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Spo0A-stimulated transcription initiation at the bacillus subtilis spoIIG promother Seredick, Stephen David 2005

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SPOOA-STIMULATED TRANSCRIPTION INITIATION AT THE BACILLUS SUBTILIS spoIIG PROMOTER. by STEPHEN DAVID SEREDICK B.Sc, University of British Columbia, 1996 A THESIS SUBMITTED IN PARTIAL F U L F I L L M E N T OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE F A C U L T Y OF G R A D U A T E STUDIES Microbiology and Immunology THE UNIVERSITY OF BRITISH C O L U M B I A July, 2005 © Stephen David Seredick, 2005 11 A B S T R A C T As a last resort to insurmountable environmental stresses, the common soil bacterium Bacillus subtilis undergoes a complicated morphological transformation to produce a metabolically dormant spore. The central regulator of sporulation is a response regulator transcription factor, SpoOA. When activated by phosphorylation, SpoOA~P directly stimulates or represses transcription at a number of promoters encoding genes involved in the early stages of sporulation. This thesis explores the mechanism by which SpoOA~P activates expression at one of these promoters, the spoIIG promoter. At the spoIIG promoter, SpoOA~P binds to OA boxes and interacts with RNA containing o A to facilitate D N A strand separation. Precisely how Spo0A~P stimulated D N A strand separation was not understood. In this thesis, I show that Spo0A~P acts prior to denaturation of the D N A strands and has no effect on transcription from D N A templates in which the -10 element is artificially denatured to expose the non-template strand. Investigation of the thermal dependence of transcription from a series of artificially denatured promoters suggested that strand separation was the primary thermodynamic barrier to transcription initiation but indicated that SpoOA~P did not reduce this energetic barrier. Kinetic assays revealed that SpoOA~P stimulated both the rate of formation of initiated complexes as well as increasing the number of RN AV-spoIIG complexes capable of initiating transcription. These results implied that SpoOA exerted its effect on RNA polymerase prior to the formation of an open complex. I present evidence for a closed intermediate complex formed by RNA polymerase, Spo0A~P and the spoIIG promoter. To isolate the effect of Spo0A~P on events prior to DNA strand separation at the spoIIG promoter I used D N A fragments that contained only promoter sequences 5' to the -10 element in electrophoretic mobility shift assays. RNA polymerase bound to these fragments readily and the RNA polymerase-spo/JG complexes recruited SpoOA~P. This complex was stabilized by SpoOA~P. In addition I investigated the effect of truncating the upstream portions of the spoIIG promoter which contain a pair of SpoOA binding sites at approximately -90 relative to the transcription start site and found that this D N A inhibited the binding of RNA polymerase. Strikingly, SpoOA~P binding depended on the presence of the D N A upstream of the OA boxes, suggesting a role for the RNA polymerase a subunit carboxy-terminal domains in transcription at this promoter. Finally, SpoOA~P binding to the promoter played two distinct roles. Promoter-distal bound SpoOA~P increased the local concentration of SpoOA~P available to bind to promoter-proximal SpoOA binding sites, while promoter-proximal bound SpoOA~P stabilized and modified pre-formed RNA polymerase-spo/JG complexes. The data is consistent with a model whereby SpoOA~P stabilizes RNA polymerase during the initial stages of open complex formation at the spoIIG promoter, but only until the non-template strand of the -10 element is fully single-stranded. These effects account for transcription activation of the spoIIG promoter by SpoOA~P. IV T A B L E OF CONTENTS ABSTRACT. ii T A B L E OF CONTENTS. iv LIST OF T A B L E S . viii LIST OF FIGURES. ix LIST OF ABBREVIATIONS A N D SYMBOLS. xi A C K N O W L E D G E M E N T S . xii INTRODUCTION. 1 1 Sporulation in Bacillus subtilis. 1 1.1 Sporulation Morphology. 2 1.2 The Sporulation Transcriptional Regulatory Network. 2 2 Regulating the Master Regulator. 4 2.1 Two-Component Signal Transduction Systems. 6 2.2 Activation of SpoOA. 6 2.2.1 The Sporulation Phosphorelay System. 