4 End -labelling of homopolymer R N A  28  2.8.5 Exoribonuclease assay  29  2.8.6 Endonuclease Assay  29  2.9 Filter binding assay  30  3 RESULTS  31  3.1 RNase E S1 domain  31  3.1.1 Overview: nature of the problem/hypothesis...:  31  3.1.2 Expression and Purification  31  iv  3.1.3 C D Spectroscopy  32  3.1.4 Protein structural properties  37  3.1.5 Dimerization  40  3.1.6 R N A binding studies  43  3.1.6.1 Filter binding studies  43  3.1.6.2 pH effects on R N A binding  44  3.1.6.3 Ionic effects on R N A binding  44  3.1.6.4 Effects of Mg++ on R N A binding  47  3.1.6.5 Monophosphorylation and triphosphorylation effects on R N A binding  47  3.1.6.6 Effects of the temperature-sensitive mutation, G 6 6 S , on R N A binding  50  3.1.7 Competition  50  3.1.8 DNA titration  54  3.1.9 Design of RNase E S1 domain mutants  58  3.1.9.1 S1 domain alignment  58  3.1.9.2 Analysis of structure in terms of potential interaction residues  58  3.1.10 Mutant S1 domain properties  60  3.1.11 RNase contamination  61  3.2 P N P a s e S1 domain  64  3.2.1 Activity of His-tagged and untagged P N P a s e  66  3.2.2 Processivity  68  v  3.2.3 Cold-sensitive phenotype of pnp mutants  71  3.2.4 Filter binding assay  71  4 DISCUSSION  76  4.1 function of RNase E S1 domain  76  4.1.1 R N A binding  76  4.1.2 Specificity  77  4.1.3 Structure and dimerization  79  4.1.4 R N A binding residues  82  4.1.5 rne-1 mutation  84  4.2 Function of P N P a s e S1 domain 5 BIBLIOGRAPHY  86 89  vi  List of Figures  Page Figure 1 Figure 2  Model of m R N A degradation in E.coli The S1 domains of enzymes of E.coli degradation  3 6  Figure 3  P N P a s e domain and quaternary structure  13  Figure 4  Purification of the S1 domain of RNase E  33  Figure 5  Purification of the me-1 S1 domain of RNase E  34  Figure 6  C D spectra of the RNase E S1 domain  35  Figure 7  C D spectra of the me -1 RNase E S1 domain  36  Figure 8  Thermal stability of S1 domains as measured by C D  38  Figure 9  The solution structure of the S1 domain of RNase E  39  Figure 10  The crystal structure of the S1 domain of RNase E  41  Figure 11  Cross linking of various S1 domain preparations  42  Figure 12  Effect of pH on R N A binding of the S1 domain  45  Figure 13  Effect of KCI concentration on R N A binding  46  Figure 14  Effect of Mg ion concentration on R N A binding  48  Figure 15  Binding of triphosphorylated and monophosphorylated R N A by the S1 domain of RNase E  49  Figure 16  Binding of homopolymeric R N A to the S1 domain  51  Figure 17  Binding of R N A to the wild type and me-1 S1 domain  52  Figure 18  Binding of R N A to the wild type and me-1 S1 domain at various temperatures  Figure 19  53  Timecourse of RNase E activity in the presence of excess S1 domain  Figure 20  55  Chemical shift perturbation of residues involved in R N A binding  57 vii  Figure 21  S1 domain sequence alignment: RNase E, P N P a s e and RNase G  Figure 22  59  R N A binding by S1 domain mutants: F57A and R109A  62  Figure 23  Nuclease activity of RNase E S1 domain preparations..  63  Figure 24  Western blot of RNase E S1 domain preparations  65  Figure 25  Phosphorolysis of wild type and his-tagged PNPase....  67  Figure 26  Schematic of P N P a s e deletion mutants  69  Figure 27  Phosphorolysis activity of P N P a s e deletion mutants  70  Figure 28  Phosphorolysis and polymerization reactions of P N P a s e AS1  72  Figure 29  Rescue of cold -sensitive phenotype by P N P a s e AS1..  73  Figure 30  Purification of the S1 domain of P N P a s e  74  Figure 31  The location of the putative phosphate binding pocket..  80  Figure 32  A model of RNase E multimerization  83  viii  List of Abbreviations 5' UTR  5' untranslated region  BR10  sequence cleaved by RNase E  CD  circular dichroism  Csp  cold shock protein  DEPC  diethyl pyrocarbonate  DNase  deoxyribonucleic acid nuclease  DTT  dithiothreitol  EDTA  ethylenediamine tetraacetic acid  FPLC  fine performance liquid chromatography  g  gravity  His-tagged  hexahistidine tagged  IPTG  isopropyl-|3-D-thiogalactopyranoside  Kd  dissociation constant  KH  K-homology domain  LB  Luria-Bertani Broth  mRNA  messenger RNA  NMR  nuclear magnetic resonance  NOE  nuclear Overhauser effect  ob-fold  oligosaccharide/oligonucleotide binding fold  PBS  phosphate buffered saline  PMSF  phenylmethylsulfonyl fluoride  PNPase  polynucleotide phosphorylase  psi  pounds per square inch  RNase  ribonucleic acid nuclease ix  me-1  me containing the mutation G 6 6 S  SDS  sodium dodecyl sulphate  SDS-PAGE  S D S polyacrylamide gel electrophoresis  TBE  tris-boric acid-sodium E D T A  T  midpoint unfolding temperature  m  UV  ultraviolet  v/v  volume/volume  X  1 Introduction Ribonucleic acid (RNA) is one of the most fundamental and important constituents of the living cell. R N A is central to the process of gene expression and assumes such diverse forms as messenger R N A (mRNA), transfer R N A (tRNA), and ribosomal R N A (rRNA) to enable the conversion of the cell's genetic information to enzymes and structural proteins. Therefore, an essential part of understanding gene expression mechanisms involves a thorough knowledge of R N A metabolism and structure. Transcription is one important aspect of R N A metabolism, but is beyond the scope of this thesis. The focus of this work is mRNA turnover, another important aspect of R N A metabolism. 1.1 mRNA degradation in E.coli The quantity of a particular protein produced in the cell is a direct result of the concentration of the corresponding mRNA and its translational efficiency. The size of a particular mRNA pool, in turn, is a function of the transcriptional activity of the gene and the lifetime of the specific transcript in the cytoplasm. While it has long been known that mRNAs in the cytoplasm are unstable due to the presence of ribonuclease activity, Apirion first proposed a mechanism for the degradation of bacterial mRNAs in 1973 [1]. This was a paradigm shift from the view of mRNA degradation as a passive recycling of ribonucleotides to a complex mechanism of endoribonuclease and exoribonuclease activities that plays an active role in gene expression. It is now evident, 30 years later, that different mRNA species have half-lives ranging from less than 2 min to greater than 15 min in Escherichia coli (E. coli), the organism for which the most data on mRNA degradation exists (for a review see Coburn 1  and Mackie [2]). The active turnover of m R N A enables this model bacterium to adapt to environmental changes and to increase the effectiveness of negative transcriptional regulatory signals. Active turnover of m R N A also enhances energy conservation in the cell. While less than 5% of R N A in the cell is mRNA, it makes up 60% of newly transcribed RNA; thus, the recycling of ribonucleotides through active turnover is critical for efficiency [3]. It has also been shown that mRNA decay mechanisms allow for the differential expression of polycistronic transcripts [4]. To add to the complexity of prokaryotic mRNA degradation, small antisense R N A s have recently been discovered to play a role in selective mRNA decay [5]. Moreover, the enzymatic machinery of mRNA decay includes a multienzyme protein complex dubbed the degradosome [6] as well as discrete ribonucleases. The complexity of mRNA degradation pathways indicates the importance of this process for control of protein expression.  1.1.2 Mechanisms and Models of mRNA decay A modern model for mRNA degradation is presented in Figure 1, adapted and paraphrased from Coburn and Mackie [2]. This illustrates the coordination of exonucleases and endonucleases and the involvement of the degradosome. The pathway begins with a typical translationally active transcript as a 5' -triphosphorylated mRNA containing 3' -terminal secondary structure. The initial step is an endonucleolytic cleavage catalyzed by RNase E leaving a 5' - fragment containing a 3' - O H which, if unstructured, is subjected to the 3' - 5' exonuclease activity of P N P a s e and/or RNase II. These enzymes degrade such fragments to short limit oligonucleotides which are subsequently degraded by oligoribonuclease [7]. The products of the initial cleavage by RNase E are usually no longer functional as mRNAs and are left with a 5' monophosphorylated end. RNase E exhibits increased activity towards 5' 2  o 5'PPP  OH 3' (rate limiting)  5'PPP  1  RNase E  OH 3'  5'P-  OH 3'  J  RNase E (fast)  RNase 11/ PNPase  o  5' P. 5'PPP  OH 3*  ii.  PAP1/PNPase  OH 3'  AAAAAA3'  Degradosome  5'P.  nucleotides and limit oligonucleotides  LA  OH 3'  Figure 1: Model of mRNA decay in E.coli of a typical transcript containing a 5' triphosphate and 3' -terminal secondary structure. Degradation of the transcript starts with a rate limiting endonuclease cleavage by RNase E. This cleavage leaves single stranded R N A (left side of the diagram) that is subject to further degradation by the single-strand-specific 3' to 5' exonucleases, PNPase and RNase II. These reactions produce limit mononucleotides and oligonucleotides as products. The initial cleavage by RNase E (right side of the diagram) produces a R N A molecule (5' monophosphorylated) that serves as a better substrate for RNase E. The terminal secondary structure is resolved by the multienzyme degradosome containing RNase E, PNPase and an ATP dependant DEAD-box helicase, RhIB. The helicase unwinds the secondary structure and produces single-stranded R N A which is subjected to the 3' to 5' exonuclease activity of PNPase resulting in the production of limit oligonucleotides.  3  monophosphorylated RNAs so that subsequent endonucleolytic cleavages occur much faster than the initial cleavage and the RNA is rapidly degraded [8]. Since the both 3' exonucleases are specific for single-stranded RNA, fragments generated by RNase E cleavages that contain secondary structure are resistant to further degradation. These fragments are can be degraded through the action of the multienzyme complex known as the degradosome [9]. This complex spatially coordinates the activity of RNase E, PNPase and RhIB, an ATP-dependent DEAD-box RNA helicase [10]. The RNA helicase is able to unwind RNA secondary structure in a reaction coupled to ATP hydrolysis that leaves unstructured single -stranded RNA as a substrate for phosphorolysis by PNPase. This mechanism provides many opportunities for differential stabilization of transcripts such as through secondary structure content and location and the accessibility of the 5' -end of the mRNA. 1.1.2 Enzymes of mRNA decay in E.coli There are least 15 different RNases in  E.coli  evenly divided among  endoribonucleases and exoribonucleases. Endoribonucleases hydrolyze phosphodiester bonds within an RNA polymer while exoribonucleases cleave phosphodiester bonds at the ends. Most of the exoribonucleases are not physiologically significant in mRNA degradation but instead process the stable tRNA and rRNA of the cell. These include RNase PH, RNase D, RNase BN (tRNA specific), RNase T and RNase R (rRNA specific). A very abundant periplasmic RNase of E.coli is RNase I, which may at most has a passive role in mRNA degradation. RNase H and RNase HI are involved in degradation of RNA primers following DNA replication. The only double strand-specific RNase is RNase III, the gene for which is dispensable for viability [11]. While there is great diversity of ribonucleases with specific substrates in 4  E.coli only  RNase E (section 1.2.1), P N P a s e (section 1.2.2) and RNase II play a general role in mRNA degradation. RNase E and P N P a s e will be the primary topic of following sections.  1.2 ob-folds A common feature of some key enzymes of R N A processing and decay is the presence of an S1 domain. The alignment in figure 2 indicates the position of an S1 domain in the N terminal region of RNase E and RNase G and in the C terminal region of P N P a s e and RNase II. The S1 domain is named after ribosomal protein S 1 , the protein in which it was initially characterized, and is a subclass of the ob-fold domains. The ob-fold domain is a compact structural motif found in all three kingdoms and is frequently used for nucleic acid recognition (for reviews see [12-14]). The domain was first characterized by Murzin who named the ob-fold for oligonucleotide/ oligosaccharide-binding fold and after O.B. Ptitsyn [15]. These folds are grouped by topology and structure without significant sequence similarity. The domain consists of a five stranded antiparallel (3-barrel ranging between 70 to 150 amino acids. This variability is largely a result of differences in variable loop structures between the conserved region of (3-sheets. The conserved fold is arranged in a 1-2-3-5-4-1 (3-sheet topology that is described as a Greek key motif. The most variable region occurs between (3 sheets 3 and 4 and is often composed of an alpha helix which packs against the bottom of the barrel effectively 'closing' the barrel. The conserved binding face has (3 strands 2 and 3 at its center and is bounded by the loops between the sheets. There are exceptions to these generalizations and there are cases where the ob-fold interacts with different macromolecules. This domain is utilized by different proteins to bind R N A , ssDNA, oligosaccharides or proteins. It can even form the catalytic site of some 5  RNase E  1 N 35  RNase G  1061 125 470  N 26  128 711  PNPase  CTJC  N  9  RNase II  691  644 N 644  Figure 2: The S1 domains of enzymes of E.co// mRNA degradation. Each ribonuclease depicted is labelled with N and C terminal amino acid numbers. Grey boxes with amino acid numbers indicate the placement of the S1 domain of the enzyme. RNase E and RNase G, both endonucleases, have an S1 domain located near their N terminus while the exoribonucleases, PNPase and RNase II, have an S1 domain near their C terminus.  6  inorganic pyrophosphatases [13]. Generally these domains do not show any binding preference for nucleic acid sequence. In the majority of ob-fold proteins complexed with oligonucleotides, the bases of the nucleotide are in close contact with the protein while the phosphodiester bonds are mostly exposed to solvent. Nucleotides interact primarily via stacking interactions with aromatic amino acid side chains and interactions with hydrophobic aliphatic side chains of polar amino acids such as lysine and arginine. The E.coli ob-fold protein, C s p B , been mutagenized to confirm the importance of basic and aromatic side chains in R N A binding [16].  