6 2.2.2 Effect of Phosphorylation. 7 3 Bacterial Gene Regulation. 9 3.1 Promoter Elements. 9 3.2 RNA Polymerase. 11 3.2.1 p and (3' Subunits. 11 3.2.2 a Subunits. 12 3.2.3 aSubunit. 13 3.3 Transcription Initiation. 15 3.4 Repression. 17 3.5 Activation. 19 4 Transcription Initiation at the spoIIG Promoter. 21 4.1 Characteristics of the spoIIG Promoter. 21 4.2 Regulatory Inputs at the spoIIG Promoter. 23 4.2.1 SinR. 23 4.2.2 Soj. 23 4.2.3 iNTPs. 24 4.2.4 SpoOA. 25 4.3 A Working Model for Activation of the spoIIG Promoter by SpoOA~P. 30 E X P E R I M E N T A L PROCEDURES 32 1 Manipulation of D N A and Oligonucleotides. 32 1.1 Preparation of spoIIG and IIG17 Promoter Fragments. 32 1.2 Heteroduplex Templates. 32 1.2.1 Construction of pSK3/ /G + and pSK3//G\ 34 1.2.2 Construction of pSK6/ /G + and pSK7//G + . 34 1.2.3 Purification Single-stranded Plasmid DNA. 37 1.2.4 Production of Heteroduplex Templates. 37 1.3 End-labeling of Restriction Fragments. 38 1.4 Oligonucleotide Labeling. 39 1.5 Preparation of Labeled D N A Fragments. 39 2 Protein Purification. 41 2.1 SpoOA. 41 2.2 SpoOA c. 42 2.3 SpoOF. 43 2.4 RNA Polymerase. 43 3 Activation of SpoOA. 45 4 In vitro Transcription. 45 5 Potassium Permanganate Footprinting. 46 6 DNase I Footprinting. 47 7 Electrophoretic Mobility Shift. 47 vi RESULTS. 49 1 RNA Polymerase Purification. 49 2 Characterization of Late Events in Transcription Initiation at the spoIIG Promoter. 49 2.1 Levels of in vitro Transcription from Heteroduplex Templates. 53 2.2 Effects of SpoOA~P on in vitro Transcription from Heteroduplex 53 Templates. 2.3 Structural Probing of Open Complexes with Potassium Permanganate. 55 2.4 Temperature Dependence of Transcription. 58 2.5 Temperature Dependence of Open Complex Formation. 62 2.6 Relative Rates of Acquisition of Heparin-Resistance. 64 3 Characterization of Early Events in Transcription Initiation at the spoIIG Promoter. 66 3.1 Evidence for Low Temperature Intermediates at the spoIIG Promoter. 66 3.2 Evidence for a Closed RNAP-s/w/JG-SpoOA-P Intermediate. 68 3.3 Complex Formation on a Truncated Derivative of the spoIIG Promoter. 70 3.4 DNase I Footprinting of the spoIIG Promoter and the TIIG Promoter Fragment. 72 3.5 Characterization of Complex Assembly on the TIIG Promoter Fragment. 75 3.5.1 RNAP Titration. 75 3.5.2 Spo0A~P Titration. 78 3.6 Definition of the Minimal Promoter Fragment Necessary for Complex Formation. 78 3.6.1 Downstream Truncations. 78 3.6.2 Upstream Truncations. 81 3.7 Characterization of Complex Assembly on the (-80)77/G Promoter Fragment. 81 3.7.1 RNAP Titration. 81 3.7.2 Spo0A~P Titration. 83 3.8 Complex Formation on Variants of the TIIG Promoter Fragment. 85 3.8.1 Complex Formation on TIIGJDown. 85 3.8.2 Complex Formation on TIIG2.1 Up and TIIG2.1 Down. 87 V l l 3.9 Relative Stabilities of Complexes Formed on TUG, TIIG2.1Up and TIIG2.1Down. 90 DISCUSSION. 93 1 Technical Advances. 93 1.1 R N A Polymerase Purification. 93 1.2 Radiolabeled Template Synthesis. 94 1.3 Alterations in E M S A Technique. 94 2 A Model for Transcription Initiation at the spoIIG Promoter. 95 3 Caveats of the Proposed Model. 99 3.1 The Role of a-UP Element Interactions in B. subtilis. 99 3.2 The Role of aCTDs in Transcription Initiation at the spoIIG Promoter. 100 3.3 In vivo Concentration of RNAP and the Recruitment of SpoOA. 103 3.4 Downstream Re-positioning of RNAP. 104 3.5 SpoOA Recruitment and aA-Dependent Mutants in SpoOA. 109 3.6 SpoOA Stimulation and D N A Topology. 110 3.7 SinRandSoj. 112 4 Transcription Initiation. 113 4.1 Closed Complex (RPC) Formation. 114 4.2 Open Complex (RPo) Formation. 116 5 Transcription Activation. 118 5.1 SpoOA. 118 5.2 Other Response Regulators. 120 5.2.1 The NtrC Subfamily. 121 5.2.2 The NarL Subfamily. 121 5.2.3 The OmpR Subfamily. 122 5.3 CAP. 123 5.4 Xcl. 126 6 Transcriptional Regulation: General Mechanisms. 127 REFERENCES. 129 V l l l LIST OF T A B L E S Table 1. Phenotypes Associated with spoOA Mutations. 26 Table 2. Plasmids Used in this Thesis. 33 Table 3. Primers Used in this Thesis. 35 Table 4. Strains Used in this Thesis. 36 Table 5. Primers Used to Prepare Radiolabeled Fragments Used in this Thesis. 40 Table 6. In vitro Transcription: Template Utilization. 54 Table 7. In vitro Transcription: Temperature-Dependence and 60 Acquisition of Heparin-Resistance. ix LIST OF FIGURES Fig. 1. Sporulation morphology. 3 Fig. 2. Regulatory gene network for sporulation. 5 Fig. 3. Activation of SpoOA. 8 Fig. 4. Basic features of the promoter and RNAP-promoter interactions. 10 Fig. 5. Functional map of a. 14 Fig. 6. Minimal model for transcription initiation. 16 Fig. 7. Simple repression mechanisms. 18 Fig. 8. Simple activation mechanisms. 20 Fig. 9. The spoIIG promoter. 22 Fig. 10. Structure of the Spo0AN receiver domain. 27 Fig. 11. Structure of the SpoOAc activation domain. 28 Fig. 12. RNAP purification. 50 Fig. 13. In vitro transcription assay templates. 52 Fig. 14. D N A strand separation at representative promoters. 57 Fig. 15. Effect of SpoOA~P on the temperature dependence of transcription. 61 Fig. 16. Effect of temperature on open complex formation. 63 Fig. 17. Time course of initiated complex formation. 65 Fig. 18. SpoOA~P and RNAP form a low temperature intermediate complex at the spoIIG promoter. 67 Fig. 19. RPOA complexes formed at low temperatures can initiate transcription i f brought to 37 °C. 69 Fig. 20. RNAP and SpoOA~P form a complex on a truncated promoter derivative, 777G. 71 Fig. 21. SpoOA~P re-models RNAP protection of the spoIIG promoter. 73 Fig. 22. SpoOA~P and RNAP form a specific complex on a TUG promoter fragment. 74 Fig. 23. Complex assembly on TUG - RNAP titration. 76 Fig. 24. Complex assembly on TUG - SpoOA~P titration. 77 Fig. 25. Complex formation on 3' nested spoIIG promoter fragments. 79 Fig. 26. Complex formation on 5' nested spoIIG promoter fragments. 80 X Fig 27. Complex assembly on (SO)TIIG - RNAP titration. 82 Fig. 28. Complex assembly on (S0)TIIG - Spo0A~P titration. 84 Fig. 29. Complex assembly on TIIG 1 Down. 86 Fig. 30. Complex assembly on TIIG2.1Up and TIIG2.1Down. 88 Fig. 31. Complex stability on TIIG, TIIG2.1Up and TIIG2.1Down. 91 Fig. 32. A model for transcription initiation at the spoIIG promoter. 97 Fig. 33. Overlapping a A and Spo0A~P contacts at the spoIIG -35 element. 106 Fig. 34. Thermodynamically separable steps in D N A strand separation at the spoIIG promoter. 117 Fig. 35. Response regulator-DNA interactions and RNAP contact surfaces. 124 LIST OF ABBREVIATIONS A N D S Y M B O L S aCTD(s) carboxy-terminal domain of the RNA polymerase a subunit(s) aNTD(s) amino-terminal domain of the RNA polymerase a subunit(s) E M S A electrophoretic mobility shift assay INTP(s) initiating nucleotides RNAP RNA polymerase R E elongating R N A polymerase RP RNA polymerase-TIIG complex RPc closed RNA polymerase-promoter complex RPo open RNA polymerase-promoter complex RPOA RNAP-777G-SpoOA~P complex RPOAc closed RNA polymerase-s/?o/JG-SpoOA~P complex RPOAo open RNA polymerase-spo//G-SpoOA~P complex SpoOAc carboxy-terminal domain of SpoOA SpoOAN amino-terminal domain of SpoOA SpoOA~P activated, phosphorylated form of SpoOA UP element upstream promoter element A C K N O W L E D G E M E N T S I have heard it said that in writing a book one gets farther and farther from the beginning but without getting closer and closer to the end. A thesis is a book (or sorts) and my experience certainly bears the statement out. Having gotten that off my chest, I 'll try to resist any further literary urges. I thank my committee members for their guidance; bacterial gene regulation is a little removed from your research interests but I think we set a departmental record for intervals between meetings. Hopefully, my thesis hasn't suffered too much in consequence. I thank my cohorts in the lab, especially Brett and Barb, for their insight and help. I've enjoyed the time spent in your company. I'm grateful for the friendship of Shelley Small, A l i Tehrani and Dave Oliver. I've profited greatly from our conversations (some of which were even related to research). Most of all I owe Barb, my wife and SpoOA co-conspirator, a debt of gratitude. How she tolerates a husband who memorizes details from 20 year-old articles but remembers her birthday by process of elimination is remarkable. Most students dedicate their theses to their family. I realize the gesture is symbolic, but I've always felt this practice odd since a thesis has limited value to those to whom it is dedicated. In fact, I'm fairly sure most spouses would have preferred a nice dinner now and then, and a toddler a trip to the park to feed the ducks. So my thesis isn't so much dedicated to my wife, parents and family as a reflection of their support and their encouragement of my intellectual self-indulgence. A l l students finish their graduate degrees less ignorant than they began, but I'm fortunate that '11 also be leaving a better person. Whether this is because of my mentor or simply happened under his watch, I'm still profoundly grateful. Thanks, George. 