1.2.1 RNase E RNase E is an essential endoribonuclease of 1061 amino acids that is widely believed to be the principle RNase for the initiation of mRNA decay [17]. The data, however, are not completely unambiguous as, for example, most oligoribonucleotides isolated from E.coli contain 5' O H termini instead of the 5' -phosphomonoesters that would be produced from an RNase E cleavage [18]. What is of interest to this study is the function and specificity of RNase E, its domain and quaternary structure, and the involvement and interactions of this enzyme in the degradosome. While the first identified function of this enzyme was the processing of 9S rRNA into pre-5S rRNA by two specific cleavage events [19], it has become clear this essential enzyme has many functions in E.coli. Additionally it has been demonstrated to be involved in the processing of the 5' -end of 16S rRNA [20], the maturation of the R N A subunit of RNase P [21], the degradation of R N A I, the antisense regulator of colE1 plasmid replication [22, 23], the processing of most if not all tRNAs [24-26], the degradation of many mRNAs including trxA (thioredoxin), ssb (single-stranded-DNAbinding protein), uvrD (DNA helicase II), car (chloramphenicol acetyltransferase), nusA 7  (N utilization substance), pnp (polynucleotide phosphorylase) [27], rpsT[28-30],  rpsO  [31, 32], malEFG, trp operon, his operon, and total pulse labelled R N A [27, 33]. It has become clear from studies of RNase E substrates that the enzyme is single-stranded RNA-specific yet has a poorly defined consensus cleavage sequence [34]. The regulatory RNA, R N A I, contains a cleavage site that has been used extensively to model a consensus sequence and has been used as the basis for determination of various substrate sequence requirements of the enzyme [34-36]. Enzyme activity has also been demonstrated to require a 5' -unpaired residue. Also relevant to this study is the increased cleavage of 5' -monophosphorylated substrates over substrates that have a 5' -triphosphosphorylated end. RNase E is, therefore, said to exhibit 5' -end dependence [8]. This has lead to the hypothesis that this enzyme must contain a phosphate-binding pocket that preferentially binds monophosphorylated R N A thus accounting for its enhanced RNase E activity on these substrates. The catalytic mechanism for RNase E is unknown. It is hypothesized, however, that the enzyme uses a two-metal-ion mechanism [37] as catalysis requires the presence of a divalent metal ion M g  + +  or M n  + +  [38]. Analysis of the amino acid sequence  of RNase E reveals three distinct domains [39]. The N-terminal 500 amino acids encode the catalytic activity of the enzyme and are required for cell viability [40]. This domain is followed by an arginine and proline-rich domain (amino acids 567-684) demonstrated to have R N A binding properties in vitro [40-42]. This R N A binding domain (RBD) has been further delineated to residues 608-622, R R K P R Q N N R R D R N E R [2]. A similar motif is also found in the HIV-1 proteins Tat and Rev [43]. While this domain may have interesting implications for RNase E catalysis, including the recognition of cryptic cleavage sites [44], it is located in a region dispensible for catalysis [44, 45]. The C -  8  terminal domain (amino acids 734-1061) of RNase E is required for protein-protein interactions stabilizing the multienzyme complex known as the degradosome [46, 47]. Proteins shown to be a part of the degradosome and therefore interacting directly with this domain include P N P a s e , RhIB and enolase [48, 49]. The C-terminal domain is dispensable for catalytic activity, is not required for cell viability [50] and is not well conserved among prokaryotes, unlike the highly conserved N-terminal region of RNase E [51]. Nonetheless, it has been demonstrated that E.coli mutants containing this domain are more competitive in continuous culture than strains lacking the C-terminal domain [52]. The intracellular concentration of RNase E is regulated by an autoregulation mechanism. The rne transcript encoding RNase E is a substrate of its own product [53]. This autoregulation is mediated through a cleavage of the 5' -UTR of the rne mRNA [45, 54]. It also is through this mechanism that the rne transcript is stabilized in response to polyadenylation [55]. RNase E autoregulation requires the degradosome [55] and, therefore, the C -terminal region of RNase E [33, 56, 57]. Within the highly conserved N-terminal region from amino acids 3 5 - 125 is the putative R N A binding domain [58], an S1 domain, a member of the ob-fold domains. The two classical temperature-sensitive mutant loci, ams-1 (G66S) and me 3071 r  (L68F), map to this domain [59, 60]. These mutations inhibit cell growth at nonpermissive temperature. The more severe of these mutations, ams-1 /me-1, completely renders the enzyme inactive at 37°C in vitro [28](GA Mackie unpublished). These mutations highlight the importance of the correct folding of the S1 domain of RNase E for activity.  9  The only previous study of the S1 domain of RNase E modeled its three dimensional structure based on the structure of the P N P a s e S1 domain [58]. This enabled the engineering of mutations that were hypothesized to interact with R N A . Residues that were mutated in the study included aromatic and basic residues that were solvent-exposed in the model. The aromatic residues were hypothesized to provide base stacking interactions, while basic residues were hypothesized to provide ionic interactions with the phosphate backbone of the R N A ligand [61]. It was found that residues affecting R N A cleavage also affected autoregulation and mapped to one surface. Mutations that affected autoregulation alone and not in vitro catalysis mapped to a region slightly separated on the structural model from this first class of mutants [61]. The second region is not the typical surface of ob-folds that interacts with oligonucleotides. These mutations led the authors to hypothesize that this second class of mutations is located on a surface which recognizes specific R N A s such as the rne mRNA 5' -UTR. Alternatively, residues in the second region could be involved in a protein-protein interaction. The superantigens, a toxic subclass of ob-folds that show recent evolutionary divergence, provide precedent for ob-fold domains interacting with proteins [13].  1.2.2 PNPase E.coli Polynucleotide phosphorylase (PNPase) is a 711 amino acid product of the pnp gene [62, 63]. P N P a s e was originally discovered and described as a templatedependent R N A polymerase enzyme with a subsequent rich history, being involved in the determination of the genetic code and site-directed mutagenesis [64, 65]. The enzyme is not essential in E.coli; however, mutant strains display a cold-sensitive phenotype [66]. P N P a s e uses inorganic phosphate in a phosphorylytic reaction that has 10  an absolute requirement for magnesium ions (or other divalent ions), and yields nucleoside diphosphates as a product. While this phosphorylytic mechanism predominates in vivo, the reverse polymerization reaction also can be induced in vitro and constitutes much of the enzyme's early history [64]. Both the requirement for a divalent ion and the structure hint at a reaction mechanism based on two metal ions with phosphate as an attacking nucleophile [37, 67]; however, details of the mechanism of P N P a s e activity are unknown. Apart from the cold -sensitive phenotype, a null mutant of P N P a s e shows a mild phenotype and is viable. It is thought that the hydrolytic enzyme RNase II can compensate in the absence of P N P a s e . In support of this hypothesis, RNase II accounts for 90% of 3' - 5' exoribonucleolytic activity in crude extracts of E.coli [3]. Moreover, a double mutant of RNase II and P N P a s e is inviable [68]. It has also been argued that the two phosphorylytic exoribonucleases of E.coli, P N P a s e and RNase P H , have an essential role in ribosome biogenesis [66]. The phosphorylytic enzymes may also have evolved for energy conservation as products of the reaction are higher energy A D P molecules while the hydrolytic mechanism of RNase II produces A M P . The role of P N P a s e in mRNA decay is evident in the activity and regulation of the enzyme. In E.coli mRNAs targeted for degradation are often polyadenylated [69] and polyadenylated substrates stimulate P N P a s e activity [69, 70]. This may be because P N P a s e is single-strand specific [71], making poly (A) R N A an ideal substrate. In addition to increasing the activity of P N P a s e towards R N A substrates, increased levels of total poly A in the cell increase the cellular concentrations of both P N P a s e and RNase E through interference with their separate autoregulatory mechanisms [55]. Poly  11  (A) tracts may sequester these enzymes to polyadenylated transcripts thereby stabilizing their respective mRNAs. P N P a s e mutant E.coli cultures show greatly decreased growth rate at low temperature in comparison to wildtype. Recently it has been shown P N P a s e has a complex role in cold growth adaptation in E.coli [72]. P N P a s e is classified as a cold shock protein (CSP) and selectively degrades other C S P m R N A species following a cold shock response [73]. During the cold shock response expression of P N P a s e increases through a loss of its autoregulatory mechanism [74]. Normally, P N P a s e autoregulates its own expression by degrading the 5' -end of the pnp m R N A following an initiating cleavage catalyzed by RNase III [75]. Unlike the autoregulatory mechanism of RNase E, P N P a s e autoregulation, while dependent on RNase III, is independent of the degradosome [55]. The domain structure of P N P a s e consists of two duplicated regions with varying homology to RNase P H and two putative R N A binding domains, K H and S1 [76] (Figure 3). The crystal structure of Streptomyces antibioticus P N P a s e has been solved as a tungstate derivative [77]. This structure reveals P N P a s e is a trimer, and in a sense the enzyme consists of a trimer of the two duplicated regions homologous to RNase P H . The two duplicated regions are the two core domains of the trimer in the x-ray crystal structure of P N P a s e . The trimer is arranged with central pore where the tungstate electron density is located, hinting that this is the location of the site of phosphate binding. The two R N A binding domains, while not ordered in the crystal structure of Symmons et al., are recognizable motifs based on sequence. A K H domain is located between amino acids 557 - 591 in P N P a s e . The most strongly conserved motif in a K H sequence is a G K x G tetrapeptide and the structure  12  Figure 3: PNPase domain and quaternary structure. A. PNPase contains two domains with homology to RNase PH termed P H and PH' domains. B. These domains constitute a dimer of RNase PH-like domains in a trimer of PNPase. A central pore is evident in the PNPase trimer from the x-ray crystal structure of the Streptomyces antibioticus PNPase [59] and is indicated in the center of the PNPase trimer. The KH and S1 domains of PNPase were not localized in the crystal structure but are thought to be located out of the plane of the depicted structure.  13  consists of (3aa(3(3a fold [78]. This domain has been shown previously to be an s s R N A binding domain by Northwestern blot [79]. Other examples of a K H domain include the KH domain of ribosomal protein S 3 which can be cross-linked to ribosome-bound R N A [80], and E.coli NusA which has been shown to bind nascent m R N A [81]. The solution structure of the K H domain of human F M R 1 , which is associated with fragile X syndrome, has been solved and presents a hypothetical R N A binding surface [78]. A mammalian Nova K H domain has also been shown to have sequence specificity in binding R N A [82]. A detailed analysis of the interaction between any K H domain and R N A is, however, lacking. An analysis of P N P a s e engineered to lack the K H domain has shown that the mutant exhibits a slightly higher rate of phosphorylysis reaction, a lower rate of polymerization, but most significantly, a greatly reduced efficiency of autoregulation [83]. The S1 domain of P N P a s e is located between residues amino acids 619 and 691. The solution structure of this domain has been solved and contains a five stranded antiparallel (3-barrel, typical of ob-fold domains [58] and exhibits strong structural homology to cold shock domains. Early kinetic studies of P N P a s e have established a hypothesis that processive catalysis by P N P a s e required the presence of distinct R N A binding sites that are separated by up to 20 - 40 nucleotides along the R N A substrate [84, 85]. These binding sites would allow the enzyme to maintain contact with the substrate during successive catalytic events and impart a processive mode of action to the enzyme. More recent mutational analysis indicates that P N P a s e lacking the S1 domain, similar to P N P a s e lacking the K H domain, is deficient in autoregulation [83]. An interesting property of the S1 domain of P N P a s e is its ability when overexpressed to suppress the phenotype of a quadruple deletion in the cspA family of 14  cold shock genes. The quadruple mutant in these cold shock proteins exhibits a coldsensitive phenotype not unlike that of a P N P a s e mutant [86]. The C s p A protein has a similar tertiary structure to the S1 domain of P N P a s e and is also classified as an ob-fold protein [87, 88]. It has been hypothesized that C s p A is an R N A chaperone and maintains R N A in an unstructured conformation, thereby acting as a transcription antiterminator [89]. An NMR experiment in which C s p A was titrated with a DNA ligand has revealed the interacting surface and residues involved in this interaction [87]. This suggests the possibility that the S1 domain of P N P a s e serves a function similar to CspA, but in the context of a 3' - 5' single-stranded exoribonuclease, and possesses a similar nucleic acid binding surface. Ob-fold domains can also be involved in protein-protein interactions, as in the case of the bacterial superantigens where it serves as a dimer interface and to crosslink MHC-II molecules [13]. The C -terminal region of RNase E (residues 844 -1045) is known to interact with P N P a s e in the degradosome, but which part of P N P a s e is responsible for this interaction has not definitively been shown. Preliminary data suggest that the S1 domain of P N P a s e may be responsible for this interaction with the C terminal region of RNase E (Miao, X., unpublished). The role of the two R N A binding domains in P N P a s e remains unclear. It is possible that the K H and/or the S1 domains constitute the distinct R N A binding surfaces previously implied to be responsible for the processive nature of P N P a s e [84]. However, it is not known whether the effect on autoregulation is a result of loss of processivity or another function of these domains.  15  1.3 Focus of this study The aim of this thesis is the examination of the role of the S1 domain in m R N A decay in E.coli. A s described previously this domain is a recurring theme in enzymes involved in this process. Two specific S1 domains will be examined in this thesis, one from RNase E and the other from P N P a s e . The S1 domain of RNase E was examined as an isolated domain. The solution structure of this fold was solved by nuclear magnetic resonance spectroscopy by Dr. Mario Schubert. This study attempts to examine the importance of specific residues and the specificity of the R N A binding capacity of this domain using the obtained structural data. The me-1 mutation and some engineered mutations are also examined in this context. Evidence for the dimerization of this domain is also established. It has been suggested that the S1 domain of RNase E constitutes the hypothesized phosphate binding pocket responsible for the enzyme's activity towards monophosphorylated substrates. My data indicate that the S1 domain of RNase E alone can not account for this observation. The role of S1 domain of P N P a s e in mRNA degradation is examined in terms of the function of the enzyme in vitro and in vivo. The phosphorolysis activity of this enzyme was examined in a construct lacking the S1 domain to test the hypothesis that this domain provides substrate contacts that account for its processive catalysis. Surprisingly the processivity of the reaction is maintained while catalytic activity is greatly reduced. In addition the function of this domain in the cold shock phenotype of a P N P a s e null mutant is examined.  ,  16  2 Materials and Methods  2.1 Source of Reagents All chemicals were purchased from commercial sources and were of reagent grade. 2.2 Bacterial Strains and culture techniques Luria-Bertani broth (LB; 86 mM NaCI, 0.5% w/v yeast extract, 1 % peptone) was used as a rich medium for culturing bacteria. Agar plates (LB supplemented with 1.5% agar) were used as a solid medium for isolation of single colonies. Plasmid selection on either medium was carried out by supplementation with one or more of ampicillin (100 p.g/mL), kanamycin (30 ng/mL), or chloramphenicol (25 ^g/mL). Manipulations of bacterial strains were performed aseptically as described (Sambrook, 1989).  2.3 Common Buffers Lysis buffer: 25 mM Tris-HCI (pH 7.5), 0.1 mM EDTA, 2 mM DTT and 0.1 mM phenylmethylsulfonyl fluoride. Buffer B: 25 mM Tris-HCI (pH 7.5), 0.1 mM EDTA, 2 mM DTT and 5% glycerol. Anion exchange buffer: 50 mM Tris (pH 8.0), 50 mM NaCI and 0.5 mM DTT. Talon buffer: 50 mM Tris-HCI (pH 8.0) and 500 mM NaCI. Filter Assay Binding Buffer (FAB buffer) was made as a five-fold concentrated stock consisting of 100 mM sodium phosphate at pH 6.5 and 250 mM NaCI. Buffer A: 20 mM Tris (pH 7.5), 1.5 mM DTT, 1.0 mM MgCI and 20 mM KCI. 2  T B E buffer: 90 mM Tris, 90 mM Boric Acid, 2 mM EDTA.  17  Formamide Loading Buffer: 45 mM Tris, 45 mM Boric Acid, 1 mM EDTA, 90% deionized formamide, 0.02% each Xylene Cyanol and Bromophenol Blue. Tricine solution: 10 mM Tricine (pH 7.9-8), 50 mM NaOAc and 5 mM EDTA. SDS-sample buffer: 120 mM Tris-HCI pH 6.8, 3% sodium dodecylsulphate (SDS), 50 mM DTT, 10% glycerol, 0.1% bromophenol blue. Laemmli buffer: 25 mM Tris-HCI, 192 mM glycine, 0.1% S D S . Coomassie Brilliant Blue stain: 0.5 mg/mL Coomassie Brilliant Blue, 45% methanol, 10% acetic acid. Destaining solution: 5% acetic acid, 5% ethanol. Transfer buffer: 3 mM N a C 0 , 10 mM N a H C 0 , 20% methanol. 2  3  3  P T B N : 20 mM sodium phosphate (pH 7.0), 0.05% Tween, 0.1 mM bovine serum albumin, 0.85% NaCI, 1 mM N a N . 3  P B S : 137 mM NaCI, 2.7 mM KCI, 10 mM N a P 0 , 1.8 mM K H P 0 , adjusted to pH 7.4 2  4  2  with HCI.  Table 1: Table of strains  Strain  Genotype  BL2KDE3)  F" ompT hsdS (rB,  DH5a  F" (/)dlacZAM15 AilacZYA -argF)  B  m ") gal, dem A(DE3) B  recA1 endA1 hsdR17(rK-, thi-1 ENS134  U169deoR  mK+) phoA supE44 X-  gyrA96relA1  F" ompThsdSB^B,  m ") gal, dem A(DE3), B  pnp::Tn5  18  4  2.4 PCR and site-directed  mutagenesis  Amplification of specific DNA sequences was achieved using the polymerase chain reaction (PCR) [90] in conjunction with a Perkin Elmer GeneAmp 2400 programmable thermocycler. P C R reactions typically consisted of up to 250 ng plasmid template or 1 \xg genomic template DNA, 200 ng each of forward and reverse synthetic oligonucleotide primers, 0.2 mM each of the four deoxynucleotide triphosphates and 5 U of commercially available Taq DNA polymerase in the buffer supplied by the manufacturer, in 100 \iL in thin walled polypropylene tubes. Site-directed mutagenesis was achieved using a commercially available DNA polymerase with 3' to 5' proof-reading activity in a P C R reaction using the manufacturer's instructions as a guide (Stratagene Quick-Change kit). Synthetic oligonucleotide primers contained sequences that spanned the desired mutation and are listed in Table 2. The products of this reaction were treated with Dpn1 to digest the native dsDNA template. The remaining product was used to transform commercially competent D H 5 a (Invitrogen) to propagate the resultant plasmid for sequencing and transformation.  2.5 Other molecular biological methods Well established molecular biological procedures such as restriction endonuclease digestion, DNA ligation, plasmid transformation, plasmid extraction, and gel electrophoresis were performed as described in Sambrook (1989) unless otherwise noted.  19  Table 2: Table of primers Mutant Primer  Sequence  generated  RE16-M1  gaaatcgaagtgggccgcgtcgttgagaaagtgaccgattacc  P N P a s e AS1  RE16-M2  ggtaatcggtcactttctcaacgacgcggcccacttcgatttc  P N P a s e AS1  RE17-M1  cataccatcaagatcaacccggaagtgggccgcgtctacact  PNPase AKH  RE17-M2  agtgtagacgcggcccacttccgggttgatcttgatggtatg  PNPase AKH  RE23-M1  gttcagatcgataaagaagaggccggcaacaaaggcgcggc  R109A  RE23-M2  gccgcgcctttgttgccggcctcttctttatcgatctgaac  R109A  RE21-M1  gttgattacggcgctgaagctcacggtttcctcccactaaaag  F28A  RE21-M2  cttttagtgggaggaaaccgtgacgttcagcgccgtaatcaac  F28A  S1R35A-F  cggcgctgaagctcacggtttcctcc  R35A, E34D  S1R35A-R  ggaggaaaccgtgagcttcagcgccg  R35A, E34D  2.6 Enzyme  Purifications  2.6.1 P N P a s e P N P a s e was expressed in E.coli BL21 containing the plasmid pGC400 [91]. The bacteria were grown in 600 mL LB supplemented with ampicillin as selection for pGC400 at 37°C, until an O D  6 0 0  of 0.4 - 0.6 was reached. IPTG was added to the  cultures to 0.1 mM final concentration for induction of the expression of P N P a s e . Cultures were grown with aeration for an additional 4 hours at 37°C. Cells were harvested by centrifugation at 4, 000 x g for 10 min at 4°C. All subsequent steps were performed in a 4°C cold room. Cell pellets were resuspended in 5 mL of lysis buffer and 20  ruptured by passage through a French Pressure Cell at 10, 000 psi. The lysate, approximately 10 mL, was treated with 10 U DNase 1 for 15 min on ice. The lysate was then centrifuged at 30, 000 x g for 45 min and the supernatant was passed through a 0.22 jam syringe filter. The filtrate was applied to a F P L C column containing agarose supported heparin (4 cm x 2.5 cm), previously equilibrated with buffer B. The column was washed at 1 mL/min for 10 min before a gradient of NaCI from 0 - 1 M in buffer B over 2 hrs was used for elution. Fractions showing high UV absorbance at 280 nm were examined by S D S - P A G E . Those fractions containing P N P a s e , which eluted around 200 mM NaCI, were pooled and subjected to further chromatography. A SourceQ anion exchange column (4 cm x 2.0 cm), previously equilibrated with anion exchange buffer, was used on an F P L C at a flow rate of 1.0 mL/min. After the injection of the pooled fractions from the heparin column the column was pumped with anion exchange buffer for 10 min. A gradient of NaCI (50 mM-1 M in anion exchange buffer) was used to elute P N P a s e . Fractions containing P N P a s e were concentrated in a centrifugal filter with a 5 kDa cut off before being subjected to gel filtration on a 0.5 M cut-off column (48 cm x 2 cm) previously equilibrated with Buffer B containing 100 mM NaCI at a flow rate of 0.5 mL/min. The fractions containing P N P a s e as determined by S D S - P A G E were concentrated using a centrifugal filter and the resulting protein fraction was frozen in 50 \iL aliquots at -80°C after the addition of glycerol to 5%.  2.6.2 S1 domain of RNase E The RNase E S1 domain was expressed in E.coli BL21 containing the plasmid 'S1B-3', previously constructed by Mike Cook. The bacteria were grown in 300 mL LB supplemented with ampicillin at 37°C until early logarithmic phase. IPTG was added to the cultures 0.1 mM for induction of expression for 4 hours. The cells were then 21  harvested by centrifugation at 4, 000 x g for 10 min at 4°C and all subsequent steps were performed in a 4°C cold room. Cell pellets were suspended in 5 mL of talon buffer and ruptured by passage through a French Pressure Cell at 10, 000 psi. The lysate, approximately 15 mL, was treated with 10 U DNase I for 15 min on ice. The material was then centrifuged at 30, 000 x g for 45 min and the supernatant was diluted to 10 mL with talon buffer. This material was added to a 5 mL C o  + 2  chelating resin (Talon®,  Clontech) in a gravity flow column and a flow rate of about 2 mL/min was achieved by hand pressure, applied with a 10 mL syringe. The column was previously equilibrated with Talon buffer. Step gradients of imidazole in talon buffer were applied to the column to elute weakly bound proteins initially and then to elute the His-tagged S1 domain in one fraction. The column was first washed with four column volumes of 5 mM imidazole followed by one column volume of 15 mM imidazole. Protein was then eluted with 1.6 column volumes of 50 mM imidazole. This was followed by a wash of 1.6 column volumes of 500 mM imidazole to remove any remaining bound proteins. Fractions were analysed by S D S - P A G E for purity. The His-tagged S1 domain which eluted in the 50 mM imidazole fraction was concentrated using a centrifugal filter with a molecular weight cut-off of 5 kDa to a volume of about 1 mL. This sample was then dialysed in the presence of 2 U of human thrombin for 16 hrs against 1 L of Talon buffer. To remove the thrombin, 200 uL of p-aminobenzamidine agarose beads were added to the sample and placed on a "neutator" shaking table at 4°C for 1 hr. The samples were centrifuged at 16, 000 x g and the supernatant was diluted with Talon buffer to a volume of 5 mL. This sample was loaded on a pre-equilibrated Co+2 column and the flow-through was again collected at approximately 2 mL/min followed by a 5 mL wash with Talon buffer. This 10 mL flow through fraction was concentrated to 1 mL using a centrifugal filter with 22  a molecular weight cut-off of 5 kDa. This sample was dialysed for 16 hrs in 1 L of F A B buffer. Sterile glycerol to 5% v/v was added to the purified protein and the sample was stored at -80°C in aliquots. Purity and identity of the isolated domain was analysed by mass spectrometry as described in section 2.7.3. Various methods were used in an attempt to remove contaminating RNase activities from the purified RNase E S1 domain. An entire preparation was applied to a reverse phase H P L C column in which a gradient of acetonitrile was used to elute the protein. Fractions of 4 mL were collected in 10 mL test tubes that had been D E P C treated, and the UV absorbance at 280 nm was monitored during the collection. Fractions showing the highest UV absorbance were lyophilized before being dissolved in F A B buffer.  2.6.3 RNase E The preparation of the RNase E used in this thesis was done as described in a directed studies project.  2.7 Protein  Analysis  2.7.1 Nuclear magnetic resonance (NMR) spectroscopy Multidimensional NMR spectroscopy was performed by Dr. Mario Schubert and Dr. Lawrence Mcintosh. These data were used by Dr. Mario Schubert to determine the solution structure of the RNase E S1 domain.  2.7.2 DNA titration A solution of 1.4 mM RNase E S1 domain was titrated with increasing amounts of a 10 residue deoxyoligonucleotide, BR10, modeled after a characterized cleavage site in R N A I; 5' - A C A G T A T T T G -3' [22, 34-36]. NMR spectroscopy was used to determine 23  residues whose chemical shifts are altered by interaction with DNA. Spectra were measured by NMR at 0, 0.2, 0.4, 0.6, 0.8, 1.0, 1.1, 1.4 and 1.6 molar ratios of DNA to protein. All data were collected and analyzed by Dr. Mario Schubert.  2.7.3 Mass spectrometry 50 \xL samples of a 0.5 mg/mL solution of the protein of interest in F A B buffer were analyzed using electrospray mass spectrometry. The analysis was done by Dr. Shouming He at the U.B.C. Laboratory of Molecular Biophysics. 2.7.4 UV quantification Quantification of purified protein preparations took advantage of their UV absorbance at 280 nm, using theoretical extinction coefficients as calculated by http://us.expasy.org/tooIs/protparam.html, based on amino acid sequence. 2.7.5 C D spectroscopy All measurements were performed on solutions containing protein of at least a 0.25 mg/mL in a 400 uL, 0.2 mm quartz cuvette. Spectra were recorded on a Jasco J 810 spectropolarimeter, corrected for buffer background and reported in molar ellipticity (degrees.cm /decimole). Temperature variation experiments to determine an 2  approximate T were done at a temperature change rate of 2°C/min with a data pitch of m  0.5°C and at a wavelength of 225 nm. This wavelength was determined to be the most informative over the temperature range tested (10 - 80°C) for the wildtype RNase E S1 domain in F A B buffer. Wavelength scans were done at 10°C temperature intervals from 25°C to 75°C and scanned from 190 nm to 250 nm.  24  2.7.6 Sodium dodecylsulphate polyacrylamide gel electrophoresis ( S D S - P A G E ) Equal volumes of sample and SDS-sample buffer were mixed and boiled at 95°C for 2 min. Samples were then loaded onto a 7.5 or 15% polyacrylamide gel (36:1 or 29:1 respectively) containing 0.1% S D S and separated using Laemmli buffer at 140-190 V. After the bromophenol blue had run off the gel, proteins were visualized by staining with Commasie Brillant Blue in acetic acid and methanol for 5 - 10 min with agitation before being destained with destaining solution for as long as overnight.  2.7.7 Western Blotting Samples were initially separated by S D S - P A G E and then blotted to a nitrocellulose membrane at 250 mA for 2 hrs in Transfer buffer. These blots were then blocked in 15 mL P T B N for 1 hr with agitation at room temperature, before the P T B N was decanted and replenished with polyclonal primary antibody at an appropriate dilution (5, 000 - 10, 000 fold in PTBN) and again incubated for 1 hr with agitation at room temperature. Three separate washes with P B S for 5 min were followed by incubation of the secondary antibody, goat anti-rabbit polyclonal antibody conjugated to horse radish peroxidase, in 15 mL of P B S for 1 hr at room temperature. This blot was then washed three times for 5 min each with 15 mL P B S . The blot was then wetted with a peroxidase chemiluminescent substrate and exposed to film for 1 to 5 min.  2.7.8 Glutaraldehyde crosslinking Reactions consisted of 45 uM total protein in F A B buffer. Samples of 50 uL were incubated at 37°C for 2 min in 0.01 % glutaraldehyde. To quench the reaction 10 uL of 300 mM Tris-HCI pH 8.0 was added and placed on ice. To concentrate the sample for 25  analysis 40 uL of 17.5% trichloroacetic acid was added and the mixture left on ice for 30min. Samples were then centrifuged at 13, 000 rpm for 15 min at 4°C and the precipitate was washed twice with 200 uL 80% acetone. For S D S - P A G E analysis samples were dissolved in 10 ul_ of S D S loading buffer and denatured by boiling for 2 min at 95°C. 2.8 RNA  substrates  2.8.1 Template preparation All substrates used were previously designed, cloned and described in the Mackie lab. Substrates S L 9 and SL9A were generated using the plasmid pTZ18U [92] linearized with BamH1 and X b a l respectively. Linearized DNA was purified from undigested DNA by agarose gel electrophoresis followed by purification using a commercial gel purification kit (QIAGEN gel purification kit). The final product was dissolved in DEPC-treated sterile water. 2.8.2 In-vitro transcription Transcription of internally labeled R N A , for use as substrates for endoribonuclease and exoribonuclease assays, was performed as described previously [93]. Essentially this reaction was catalyzed by T7 R N A polymerase (6  U/JJL)  using 0.6  (j.g of linear DNA template (see section 2.8.1). The reaction also included all 4 ribonucleotides with G T P , A T P , and U T P at 0.5 mM, C T P at 50 pM and a - P - C T P at 3 2  375 nM as a label. The reaction contained 0.8 \JI\A. of a commercially available porcine RNase inhibitor (RNasei). The reaction was incubated at 30°C for 2 hrs before being quenched with three reaction volumes of 10 mM EDTA and 2.6 M  26  N H 4 O A C .  Quantification took place by comparison of total 3 2  P radioactivity to total precipitatable  P in 2.5 uL samples. One sample was counted directly, the other after precipitation  with 5% trichloroacetic acid (TCA) in the presence of 10 u.g/mL B S A on ice for 15 min. The precipitated R N A was then filtered under vacuum, washed with 5% T C A , and the filter dried by washing with acetone. The remainder of the reaction was extracted twice with phenol/chloroform/isoamyl alcohol (25:24:1) (PCI) and then precipitated with 2.5 volumes of ethanol. The pellet was washed with 80% ethanol to remove salt. Based on calculated yields, the transcribed R N A was diluted to 0.2 u.M for use in ribonuclease and binding assays. 2.8.3 Monophosphorylation of R N A For substrates where it was desired to have a 5' monophosphorylated R N A product, R N A was prepared as in section 2.8.4 but the pellet was dissolved in 75 uL of Tricine solution. The R N A was passed through a commercially available size exclusion spin column (Amersham-Pharmacia, G-25 spin column) to remove unincorporated deoxynucleotides and short abortive transcripts. R N A is synthesized with a triphosphorylated 5' end; these phosphates were removed enzymatically by 150 U/mL calf intestinal phosphatase (Amersham Biosciences) in the supplied buffer. The phosphatase reaction was incubated at 37°C for 30 min before the enzyme was inactivated by continued incubation at 37°C in the presence of 0.1 % S D S , 5 mM EDTA and 50 u.g/mL proteinase K for 15 min. The protease treatment was stopped by diluting the reaction with one volume of 0.25% S D S and 300 mM sodium acetate. The digest was extracted twice with PCI and then precipitated with 2.5 volumes of 95% ethanol. The precipitated R N A was washed with 80% ethanol, and then dried at ambient temperature and pressure. The phosphorylation reaction was initiated by dissolving the 27  R N A pellet in DEPC-treated water and incubating the R N A for 35 min at 37°C in the presence of 10 mM Tris-acetate pH 7.9, 10 mM Mg-acetate, 50 mM K-acetate, 50 \iM DTT, 500 uM A T P , 0.8 U/uL RNasin (RNase inhibitor, Amersham) and 0.3 U/u.L polynucleotide kinase (PNK). This reaction was quenched by the addition of 2 volumes of 7 mM EDTA, 600 mM N H O A c and extracted once with PCI. The product was then 4  precipitated overnight at -80°C after the addition of 2.5 volumes of ethanol. After centrifugation and decantation the pellet was dissolved in DEPC-treated H 0 , quantified 2  by scintillation counting, and then further diluted in DEPC-treated H 0 to an estimated 2  concentration of 0.2 p M . All transcripts were kept at -20°C in 50 \iL aliquots.  2.8.4 End -labelling of homopolymer R N A Homopolymeric R N A of an undefined length was obtained from Sigma chemical company. Each polymer was separately dissolved in 0.1 M Tris-HCI pH 7.0 and quantified assuming that 1  A 60nm 2  unit is 40 ug/mL R N A and an average polymer length  of 100 nucleotides to estimate the molarity. Each homopolymer was then dissolved to a concentration of 15 uM for use in an end-labelling reaction. The reaction consisted of about 30 pmol R N A , 10 mM Tris-acetate pH 7.9,10 mM Mg-acetate, 50 mM K-acetate, 875 nM A T P , Y - P - A T P as a label and 0.5 UVuL polynucleotide kinase and was 3 2  incubated for 35 min at 37°C. Following the incubation the reactions were chilled on ice and quenched with 160 mM NH -acetate, 8 mM EDTA and the reaction volume was 4  increased to 50 uL with DEPC-treated water. All reactions were extracted once with chloroform/isoamyl alcohol (24:1), speed vacuumed to dryness, and resuspended in 40 uL DEPC-treated water. To rid the reaction of unincorporated nucleotides a G-25 sephadex spin column was used according to the manufacturer's instructions. The volume of the sample was increased to 150 ul_ and aliquots were frozen at -80°C. 28  2.8.5 Exoribonuclease assay Internally  3 2  P labelled S L 9 A R N A was prepared as described in section 2.8.2.  The assay mixture (32 uL) contained 0.8 pmol of R N A in Buffer A. Before addition of the enzyme this mixture was heated for 2 min at 50°C, then at 37°C for 10 min and finally placed on ice for at least 2 min. This heating regime was intended to ensure that the RNA refolded into the expected secondary structure. At time zero 8 uL enzyme was added in a 50 mM phosphate buffer pH 7.5. A zero time point was taken by immediately removing a 4 uL sample and mixing with 12 uL Formamide Loading Buffer. The assay continued at 30°C and 4 u.L samples were removed at various time points, as in the zero timepoint, and mixed with 12 uL Formamide Loading Buffer. Following the assay portions of the 16 u L samples were separated on polyacrylamide gels containing 8 M Urea in T B E buffer.  2.8.6 Endonuclease Assay Internally labelled 9S R N A substrate [93] was prepared as described in section 2.8.2. The assay mixture contained 5 mM Tris pH 7.5, 1 mM MgCI , 20 mM NH CI, 50 2  4  u.M DTT and 12 mM KCI along with 6.7 nM of the prepared labelled R N A . Before addition of the enzyme this mixture was heated for 2 min at 50°C, then at 37°C for 10 min and finally placed on ice for at least 2 min. This heating regime was intended to ensure that the R N A refolded into the expected secondary structure. At time zero 8 ixL enzyme was added to the assay buffer above with glycerol resulting in a final glycerol concentration of 0.05%. A zero time point was taken by immediately taking a 4 u.L sample and mixing with 12 u L Formamide Loading Buffer. The assay continued at 30°C and 4 u.L samples were removed at various time points, as in the zero timepoint, and 29  mixed with 12 \xL Formamide Loading Buffer. Following the assay portions of the 16 \xL samples were separated on polyacrylamide gels containing 8 M Urea in T B E buffer. 2.9 Filter binding assay The filter binding assay used in this study was adapted from an established double-filter method specifically for protein-nucleic acid interactions [94]. The assay utilized a 96 well dot blot apparatus in which samples were placed in separate wells on a 0.45 p,m nitrocelllulose membrane overlaying a nylon membrane. Both membranes had previously been wetted in distilled water for 5 min. A central vacuum source was used to pull 50 \xL of 1x F A B buffer through both membranes for equilibration. Samples were prepared on ice and contained 4 nM labelled R N A and various dilutions of protein in 1x FAB. These samples were added to each well and left at room temperature for 5 min. The vacuum was then again applied to the apparatus for 2 min before each well was washed with 100 pL F A B buffer at room temperature. The vacuum was continued for an additional 5 min before membranes were removed from the apparatus and allowed to dry. Membranes were exposed to a phosphoimager screen which was quantified using Image Quant software. To calculate the fraction of R N A that is bound at equilibrium for each well, the intensity of the signal on the nitrocellulose is divided by the sum of the signal from the nitrocellulose and the nylon membranes (ie. Bound/(Bound + Unbound) = Fraction Bound). Non-specific binding of the R N A or DNA to the nitrocellulose was found to be negligible and therefore not included in the calculation.  30  3 Results 3.1 RNase E S1 domain  3.1.1 Overview: nature of the problem/hypothesis To investigate the function of the S1 domain of RNase E the domain was examined in isolation with a working hypothesis that this domain functions bind to substrate R N A molecules. It was also hypothesized that this domain provides the phosphate binding pocket that is responsible for the increased endonuclease activity of RNase E on monophosphorylated substrates relative to triphosphorylated substrates. This problem was first examined in a structural context to confirm sequence alignments that this domain is indeed an ob-fold. The capacity of this domain to bind R N A was also examined using a filter binding assay. In addition, mutants in the S1 domain, both engineered and a classical-temperature sensitive mutant, were examined for their structural and functional properties.  3.1.2 Expression and Purification The DNA sequence of the E.coli RNase E S1 domain, encoding amino acids 35 125 of RNase E, was cloned into the BamH1 site of the plasmid vector pET15b (see Table 3). The vector, pET15b, contains a T7 polymerase promoter which allows for protein production in strains expressing T7 polymerase under lac operon control. The cloning work was done previously by Mike Cook and the plasmid named S1 B-3.The protein was expressed and purified from BL21 as described in Materials and Methods (section 2.6.2). Steps in the purification procedure were monitored by separating samples from intermediate steps in the procedure on an S D S - P A G E gel (see Materials and Methods, 31  section 2.7.6). Typical results are shown in Figure 4 for the wild type S1 domain of RNase E and Figure 5 for the rne-1 G 6 6 S mutant. In Figure 4A and Figure 5A it can be seen that the majority of pure protein elutes at approximately 50 mM imidiazole. After the thrombin cleavage reaction most of the protein elutes in the Talon column flow through (Figure 4B lane 3 and Figure 5B lane 3). Both purification procedures show a mobility change after dialysis in the presence of thrombin confirming that the hexahistidine tag was cleaved. Samples of purified protein used in subsequent experiments are shown in Figure 4B Lane 4 for the wild type RNase E S1 domain and in Figure 5B Lane 5 for the rne-1 G 6 6 S mutant. A protein contaminant of about 70kDa is visible in the final purified fraction of both preparations. The ams-1 (also called rne-1) mutation is a temperature-sensitive mutation in RNase E that maps to amino acid 66 changing glycine to serine (G66S). This mutation renders cells inviable at the non-permissible temperature and RNase E catalytically inactive at > 37°C. The rne-1 mutation was inserted into the S 1 B - 3 construct using sitedirected mutagenesis (see Materials and Methods, section 2.4) by Mike Cook. Since this temperature-sensitive mutation is responsible for abolishing RNase E activity both in vivo and in vitro, its effect was investigated in the context of the isolated domain.  3.1.3 C D Spectroscopy  '  The stability of the domain was examined using circular dichroism (Materials and Methods, section 2.7.5). This allowed an estimate of the melting temperature (T ) of the m  domain, and a comparison to the rne-1 domain. A s the latter mutation is temperaturesensitive, an altered T was anticipated. It is apparent from the spectra, even at 25°C, m  that the rne-1 mutant domain has a different absorbance profile than wild-type (see Figures 6 and 7). Moreover, both protein samples show altered spectra at 75°C 32  8 200kDa_^ 116kDa 66kDa 45kDa_ 31kDa_  21.5kDa_ 14.4kDa. 6.5kDa.  B 200kDa_J 116kDa_ 66kDa4SkDa  31kDa_|  21.5kDaJ 14.4kDa 6.5kDa  Figure 4: Purification of the S1 domain of RNase E. The S1 domain of RNase E was purified as described in Materials and Methods (section 2.6.2). Various fractions were separated on a 15% SDS-PAGE gel (see Materials and Methods section 2.7.6). A: Lane 1, Molecular size markers; the sizes of each marker are indicated to the left. Lane 2, the S30 fraction; Lane 3, Talon column flow through; Lane 4, 5mM imidazole wash; Lane 5, second 5mM imidazole wash; Lane 6,15mM imidazole wash; Lane 7, 50mM imidazole wash; Lane 8, 500mM imidazole wash. Pooled fractions corresponding to Lane 7 were concentrated and dialysed overnight in the presence of thrombin as described in Materials and Methods (section 2.6.2). B: Lane 1, Molecular size markers; Lane 2, preparation before dialysis in the presence of thrombin; Lane 3, preparation after dialysis in the presence of thrombin; Lane 4, flow through; Lane 5, 500mM imidazole wash. 33  200kOa  3  4  5  6  66kOa 45kDa 31 kDa 21.5kDa  14.4kDa_| 6.5kOa  200kDa. 116kDa. 66kDa-| 45kDa_ 31kDa. 21.5kDa. 14.4kDa. 6.5kDa_l  Figure 5: Purification of the rne-1 S1 domain of RNase E . The rne-1 S1 domain of RNase E was purified as described in Materials and Methods (section 2.6.2). Various fractions were separated on a 15% S D S - P A G E gel (see Materials and Methods section 2.7.6). A : Lane 1, Molecular size markers; the sizes of each marker are indicated to the left; Lane 2, the S30 fraction, Lane 3, Talon column flow-through of S30; Lane 4, 5mM imidazole wash; Lane 5, second 5mM imidazole wash; Lane 6 , 1 5 m M imidazole wash; Lane 7, 50mM imidazole wash; Lane 8, 500mM imidazole wash. Pooled fractions corresponding to Lane 7 were concentrated and dialysed overnight in the presence of thrombin as described in Materials and Methods (section 2.6.2). B: Lane 1, Molecular size markers; Lane 2, preparation after dialysis in the presence of thrombin; Lane 3, Talon column flow through; Lane 4, 500mM imidazole wash; Lane 5, flow-through after concentration.  34  600000  250>  Wavelength (nm)  Figure 6: Circular Dichroism spectra of the RNase E S1 domain. Spectra were recorded on a Jasco J-810 spectropolarimeter with a protein concentration of 35 pM in FAB buffer. Each plotted spectrum is an average of 2 accumulations at the indicated temperature with a resolution of 0.5 °C. Molar ellipticity is reported on the vertical axis in degrees.cm /decimole at 25 °C {•) and at 75 °C (•) plotted against 2  the incident wavelength.  35  200000'! 100000 4  Wavelength (nm)  Figure 7: Circular Dichroism spectra of the RNase E S1 domain containing the rne1 mutation. Spectra were recorded on a Jasco J-810 spectropolarimeter with a protein concentration of 35 uM in FAB buffer. Each plotted spectrum is an average of 2 accumulations at the indicated temperature with a resolution of 0.5 °C. Molar ellipticity is reported on the vertical axis in degrees.cm /decimole at 25 °C (•) and 2  at 80 °C (•) plotted against the incident wavelength.  36  compared to 25°C. A C D melting curve was generated for both the wild type and rne-1 domains using the molar ellipticity at 225nm as an indicator of secondary structure content. Examination of the resultant curves reveal an estimated T for the wild type m  domain of 50°C while the rne-1 mutant has an estimated T  m  at least 5°C lower (Figure  8). These melting curves also reveal that the structure of the rne-1 mutant domain is much different than the wild type domain even at low temperatures. The data indicate that the rne-1 mutation leaves this domain with less secondary structure at physiological temperatures. In addition, the structure is likely in equilibrium between a folded and unfolded state that favours the unfolded state to a greater extent than in the wild type domain. 3.1.4 Protein structural properties Nuclear magnetic resonance (NMR) experiments required specific isotopic labelling of the protein. To achieve this, cells containing S1B-3 were grown in M9 media supplemented with appropriate isotopically labeled substrates. Labelling of the domain enabled use of multidimensional nuclear magnetic resonance (NMR) and resulted in a successful structure determination by Dr. Mario Schubert working with Dr. Lawrence Mcintosh. The structure of this domain was confirmed as a classical ob-fold with 5 stranded anti-parallel B sheets (Figure 9). The B sheets are arranged in a Greek key topology with a 1 -2-3-5-4-1 arrangement. The domain contains extended loops between sheets 3 and 4 that contain alpha helical content. Unexpectedly, crystals formed during the storage of the protein sample used for NMR experiments. These crystals were found to diffract to resolution sufficient for structure determination. Additional protein samples were produced, were crystallized using the initial conditions, and were used by Dr. Mario Schubert and Dr. Paula Lario for 37  -1.50E+05-1.75E+06-2.00E+054*  C  QL  ED i_  eg  o  $£ wild type S1 domain  \  -2.25E+05-J^ -2.50E+05-2.75E+05H  -3.00E+05-I  rne-1 S1 domain ^ ^ s ^ , 3R  -3.25E+05-3.50E+05H -3.75E+05-4.00E+05  — I  1  1  1  1  1  1  1  10 15 20 25 30 35 40 45.50  1  1  1  1  1  1  55 60 65 70 75 80  Temperature fC) Figure 8: Thermal stability of S1 domains as measured by CD spectroscopy. Ellipticity of wild type and rne-1 RNase E S1 domains in pH 6.5 phosphate buffer, each at a concentration of 35 u,M was measured at 225nm over the temperature range indicated (see Materials and Methods, section 2.7.5). Molar ellipticity is reported on the vertical axis in degrees.cm /decimole. 2  38  Figure 9: The peptide backbone solution structure of the S1 domain of RNase E as determined by Dr. Mario Schubert and Dr. Lawrence Mcintosh by NMR. This is a superposition of 7 different best fit structures with |3-sheets shown as flat bands ending in arrows pointing in the carboxy terminal direction and a helices as flat bands without arrows. Structures with the least defined structure which show the most variation between possible structures and which are therefore considered to be the most dynamic are indicated only by the grey lines .  39  the determination of the x-ray crystal structure of this domain (Figure 10). The native and Pb-derivative diffracted to 1.8 A and 2.0 A resolution, respectively. The Rwork / Rfree for the determined structure is 0.189. This structure provides a structure with greater resolution and provides some additional evidence that the S1 domain of RNase E may dimerize specifically. 3.1.5 Dimerization Interestingly the asymmetric unit of the crystals of the RNase E S1 domain included two monomers (Dr. Schubert, personal communication). Moreover, N O E ' s (Nuclear Overhauser Effect) from the N M R structure were consistent with intermolecular interactions between monomers. These interactions lie on a surface of the domain separate from the putative R N A interaction sites located on (3-1 and (3-4. NOE's and concentration-dependent chemical shifts implicated residues Y42, I46, E76, E99, V100, and Q103 (Dr. Mario Schubert, personal communication). These observations suggest that the RNase E S1 domain may dimerize specifically. In support of this notion, the NMR tumbling time is concentration dependent, 7.4 ns at 0.6 mM and 8.3 ns at 1.2 mM, and is somewhat slow for a 10kDa monomer [95]. In contrast, static light scattering suggested that purified RNase E S1 domain is monomeric (data not shown). To investigate the possibility that the S1 domain of RNase E multimerizes in solution, crosslinking experiments were performed with glutaraldehyde (see Materials and Methods, section 2.7.8). The gels in Figure 11 shows that prior treatment of either the wild type or rne-1 RNase E S1 domains produces additional bands around 26kDa in lanes 8 and 9 denoted by the arrow in the margin. Visual inspection suggests that as much as 10 - 1 5 % of the protein was converted to the putative dimer under the  40  Figure 10: The crystal structure of the S1 domain of R N a s e E. A : The structure with labelled p-sheets and loops. The N terminus of the protein is the end nearest to psheet labelled p i . For the labels on the figure p is the prefix for p-sheets and L is the prefix for the loops in between the sheets. A n axial view of the p-barrel as seen from the top of A. This figure and the data used to produce it are the work of Dr. Mario Schubert.  41  7  8  9  10 11 12  200kDa-^ 116kDa. 66kDa. 45kDa. 31kDa_ 21.5kDa_ 14.4kDa_ 6.5kDa_  Figure 11: Crosslinking various S1 domain preparations. Samples of the RNase E S1 domain and its rne-1 counterpart were crosslinked in 0.01% glutaraldehyde (see section 2.7.8). Samples containing 2.25 nmol were separated on a 15% S D S - P A G E gel and stained (see Materials and Methods, section 2.7.6). Lanes 1 and 7 contain standard molecular weight markers; lanes 2 - 6 contain the various preparations without glutaraldehyde; lanes 8 - 1 2 are the same preparations crosslinked with glutaraldehyde. Lanes 2 and 8 contain the wild type R N a s e E S1 domain; lanes 3 and 9 contain the me-1  S1 domain; lanes 4 and 10 contain the S1 domain of  PNPase; lanes 5 and 11 contain a 1:1 mixture of wild type R N a s e E S1 domain and the S1 domain of P N P a s e ; and lanes 6 and 12 contain a 1:1 mixture of me-1  S1  domain and the S1 domain of P N P a s e . The arrow (*-) in the right margin denotes the position of the putative S1 dimer.  42  conditions tested. Comparable treatment of the P N P a s e S1 domain results in a much weaker band, showing that cross-linking efficiency depends on the particular S1 domain examined, preparation around 28kDa in lane 10. Lanes 11 and 12 of figure 11 show similar bands as those in lanes 8 and 9 suggesting that heterologous crosslinking is inefficient in mixtures of S1 domains from RNase E and P N P a s e . Thereby demonstrating the specificity of cross -linking of the S1 domain of RNase E. 3.1.6 R N A binding studies  3.1.6.1 Filter binding studies To determine the binding properties of the RNase E S1 domain, a two membrane filter binding assay protocol was used to permit quick analysis of many samples using a 96 well dot blot apparatus (see Materials and Methods, section 2.9). This method facilitated the examination of various factors that are involved in binding of this domain to RNase E substrates as well as the specificity of this interaction both with regards to substrate and to particular residues on the domain. In these experiments the K is defined as the concentration of one component at d  which 50% of the other component of the interaction is bound. For these experiments the R N A concentration is limiting (4 nM) and the protein concentration is varied above and below the K (see Materials and Methods, section 2.9). d  The value of the K is a reflection of the strength of the interaction between the d  two components, the higher this value the weaker and more transient the interaction. Thus differences in the dissociation constant provide a way of quantifying the various contributions of different factors involved in the binding interaction. Factors whose role  43  in binding were investigated include the 5' phosphorylation state of the R N A , the sequence of the R N A molecule, pH, salt concentration and magnesium concentration.  3.1.6.2 pH effects on R N A binding Based on its sequence, the pi of the RNase E S1 domain is calculated to be 8.2. This relatively high pi indicates that the positive charge at physiological pH of this domain could offer strong contributions to binding the phosphate backbone of R N A . In order to examine the contribution of ionizable side chains to the interaction, the filter binding assay was performed in F A B buffer at varying pH (see Figure 12). The plot of Figure 9 shows a distinct dependence of binding on solvent pH. These data indicate that charge interactions are essential for binding because as the pH approaches the pi of the domain where the net charge is neutralized, the interaction is eliminated.  3.1.6.3 Ionic effects on R N A binding A s an alternative method to probe the importance of change interactions between the S1 domain and an R N A ligand, the filter assay was performed in the presence of varying KCI concentrations. Potassium and chloride ions in the binding buffer are able to neutralize charges on the protein and on the R N A phosphate backbone, thereby competing for ligand binding. Additionally, by increasing the ionic strength of the buffer, these ions favour hydrophobic interactions. Thus this assay enables a rough method of determining which forces dominate the interaction of the two molecules. Results of such an experiment are graphed in Figure 13. These data indicate that increasing the ionic strength of the solvent with KCI decreases the affinity of the RNase E S1 domain for the substrate R N A . Thus, data from filter binding  44  1 -I  pH Figure 12: Effect of pH on RNA binding by the RNase E S1 domain. Filter binding assays were performed and the fraction bound was calculated as described in Materials and Methods (section 2.9). These assays were done in sodium phosphate buffer at various pH with 6 jiM RNase E S1 domain. The RNA substrate is 4 nM triphosphorylated SL9A (Materials and Methods, section 2.8.2). Each point represents an average of two experiments.  45  Figure 13: Effects of varying KCI concentration on the binding of S L 9 A by the RNase E S1 domain. Filter binding assays were performed as described in Materials and Methods, section 2.9. The protein concentration was maintained at 6 p,M while the potassium chloride concentration was varied from 20 mM to 200 mM. The Fraction Bound is plotted against the KCI concentration with standard error bars and the solid line connecting each point is an approximation of a best fit line.  46  experiments performed with varying pH or KCI concentration confirm the importance of ionic interactions for binding of R N A to this domain.  3.1.6.4 Effects of M g  + +  on R N A binding  The impact of divalent cation in the binding interaction was also investigated. RNase E activity requires the presence of divalent metal ions [38]; therefore, I tested whether the binding of the S1 domain to substrate might be the source of this requirement. Figure 14 shows the fraction of bound R N A is plotted versus Mg ion concentration in a standard filter binding assay. Altering the magnesium chloride concentration in the binding assay does not change the fraction bound significantly over a range of [Mg ] between 0 - 4mM. Since the examined interaction does not require ++  the presence of magnesium ions, binding of R N A by the S1 domain is not the source of the requirement of magnesium ions for the catalytic activity of RNase E.  3.1.6.5 Monophosphorylation and triphosphorylation effects on R N A binding If the S1 domain of RNase E facilitates higher enzymatic activity on monophosphorylated substrates, then the domain may have a lower Kd for such R N A s . This was tested in a filter binding assay in which the binding of S L 9 A R N A containing a 5' -monophosphate was compared to the same R N A molecule initiated with a 5' triphosphate. The synthesis of these R N A molecules is described in Materials and Methods (sections 2.8.2 and 2.8.3). Monophosphorylated SL9A (•) and triphosphorylated S L 9 A (•) each exhibited apparent K s of approximately 4 x 10" M 6  d  (Figure 15). These results indicate that the domain exhibits no preference for monophosphorylated RNA. Moreover, the S1 domain also displays very similar K for d  47  1 0.90.8-  1  0.70.6  H  c 0.5 o 0.4  0.3 0.2 0.1 ^ 0  -i  2  3  4  5  MgCI {mM) 2  Figure 14: Effects of Mg ion concentration on RNA binding to the RNase E S1 domain. The fraction of bound RNA was determined by the filter binding assay described in Materials and Methods (section 2.9). The concentration of RNase E S1 domain was held constant at 6 mM, the concentration of RNA at 4 nM, while the concentration of magnesium chloride was varied as indicated from 0-4 mM. Data points are an average of two experiments.  48  O.OOE+00 2.00E-06 4.00E-08 6.00E-06 8.00E-06 1.00E-05 1.20E-05 1.40E-05  Protein Concentration (M) Figure 15: Binding of monophosphorylated SL9A, triphosphorylated SL9Aand monophosphorylated SL9 by the RNase E S1 domain. The concentration of RNA substrates in the filter binding assay was 4 nM. The structure of the RNA is described by Spickler and Mackie (2000) [92] and its preparation is described in section 2.8.2. The concentration of the RNase E S1 domain protein is plotted against the average fraction of bound RNA from three replicate experiments. The fraction bound was calculated as described in Materials and Methods (section 2.9). A line connects each data point; a solid line for triphosphorylated SL9A; a dotted line for monophosphorylated SL9A and a dashed line for monophosphorylated SL9.  49  monophosphorylated S L 9 U ) R N A , indicating that the 30 adenylate residues at the 3' end of the molecule do not impact the interaction (Figure 12). To examine the possibility that the S1 domain of RNase E exhibits some sequence preference, the affinity of the domain for different homopolymeric R N A s was examined. The results plotted in Figure 16 show that both poly (U) (•) and poly (A) (•) exhibit apparent K values of approximately 2 x10" M. Moveover, the binding curves are 6  d  virtually superimposable. These results suggest that the RNase E S1 domain does not display any specificity for one homopolymer relative to the other. 3.1.6.6 Effects of the temperature-sensitive mutation, G 6 6 S , on R N A binding. To investigate the effect of this mutation on R N A binding, filter binding assays were performed to compare the wild type to the rne-1 mutant domain. The data in Figure 17 show that the wild type S1 domain exhibits an apparent K for S L 9 A R N A of 5 d  x10" M, slightly higher than measured in Figure 15. The me-1 mutant domain exhibits a 6  slightly lower apparent K of approximately 2 x10" M. A similar assay was performed 6  d  using end-labelled poly (C) at different temperatures to check the R N A binding capacity of the rne-1 mutant domain. A s can be seen from Figure 18 the binding capacity of the mutant domain for poly C R N A appears largely unaffected by temperature. The wild type domain appears to display limited binding, especially at higher temperature (Figure 18, row 3 and row 5). However, as noted in section 3.1.1.1 the preparation of this domain contains RNase activity. At the higher concentrations and higher temperatures this activity may be increased resulting in degradation of the R N A and a loss of signal on both membranes.  50  1.20  0.00 •+= » O.OOE+00 5.00E-06  1  1  1  1  1.00E-05  1.50E-05  2.00E-05  2.50E-05  > 3.00E-05  i 3.50E-05  Protein Concentration (M)  Figure 16: Binding of end-labelled home-polymeric RNA to the RNase E S1 domain. The fraction bound was determined for each protein concentration by dot blot filter binding assay as described in Materials and Methods, section 2.9. The end-labelled homopolymeric RNA, at a concentration of 4 nM, was prepared as described in Materials and Methods, section 2.8.4. The solid line (poly (A)) and dashed line (poly (U)) are approximations of a best fit line.  51  0  0.000005 0.00001 0.000015 0.00002 0.000025 0.00003 0.000035 Protein Concentration (M)  Figure 17: Binding monophosphorylated SL9A RNA to wild type (solid line) or me1 (dashed line) S1 domains. SL9ARNA, at4nM in binding mixture, was prepared as described in 2.8.2. Protein concentrations are plotted against the fraction of RNA bound (see Materials and Methods, section 2.9). This data is representative of three aliquots of the same dilution in three different wells of the 96-well plate for each spot. The fraction bound is an average of the signals with standard error shown as the Y-axis error bar and lines connecting data points as an approximation of best fit lines.  52  Protein Concentration 0 1  +  2  • •  3  Nitrocellulose membrane  %  4 5 6 7 8  • •  10  %  11  Hi  •  w  # •  *  9  12  1  #  • •  •••  •  •  • 27 13.5 6.8  3.4  1.7  .8  .2  .4  Nylon membrane  in w  .1  Protein Concentration (jiM)  Figure 18: Binding of poly (C) to wild type or me-1 R N a s e E S1 domain. The nitrocellulose, retaining bound RNA, and nylon membranes, retaining free R N A are shown. Filter binding was performed as described in the Materials and Methods (section 2.9). Odd numbered lanes contain a dilution series of the wildtype S1 domain while even numbered lanes are a dilution series of me-1 S1 domains from 27yJMI to 0.1 u.M as indicated below the nylon membrane blot. The first column is a control containing no protein. Lanes 1 and 2, 7 and 8 of the experiment were done at room temperature; lanes 3 and 4, 9 and 10 were incubated first at 44 °C for 10 min and then transfered to ice for 10min before being applied to the well; lanes 5 and 6,11 and 12 were incubated at 44 °C for 10m in and then directly applied to the well.  53  3.1.7 Competition The hypothesis that the S1 domain of RNase E provides preference for 5' monophosphorylated over 5' triphosphorylated substrates provided another testable property of the domain. First, the activity of RNase E should be hampered in the presence of a domain that binds to substrate molecules leaving them protected from endonuclease activity. This effect would be exacerbated if the domain binds to the same substrate recognition sites as the complete enzyme. Also, if indeed the S1 domain provides specificity by preferentially binding R N A molecules at the 5' -end, then in the presence of excess autonomous S1 domain the end-dependent activity of RNase E may be reduced. The results of such a mixing experiment are shown in Figure 19. Full length 9S RNA, labelled S, is initially endonucleolytically cleaved by RNase E at one of two sites to produce two alternative products, 8 S and 7S (Figure 19, panel A). A second cleavage produces pre-5S R N A , labelled p5S. If the S1 domain mediates the 5' -end dependence of RNase E through stronger binding to monophosphorylated 9S R N A , then 8S and 7S intermediates would accumulate in the presence of competing free S1 domain. The S1 domain preparation alone displays little or no endonuclease activity. Panel C shows the effect of adding a 1, 000-fold excess of the S1 domain to the assay. The rate of formation of p5S product is not significantly different from that observed in panel A. The intensity of the p5S band in panels A and C of Figure 19 after 30 minutes are approximately equivalent. These results clearly show that even in the presence of a large excess of the S1 domain RNase E activity is unaffected.  3.1.8 DNA titration NMR offers a powerful tool to examine the dynamics of the interaction between the S1 domain and an R N A ligand. Knowledge of the structure makes possible 54  Time (min) 0  5  10  30  60  0  5  10  30  60  0  5  10  30  60  •95  •as -7S  -pSS  5'end  Panel A  Panel B  Panel C  Figure 19: Time course of RNase E activity on 6.7 nM internally labelled 9S RNA transcript at 30 °C see Materials and Methods (section 2.8.6). Panel A: 2.8 nM of RNase E; Panel B; 2.8 uM S1 domain; Panel C: 2.8 nM RNase E in the presence of 2.8 uM S1 domain. (Abrevations: 9S = 9S RNA substrate, 8S = product of an initial cleavage, 7S = product of an alternative initial cleavage, p5S = pre-5S RNA product of both cleavages, 5' end = product of cleavage producing 7S)  55  monitoring of changes in chemical shift of specific amide backbone residues of the protein in the presence of ligand. The chemical shift of individual amide hydrogens and nitrogens reflects the environment of the residue and can be used as a sensitive indicator of changes in this environment. Therefore, an experiment was designed in which a model ligand, BR10 DNA, was titrated into a solution of the RNase E S1 domain and the chemical shifts were monitored by NMR. Residues that show a change in chemical shift during the titration are changing in position or interacting with DNA. The ligand used for the NMR titration was a DNA mimic of a known R N A substrate (see Materials and Methods, section 2.7.2). It was observed that degradation of R N A occurred under the conditions of the NMR experiment due to small amounts of contaminating exonuclease activity that could not be removed from the protein preparation. Several additional chromatographic steps were added to the procedure in Materials and Methods (section 2.6.2) in an attempt to eliminate this activity. This is examined in more detail in section 3.1.11. Data obtained from the N M R titration were analyzed by Dr. Mario Shubert and revealed several chemical shifts consistent with a binding interaction. These chemical shifts implicate certain residues that are potentially directly or indirectly involved in the binding of BR10 DNA. These residues and an indication of their relative change are illustrated in Figure 20 (prepared by Dr. Mario Schubert). A total of 15 residues show altered chemicals shifts on a two dimensional H N N M R spectrum. The amino acids 1  1 5  are numbered as they would be in the full length RNase E protein. Several positively charged amino acid side chains are associated with these changes: R64, K71, K106, R109, and K112. Aromatic residues that appear to be solvent exposed in the NMR structure and show a chemical shift perturbation in the titration include F57, Y60 and  56  chemical shift perturbation  Figure 20: NMR-derived space filling model of the S1 domain of RNase E highlighting residues which show chemical shift changes upon binding to DNA. The relative change in chemical shift is indicated on the model by the shading. Those residues showing significant chemical shift perturbations upon the addition of DNA to the domain are labelled. The opposite side is shown as a 90 clockwise rotation about a vertical axis. This figure was provided by Dr. Mario Schubert.  57  F67. All of the chemical shift changes are localized to a single face of the domain indicating that only one side of the protein is involved in interaction with DNA, and presumably with R N A also.  3.1.9 Design of RNase E S1 domain mutants  3.1.9.1 S1 domain alignment An alignment of the S1 domains of three enzymes of mRNA degradation in E.coli shows many regions of conservation (Figure 21). It is evident from the alignment that there is more conservation between the two endonucleases, RNase E and RNase G , than between RNase E or RNase G and P N P a s e . This may reflect the fact that RNase E and RNase G are both 5' -end-dependent endonucleases so that residues that may be important for RNase E function therefore would be more likely to be conserved in RNase G than P N P a s e . 3.1.9.2 Analysis of structure in terms of potential interaction residues. A visual inspection of the structure presented in Figure 20, reveals the position of solvent exposed residues that are sterically more available for binding ligand. While residues that show a chemical shift such as A115 and L116 may provide hydrophobic pockets for the exclusion of water and interaction with bases on R N A , they also may show a chemical shift change simply from a conformational change in the backbone of the protein upon binding. Residues such as E54, D105 and E107, which also show significant chemical shift perturbation, may provide hydrogen bonding opportunities for nucleic acid bases. These residues may be providing the domain with specificity through particular hydrogen bonds to certain bases of the ligand. Therefore, a shift upon binding due to direct interaction with nucleic acid most likely implicates residues that 58  75 80 85 90 K J B J R E Y FP A N Y S A H G R P N D IE P H T E C V A G E E Q K Q F T EDK R  115  120  125  rne rng pnp  Figure 21: Alignment of three S1 domains from enzymes involved in m R N A degradation in E.coli. RNase E, RNase G and P N P a s e are labelled rne, rng and pnp respectively. The numbering system is based on full length R N a s e E. The alignment was calculated using Clustal W (http://clustalw.genome.ad.jp/) using a gap open penalty of 10 and gap extension penalty 0.05 and using the B L O S U M (for protein) scoring matrix. Amino acids that are conserved are highlighted in black while those only residues conserved between two domains are highlighted in grey.  59  protrude into the solvent and have side chains which participate in interactions with nucleic acid. Residues that showed conservation (Figure 21), a chemical shift change (Figure 20), and possessed structure and side chain properties suitable for binding DNA or R N A were designated as targets for site-directed mutagenesis, F57, R80 and R109 in particular. Phenylalanine residues can undergo hydrophobic interactions and stack with nucleotide bases in an oligonucleotide binding protein or domain [16]. Arginine residues can provide ionic interactions with the phosphate groups of the polyribonucleotide ligand to facilate binding of a protein or domain. Mutating these residues to alanine would eliminate these properties. The site-directed mutagenesis and analysis of the resultant polypeptides is described in section 3.1.10.  3.1.10 Mutant S1 domain properties Mutant RNase E S1 domains were made using the same construct produced by Mike Cook as template (Materials and Methods). Mutants were designated by the original amino acid followed by the number of this amino acid in the full length RNase E protein sequence, followed by the amino acid replacing it in the one letter code. The mutant plasmids were sequenced and confirmed as F57A and R109A; however, the R35A mutant contained an additional unintentional mutation, E34D. Nonetheless, it was subjected to the same studies as other mutants. The mutant proteins exhibited no significant difference in solubility or concentration during their preparation. The wild type S1 domain of 10, 632Da contains 14 basic residues, and 13 acidic residues resulting in a predicted pi of 8.2. While the F57A mutation had little impact on the predicted pi of the domain, the R109A and R64A + E63D mutations altered the predicted pi based on sequence from 8.2 to 7.04.  60  The isolated mutant domains, R109A and F57A, were subjected to filter binding assays to examine the contributions of these residues to the binding interaction. A s can be seen from the filter binding results with these mutant domains (Figure 22), there is no detectable difference in R N A binding under the conditions tested. The K ' s for both d  mutant domains are approximately 1 x10" which is very close to that for the wild type 6  domain (Figure 16), within error of the experiment. Moreover, neither mutant domain showed a detectable difference in K for poly A compared to poly U. d  3.1.11 RNase contamination As noted in section 3.1.8, RNase activity was found in preparations of the S1 domain of RNase E. Panel A of Figure 23 shows that poly C R N A is almost completely converted to low molecular weight products after a 10 minute incubation with the wild type S1 domain at 30°C. This activity complicates assays of R N A binding and eliminated the possibility of using R N A in NMR experiments. Alternative purification techniques were utilized in an attempt to rid the preparation of the nuclease activity, including reverse phase high performance liquid chromatography (see Materials and Methods, section 2.6.2). A single, apparently homogeneous peak of an UV absorbance at 280 nm eluted from this column (data not shown); however, the resulting purified domain still contained a level of nuclease activity similar to the starting material. This result suggested that the S1 domain itself could have intrinsic nuclease activity. To test this hypothesis, the activity of the wild-type protein preparation, was compared to that of three mutants (Figure 23). After 10 min at 30°C the wild-type RNase E S1 domain has degraded the poly (C) R N A to small oligonucleotides and nucleotides whereas longer R N A persists longer in the reactions with rne-1, F57A and R109A mutants (Figure 23, Panels B, C, D respectively). Although not a quantitative assay, it can be seen that the 61  1• A  0.90.8 • 0.7 •  X X A  •  0.60^ 0.40.30.2 • 0.1 • 0-  I  I  I  I  0.000002 0.000004 0.000006 0.000008 0.00001 0.000012 0.000014 Protein Concentration (M)  Figure 22: RNA binding by RNase E S1 domain mutants, F57Aand R109A. Each protein was prepared as described in Materials and Methods (section 2.6.2) and tested for filter binding with 4 nM of end-labelled poly (A) or poly (U) as indicated. The mutant F57Awas tested with poly (A) (•) and poly (U) (>), similarly R109Awas tested with poly (A) {*) and poly U (x). The fraction bound was calculated described in Materials and Methods section 2.9. For clarity, best fit curves are omitted.  62  Time (min) 0  10  30  lis  60 120  3jg^.  • 0  10  30  60 120  0  10  30  60 120  0  10  30  60  120  ft  >«Hh.  wt RNase E S1  me-1 RNase E S1  F57A RNase E S1  R109A RNase E S1  Figure 23: Nuclease activity of various RNase E S1 domain protein preparations. 5' end labelled poly C (5 nM) was incubated with the indicated RNase E S1 domain (25 uM) for the indicated time at 30 °C. The reactions were quenched in formamide buffer and separated by electrophoresis on 8% TBE UREA polyacrylamide gels. S indicates the signal from the largest species in the gel while the P indicates the signal from the smallest fragments in the gel. (Abbreviations: S = substrate poly C RNA, P = product oligo C RNA).  63  rne-1 S1 domain contains the least amount of RNase activity followed closely by the F57A S1 domain mutant. It could therefore, be argued that the S1 domain exhibits nuclease activity in the absence of the full length enzyme and mutations in the domain interrupt this activity. To investigate this further a western blot of two S1 domain preparations was used to identify potential contaminants. Both preparations contain some RNase G , shown by the arrow, but no detectable RNase E (Figure 24, top and bottom panels). A s can be seen from Figure 24, middle panel, the wild-type S1 domain contains higher amounts of contaminating full length P N P a s e than does the rne-1 construct. The slightly faster migrating bands are likely products of proteolysis of P N P a s e corresponding to the elimination of the KH and S1 domains. These domains have been shown to be sensitive to proteolysis previously and are detected by the P N P a s e polyclonal antibody [84] (Janet Hankins, personal communication). This would be consistent with the observed lower nuclease activity of the rne-1 construct, if full length P N P a s e is the source of the RNase activity. The presence of 20mM phosphate buffer at pH 6.5 in all the preparations provides the necessary reaction conditions for phosphorylysis catalyzed by P N P a s e . Therefore, it is likely that at least some of the nuclease activity of the preparation is due to the contamination by this enzyme. It has not been shown conclusively, however, that the S1 domain of RNase E lacks intrinsic exonuclease activity independent of P N P a s e contamination or that another contaminating RNase enzyme is responsible for this activity.  3.2 PNPase S1 domain To investigate the role of the S1 domain of P N P a s e in mRNA degradation in E.coli, a P N P a s e construct was made with a deletion in this domain. At the outset, it 64  RNase G  k~ P N P a s e  1  9  8  k  D  a  H  k RNase E  —  Figure 24: Western blots of RNase E S1 domains. Lane 1 contains a whole cell extract (courtesy of Dr. George Mackie); lane 2 contains the wild type RNase E S1 domain; lane 3 contains the rne-1 R N a s e E S1 domain. Each blot was probed with the indicated polyclonal antibodies and visualized using a secondary antibody conjugated to horse radish peroxidase (see Materials and Methods section 2.7.7). Molecular weight markers migrated as indicated to the left of each blot.  65  was essential to determine whether an N terminally His-tagged P N P a s e would serve as a good model. This type of construct has been used before but has lacked activity in our hands [96]. It was found that the His-tagged P N P a s e , while active, had different properties under the conditions tested (section 3.2.1). An untagged P N P a s e lacking the S1 domain and a construct lacking the K H domain were assayed for activity to test the hypothesis that these putative R N A binding domains contribute to the processivity of P N P a s e (section 3.2.2). Additionally the P N P a s e lacking the S1 domain was tested for its ability to rescue the cold-sensitive phenotype of a P N P a s e null mutant (section 3.2.3). 3.2.1 Activity of His-tagged and untagged P N P a s e . Untagged P N P a s e was expressed and purified from BL21 harbouring the plasmid pGC400, while the construct P E P a 1 8 in BL21 was used for expression and purification of His-tagged P N P a s e (see Materials and Methods, section 2.6.1). While both preparations of P N P a s e showed phosphorylytic activity dependent on the presence of phosphate, there were detectable differences in the sizes of products produced from reactions with internally labelled rpsT (268 - 447) poly A  3 0  mRNA (Figure  25 compare lanes 6 - 1 0 with 16 - 20). The data show that the native enzyme is more active than its tagged counterpart. The digestion catalyzed by wild type (lanes 6 -10) is complete within 5 minutes whereas His-tagged P N P a s e requires 30 - 60 min to digest all the substrate (Figure 25, lanes 16 - 20). Moreover, while wild type P N P a s e produces the product labelled P, the His-tagged protein produces an extended R N A product band labelled P' in addition to small amounts of P. The source of this difference is unknown as is the identity of the extended product. Accordingly, it was concluded that wild type P N P a s e is a more useful in vitro model than the tagged enzyme. 66  Time (minutes) B  5  19 30 60  0 5  10 3060  0 m m m m  1  2  3  4  PNPase -P04  5  0  5  A  A  ^Kjpp  6  7 8  9 10  10 30 60  *  flmpp  A  10  3060  A 'HHP  11 12 13 14 15  PNPase +P04  0 5  His-tagged PNPase -PO4  16 17 18 19 20 His-tagged PNPase +PO4  Figure 25: Rate of phosphorolysis of wild type and his-tagged P N P a s e . Reaction mixtures contained 0.8 pmol of internally labelled rpsT poly A mRNA, 0.8 pmol of P N P a s e and were incubated with and without 10 mM phosphate buffer pH 7.5 as described in Materials and Methods (section 2.8.5). A representative experiment is shown where either native or hexahistidine tagged P N P a s e was the source of activity. (Abreviations: S = rpsT/poly A substrate, P = phosphorylysis product, P' = elongated product (see text)).  67  3.2.2 Processivity To investigate the role of the S1 domain in the processivity of P N P a s e , two different deletion constructs were made (Figure 26). The P N P a s e A K H mutant eliminates the entire KH domain, while the P N P a s e AS1 deletion eliminates the first three p-sheets of the P N P a s e S1 domain. These deletion mutants have been previously designed and partially characterized by Jarrige et al. 2002 [83]. To recreate these mutants, site-directed mutagenesis was employed using primers RE16-M1, RE16-M2 and RE17-M1, RE17-M2 to generate P N P a s e AS1 and P N P a s e A K H , respectively (Table 2, section 2.4). Neither construct encoded a hexahistidine-tag. The resultant proteins were overexpressed and purified using a three step chromatographic procedure as described in Materials and Methods (section 2.6.1). The model substrate SL9A, described previously [92], was used to assay phosphorolysis activity of the P N P a s e AS1 and P N P a s e A K H enzymes compared to the wild type P N P a s e . A s can be seen from Figure 27 there are very significant differences in the specific activity of these different deletions. The P N P a s e AS1 (Figure 27, Panel B) can be estimated from the time course to be at least 100 -fold less active than the wild type (Figure 27, Panel A). The P N P a s e A K H mutant (Figure 27, Panel C) can be estimated to be about 20 -fold less active than the wild type enzyme shown in Panel A. No intermediates could be detected between the substrate (S) and product (P) bands even when similar samples were separated at higher resolution on a sequencing gel (data not shown). The absence of detectable intermediates indicates that all three exhibit processive phosphorylytic activity. In the cell P N P a s e is under conditions that favour phosphorolysis; however, in vitro conditions can be designed such that R N A polymerization occurs. Both the 68  Wild type PNPase  PH'  •  PH  I I  KH  S1  II I I  0-210  312-541  PH'  PH  II  551-591622-690  PNPase A K H  yA  III  S1  C Z I  563-619 PNPase  AS1  i  PH'  PH  i  i  KH  /  ~n~rf 625-658  Figure 26: Schematic of PNPase wildtype and deletion mutants. Both deletions were created using site-directed mutatgenesis and confirmed by DNA sequence data (see Materials and Methods, section 2.4). The numbers in the constructs labelled PNPase AKH and PNPase AS1 indicate those residues that have been eliminated. All constructs were made in the vector pET11.  69  Time (min) 0  1  5  10 20 30  45  60 90  B  Figure 27: Rate of phosphorolysis of internally labelled S L 9 A RNA. Reaction mixtures contained 0.8 pmol RNA, 10mM phosphate buffer pH 7.5 and were incubated at 30 °C as described in Materials and Methods (section 2.8.5). A representative experiment is shown using 0.65 u.M wild-type P N P a s e (Panel A), 22 pjvl PNPase-S1 (Panel B) and 4.5 uM P N P a s e - K H (Panel C). (Abreviations: S = S L 9 A substrate, P = phosphorylysis product).  70  phosphorylysis and polymerization activities were tested for processivity using the mutant P N P a s e A S 1 . These results are shown in Figure 28. Panel A shows no detectable intermediates. Unfortunately under these conditions the reaction occurs too quickly to conclude that no intermediates would be detected. It is clear, nonetheless, that the S1 domain of P N P a s e is not necessary for either activity of P N P a s e under the conditions tested.  3.2.3 Cold-sensitive phenotype of pnp mutants. As the S1 domain closely resembles a cold shock domain and a P N P a s e deletion strain of E.coli shows a cold-sensitive phenotype, the P N P a s e AS1 plasmid was tested for its ability to rescue this phenotype. Figure 29a - c shows that a P N P a s e deletion strain (pnp::Tn5) is viable at 37°C. When this strain contains a plasmid with an antisense insert it does not show growth when cultured at 16°C (Figure 26 e). A wild type P N P a s e construct can rescue the cold-sensitive phenotype of this strain and the culture is viable at low temperature (Figure 29 d). The P N P a s e AS1 construct cannot rescue growth of the mutant and exhibits the same phenotype as the negative control (Figure 26 f).  3.2.4 Filter binding assay The S1 domain of P N P a s e (corresponding to residues 617 - 700) was overexpressed and purified as described in section 2.6.2. Results from the purification procedure were followed by separating samples from intermediate steps in the procedure on an S D S - P A G E gel (see Materials and Methods, section 2.7.6); (Figure 30). The purified protein is shown in Figure 30 B Lane 2 and appears free of detectable contaminants. The smear in this lane is thought to be from overloading the gel.  71  Time (min) 0  5  10  -wpps-  111  30  60  0  5  10  30  60  M—E  •man  <«-s P  Panel A  Panel B  Figure 28: Rate of phosphorylysis and homopolymer tailing by P N P a s e AS1. Reaction mixtures contained 0.8 pmol of internally labelled rpsT poly A m R N A and 2.3 u M P N P a s e - S 1 incubated in the presence of 3 m M A D P (Panel A) or 12 m M sodium phosphate buffer pH 7.0 (Panel B). (Abbreviations used: S = substrate, E = extended R N A , P = phosphorylsis product.)  72  c: PNPase AS1  f. PNPase AS1  LB agar at 37 degrees  LB agar at 16 degrees  Figure 29: Rescue of the cold-sensitive phenotype of P N P a s e null mutants by wildtype or PNPase-S1 constructs. E. coli pnp::Tn5 transformed with plasmids pGC400, containing full length PNPase (a, d), pRE75 (PNPase AS1) (c, f) or pRE74 (PNPase AS1 ligated in the vector in the opposite orientation) (b, e) were plated on LB agar containing 100ug/mL ampicillin. Plates were incubated at 37 and 16 degrees.  73  200kDa_j 116kDaIJ  200kDa. 116kDa. 66kDa_| 45kDa_  31kDa.  21.5kDa.  Figure 30: Purification of the S1 domain of P N P a s e . The S1 domain of P N P a s e was purified as described in Materials and Methods (section 2.6.2). Various fractions were separated on a 15% S D S - P A G E gel (see Materials and Methods section 2.7.6). A: Lane 1, Molecular size markers (sizes of each marker are indicated to the left); Lane 2, the S30 fraction; Lane 3, Talon column flow through of the S30; Lane 4, 5 mM imidazole wash; Lane 5, second 5 mM imidazole wash; Lane 6 , 1 5 m M imidazole wash; Lane 7, 50 mM imidazole wash; Lane 8, 500 m M imidazole wash. The fraction corresponding to Lane 7 was concentrated and dialysed overnight in the presence of thrombin as described in Materials and Methods (section 2.6.2). B: Fractions after thrombin treatment Lane 1, Molecular size markers; Lane 2, concentrated flow-through; Lane 3, flow-through Lane 4; 500 mM imidazole wash. 74  The ability of the purified P N P a s e S1 domain to bind R N A was also examined using a filter binding assay. Both S L 9 A and S L 9 R N A s were tested in either monophosphorylated or triphosphorylated form (Materials and Methods, section 2.9). The K of the P N P a s e S1 domain was estimated to be over 200 u.M d  (data not shown). Thus, this domain exhibited a much higher K than the S1 d  domain of RNase E.  75  4 Discussion This thesis has addressed many questions about the function of S1 domains in two key enzymes of mRNA degradation in E.coli. The importance of this domain is evident from its presence in multiple enzymes involved in this fundamental process (see Figure 2). Two basic hypotheses were tested in this thesis. It was thought that these domains served different functions in RNase E and P N P a s e through their ability to bind RNA. In RNase E the S1 domain was hypothesized to bind to substrate mRNA molecules and provide the phosphate binding pocket that enhances RNase E activity against monophosphorylated R N A substrates. In P N P a s e the S1 domain was hypothesized to bind to substrate mRNA molecules separately from the active site of the enzyme and therefore impart processivity to its exonuclease activity. In short this work has shown that both of these hypotheses will need revision. A simple scheme for the purification of hexahistidine-tagged S1 domain of RNase E or P N P a s e has been established and has proven to be effective in purifying each protein in sufficient quantities to permit strcutural investingations.  4.1 Function of RNase E S1 domain  4.1.1 R N A binding Although studied in isolation from the intact enzyme, the S1 domain is implicated in playing a role in binding substrate R N A to full length RNase E. The results from the NMR titration of the domain with nucleic acid, although represented graphically in Figure 17, clearly showed that the change in chemical shifts followed a sigmoidal titration 76  curve. This result indicates that this is a genuine binding interaction as it is saturable. In addition, the localization of these altered chemical shifts to one area of the structure confirms that this interaction is specific. Filter binding results also quantify the interaction of the S1 domain of RNase E with S L 9 A R N A molecules (Figures 12 and 13). These experiments estimate a K s of between 1 and 2 u.M. This apparent K is d  d  consistent with those of cold shock domain proteins from the C s p A family. C s p B, C s p C and C s p E have K for binding substrate R N A in the low micromolar range (12.6 u.M, d  3.9 u.M, and 0.9 u,M respectively), consistent for a role in an R N A chaperone function [97, 98]. 4.1.2 Specificity The issue of the specificity of the RNase E S1 domain has been addressed in this work through filter binding assays. The dependance of binding on pH or KCI concentration was analyzed in two filter binding experiments (Figures 9 and 10). These experiments indicate that charge interactions are very important in the interaction of the RNase E S1 domain with substrate R N A molecules. In fact, ionic interactions may dominate over hydrophobic interactions as increasing the ionic strength weakens binding instead of strengthening it. In terms of RNase E's substrate specificity, this may indicate that the S1 domain predominately interacts with the phosphate backbone of the R N A and may not provide a means of R N A substrate recognition, as the negative charge of the phosphate backbone is sequence-independent. Confirmation of the lack of sequence specificity of this domain is shown in Figures 13 and 14. These results show that the wild type RNase E S1 domain does not preferentially bind 5' -end-labelled poly A, poly U or poly C. While this strongly suggests that the domain does not have any broad sequence preference for binding, it does not 77  eliminate the possibility that there exists a specific sequence to which this domain binds with a much lower Kd. It has been shown, for example, through the S E L E X method that Csp B, C s p C and C s p E have lower K s for particular nucleic acid sequences [97]. d  However, sequence-independent binding does not contradict a role for the S1 domain in the function of RNase E as an endonuclease having a general role in m R N A decay and being able to recognize many mRNAs. An alternate site on this enzyme could impart specificity for the AU-rich consensus sequences of targets. Performing a S E L E X experiment using the isolated S1 domain would reveal whether the domain possesses any sequence specificity. This thesis tested a major hypothesis that the RNase E S1 domain exhibits specificity for binding monophosphorylated R N A , thereby imparting RNase E with the observed increased activity on monophosphorylated R N A as compared to triphosphorylated R N A . The results in Figure 12 show that the domain binds to monophosphorylated and triphosphorylated S L 9 A with similar affinities. This suggests that the S1 domain of RNase E is not the putative phosphate binding pocket [99]. Although the stoichiometry of the interaction of the S1 domain with R N A is unknown, its size indicates that multiple domains could bind single-stranded R N A of the length used in the filter binding assays. The domain interacts with only a few nucleotides in determined structures of ob-fold proteins complexed to nucleic acids [12]. Interestingly, the stretch of 30 A residues in S L 9 A does not alter the K of binding d  relative to S L 9 indicating that the S1 domain does not bind preferentially to the 3' -poly A tract. The data in Figure 8 do show that the domain does bind to poly A R N A . Moreover, the domain appears to bind homopolymeric R N A with the same affinity as the S L 9 A and S L 9 RNAs. All the tested substrates, monophosphorylated SL9A,  78  triphosphorylated SL9A, monophosphorylated S L 9 and homopolymeric R N A , share limited common properties. These include a 5' -end and 3' -end as well as some single stranded character at their 5' and 3' ends. It is conceivable, therefore, that this domain imparts the preference of RNase E for substrates with a 5' -single stranded terminus [8] but not its specificity for 5' monophosphorylated R N A (see Figure 31). Thus, this thesis has established that the putative phosphate binding pocket is independent from the S1 domain. It, however, is possible that in the context of a tetramer of RNase E the S1 domain forms a phosphate binding pocket. In Figure 31 the N terminal domains are represented as independent; however, the relative position of these domains in the three dimensional structure of RNase E is completely unknown. For example, the S1 domain may be in close proximity to the active site and therefore impart the enzyme's single -strand -specific activity. And it is also possible that the phosphate binding pocket lies within the catalytic domain of RNase E as suggested previously [2].  4.1.3 Structure and dimerization The S1 domain of RNase E has been confirmed structurally as an S1 domain, a subclass of ob-fold domains. While this has been determined by sequence analysis previously, a high resolution structure is now available for aiding the determination of the function of this domain (Figure 7). The previous model of the S1 domain of RNase E was constructed by threading the RNase E sequence onto the P N P a s e S1 domain structure and has now been confirmed to be a reasonable estimate and a useful structure [61]. The general fold of the S1 domain predicted by Diwa et al. agrees with the NMR -derived and x-ray crystal structure. There are differences, however, the most obvious of which is the lack of secondary structure in the loop connecting (3-sheet 3 and (3-sheet 4 in the modelled domain (L3 in Figure 7). 79  Cterm  o Figure 31: Domains in RNase E. A cartoon schematic of RNase E is shown with the amino terminus on the left and a substrate R N A molecule with the 5' end on the left. The S1 domain studied in this thesis is represented by a barrel binding substrate, the single -strand -specific catalytic domain is represented by the pair of scissors cutting the R N A substrate and the scaffolding region is represented as having binding sites for the components of the degradosome. The domain labelled * is the putative phosphate binding pocket. While this domain is represented at the extreme amino terminus, the location of this putative domain is unknown.  80  Multiple lines of evidence that the isolated S1 domain forms a dimer have been presented in this work. The N M R experimental work, conducted by Dr. Mario Schubert, indicates a concentration -dependent tumbling time that is longer than expected for a dimer. The tumbling time is consistent with an equilibrium between monomer and dimer. The tumbling time increases with increased S1 domain concentration, also consistent with an equilibrium in solution between monomer and dimer [95]. Additional NMR data contains intermolecular N O E s that are located on a putative dimerization surface distinct from the surface implicated in substrate binding. The high resolution x-ray crystal structure of the RNase E S1 domain (Figure 7), also the work of Dr. Mario Schubert, reveals an assymetric unit that contains two monomers. While not conclusive, this is also consistent with an ability of the domain to dimerize. Finally, glutaraldehyde crosslinking experiments show that the RNase E S1 domain can be crosslinked readily to form a larger species, likely a dimer (Figure 8). The finding that the isolated RNase E S1 domain can form a dimer has consequences for the multimeric state of the full length enzyme. This is consistent with previous estimates of the stoichiometry of RNase E in the degradosome, which predict RNase E to exist as at least a dimer. This is also consistent with the quaternary structure of the highly homologous enzyme, RNase G , that suggest this endonuclease exists as a tetramer [100]. The finding of interacting amino acids in this domain that are separate from the R N A binding site suggest that this domain may have two functions within RNase E. The domain would serve to bind substrate and to provide for proteinprotein contacts that stabilize an RNase E multimer. While the S1 domain may provide some contacts involved in stabilizing an RNase E multimer, it is clear that there must be others as the equilibrium in solution does not strongly favour a dimer (eg. Figure 8).  81  A model is presented in Figure 32 which attempts to account for the observed dimerization of the S1 domain of RNase E and recent evidence that RNase E is active as a tetramer [101]. The model predicts that the temperature -sensitive mutation G 6 6 S interrupts interactions in the RNase E tetramer as it does not appear to interrupt R N A binding or S1 domain dimerization. 4.1.4 R N A binding residues The NMR titration data show a localized R N A interaction surface in the domain. This local surface is consistent with previous mutational studies of the RNase E S1 domain [61] and is the same region utilized by most ob-fold proteins to interact with nucleic acids [12]. Filter binding assays showing the importance of charge interactions in the recognition of nucleic acid by this domain were confirmed by the NMR titration experiment. The residues showing chemical shift perturbation included 3 lysines and 2 arginines (see figure 17). A large number of hydrophobic amino acids also showed alterations in chemical shift during the titration. These included a phenylalanine residue (F57) that in the homologous S1 domain of P N P a s e was predicted to bind R N A (see Figure 18) [58]. The three designed mutations in this study, F57A, R64A and R109A all showed defects in autoregulation in vivo in a previous study by Diwa et al. [61]. Diwa et al. went on to show that the RNase E catalytic domain (amino acids 1 - 601) containing F57A exhibited lower cleavage rates on model substrates in vitro. Surprisingly, although these mutants show defects in vivo and in vitro, the corresponding isolated S1 domain shows very little effect of these mutations on its affinity for R N A (Figure 19). It is possible that the R N A binding capacity of the domain is sufficiently different in the  82  Figure 32: A model of RNase E multimerization. An RNase E tetramer is shown with each monomer containing an S1 domain represented by a barrel structure and the remaining carboxy terminus represented by an oval. The interactions uncovered in this work between S1 domains are indicated by solid double headed arrows. The model shows how a dimer of S1 domain may form in the context of an RNase E tetramer. The hypothesized interactions labelled * , indicated by dashed double headed arrows, may be involved in the dimerization of dimers of RNase E and may be disrupted by the me-1 mutation leaving inactive RNase E monomers. The close coordination of S1 domains may also increase the affinity of RNase E for substrate.  83  context of the full length protein that these mutations become much more deleterious to RNA binding. It is also possible that the observed effects on autocontrol and in vitro catalysis of RNase E are not a function of the domain's ability to bind R N A , but reflect some other function of these residues (eg. dimerization).  4.1.5 me-1 mutation The structure of the S1 domain of RNase E as examined by C D revealed that the rne-1 mutation disrupts the structure of the domain, as predicted by Diwa et al. [61]. The extent of this disruption was previously unknown. It is apparent that this mutation results in significant alteration of structure even at low temperatures (Figure 5). The effect of this temperature-sensitive mutation has been studied extensively [2]. Much like the engineered mutations, this mutation shows very little effect on R N A binding capacity as determined by the filter binding assay even at higher temperatures (see Figures 14 and 15). This points to another effect of this mutation in the context of the full length RNase E. It can be hypothesized that this mutation renders the domain unable to dimerize as effectively as the wild type. However, the me-1 mutation in the S1 domain of RNase E does not prevent dimerization in the crosslinking experiment of Figure 8. The reason why this mutation eliminates catalytic activity from RNase E thus remains unknown. Dimerization may be more favorable in the full length protein and this would decrease the K of the domain for R N A and alter other aspects of R N A binding. Thus, d  mutations with no obvious effect in the isolated domain would have an effect in the full length or catalytic domain of RNase E. Alternatively, these mutations may have an effect on the folding or stability of the full length RNase E that are not evident in the isolated domain.  84  Although problematic experimentally, the RNase contamination of the RNase E S1 domain preparations was interesting in itself. P N P a s e , which interacts with the distal C terminal end of full length RNase E is predicted to have different properties than the S1 domain of RNase E, yet persistently copurifies with this domain. Curiously an inverse form of interaction has been found between the S1 domain of P N P a s e the C terminal scaffolding region of RNase E (X. Miao, unpublished work). However, if the S1 domains of RNase E and P N P a s e do interact to promote copurification of P N P a s e and S1 domains, such an interaction can not be dectected by crosslinking (Figure 8 lanes 11 and 12). Many possible avenues of investigation remain to clarify the role of the S1 domain in RNase E. Single alanine mutations in this domain may not exert much effect on its R N A binding properties. Thus, multiple mutations and/or mutations generating charge reversal might be more useful in determining important interacting residues. It is also possible that the engineered mutations made to date are not in residues that are important for R N A binding. Determination of the structure of the rne-1 RNase E S1 domain would be an interesting experiment if it can be performed. This may reveal structural features that are altered by this mutation and whether they lie in or near the R N A binding surface or closer to residues hypothesized to be involved in dimerization. Thus, allowing insight into which function of the domain is responsible for the severe phenotype of this mutation.  85  4.2 Function of PNPase  S1 domain  This thesis has also investigated the properties of the S1 domain of P N P a s e and examined a long standing hypothesis that this domain imparts processive activity to the exonuclease [84, 85]. An important finding of the work examining P N P a s e is the altered activity of histagged P N P a s e as compared to wild type. While this difference was not examined in detail it is apparent that the different product and the lower activity of the N terminally tagged construct (Figure 22) is reason to abandon it as a model for endogenous PNPase. My data show that the elimination of either the S1 domain or the KH domain of P N P a s e does not cause this enzyme to act distributively. It is possible that processivity is conferred by another site in P N P a s e or that the removal of one of the putative R N A binding domains is not sufficient to eliminate the processivity of P N P a s e . RNase P H is highly homologous to P N P a s e , consisting of a hexamer of P H domains arranged around a central pore like P N P a s e [67]. However, RNase P H lacks the 'accessory' R N A binding motifs, the K H and S1 domains found in P N P a s e . However, Isii et al. have shown that RNase P H may be processive [102]. Their results do not prove the processivity of RNase P H with certainty since the assay in Figure 6 of their paper appears complete at the first time point. Very interestingly, removing the S1 domain from P N P a s e blocks the ability of this enzyme to rescue the cold -sensitive phenotype of a P N P a s e null mutant (see Figure 26). This experiment was initially designed for three reasons. First, the P N P a s e S1 domain resembles cold shock domains [58]. Second, the cold-shock phenotype of a quadruple mutant of the C s p A family can be rescued by overexpressing the S1 domain 86  of P N P a s e [86]. Third P N P a s e expression is increased about 5-fold in cold shock and P N P a s e is, therefore, a cold-shock protein [72]. It is reasonable to hypothesize, therefore, that the S1 domain of P N P a s e plays a role during cold shock. The experiment in Figure 26 seems to confirm this possibility. However, the lower specific activity of the P N P a s e AS1 enzyme may account for its inability to rescue the pnp null mutant at low temperature. Expression of the constructs relies on leaky expression from the vector. Moreover, the constructs do not contain the 5' UTR and are not subject to autoregulation. Another interesting avenue would be to test P N P a s e AS1 phosphorolysis at lower temperatures to examine the possiblility that the S1 domain destabilizes secondary structures and enhances P N P a s e activity under these conditions. Finally, the S1 domain of P N P a s e may play a direct role in the autocontrol of the enzyme. The S1 domain may recognize features of the 5' -UTR of the pnp transcript and facilitate its degradation. It is known that the K H and S1 domain are necessary for efficiency of the autoregulatory mechanism of P N P a s e [83]. Previous results from this laboratory indicate that the S1 domain of P N P a s e may be necessary for the interaction of P N P a s e with the degradsome (Miao, X., unpublished). This possibility suggests a dual function for the domain in protein-protein interaction and R N A binding. A s seen with the S1 domain of RNase E, the S1 domain of P N P a s e contains protein interaction and R N A interaction surfaces. The results of this thesis have shed light on the function of S1 domains in E.coli mRNA degradation. It is anticipated that many of the properties discovered in this work are also present in the S1 domains of the endoribonuclease RNase G and the  87  exoribonuclease RNase II. Many new questions about the S1 domain now have a basis from which to start.  88  5 Bibliography 1.  Apirion, D., Degradation of RNA in Escherichia coli. A hypothesis.  Mol Gen  Genet, 1973. 122(4): p. 313-22. 2.  Coburn, G.A. and G.A. 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