1 INTRODUCTION The ability to respond to the environment is a definitive feature of life, and identifying the mechanisms by which cells or organisms modify their metabolic activity or behavior has been one of the central problems confronting biologists. Insight into the molecular nature of cellular responses grew out of Jacques Monod's work on preferential sugar use in Escherichia coli and, though not widely appreciated, Bacillus subtilis (Judson, 1996). Eventually, Monod's group would concentrate on the regulation of diauxic growth in E. coli. Understanding this deceptively simple cellular response in a common intestinal bacterium had profound consequences, leading to the creation of the field of molecular biology (Echols and Gross, 2001), a Nobel prize for Monod, Jacob and Lwoff for their discoveries concerning genetic control of enzyme and viral synthesis, and the adoption of E. coli as the model "model organism". Though relegated to runner-up status in prokaryotic biology, in part to Monod's choice and subsequent successes, B. subtilis has been intensively studied due to the ease with which it can be genetically manipulated and because of a compelling morphological transformation, sporulation, in response to deteriorating conditions. 1 Sporulation in Bacillus subtilis. Bacillus subtilis is a useful model system for molecular biology for two reasons. Firstly, in its relative simplicity B. subtilis provides an excellent opportunity for understanding how cellular networks make decisions. Specifically, B. subtilis cells embark on one of several alternative stationary phase survival strategies following exponential growth - the production of flagella to enhance motility, the development of competence, the secretion of degradative enzymes, the secretion of secondary metabolites and sporulation. The isolation of mutants affecting each of these processes identified the genetic elements for control and therefore permitted molecular studies. Secondly, since vegetatively growing cells and spores are physiologically and morphologically distinct, the transition between the two cell types represents one of the simplest known developmental processes (along with the stalked-swarmer cell transition in Caulobacter crescentus). Before examining the molecular control systems guiding sporulation, we should first consider the morphological transformation. 2 1.1 Sporulation Morphology. The successive morphological stages in sporulation are shown in Fig. 1 (Errington, 1993; Errington, 2003; Stragier and Losick, 1996). Sporulation is usually induced by starvation and high population density. Cells that have ceased to grow and have passed through the stage of competence but show no other morphological features are said to be in stage zero of sporulation. The first dramatic event in sporulation is the formation of a medial or axial filament of D N A containing a pair of complete chromosomes. As no mutants have been found that block sporulation at this stage, it is not assigned a specific name (though when the stages of sporulation were morphologically rather than genetically defined, cells in such a condition where termed stage I). The second stage of sporulation begins with an asymmetric division of the cytoplasm creating cells of unequal size. In the third stage, the larger or mother cell engulfs the smaller cell and membrane fusion around it creates the forespore protoplast, such that the forespore exists as a cell-within-a-cell. In subsequent stages the cortex, a modified form of cell wall, is synthesized between the forespore and mother cell membranes, while the mother cell assembles a proteinaceous coat on the outside surface of the spore. During the deposition of these layers, small acid soluble proteins are synthesized in large amounts to coat the forespore chromosome conferring resistance to U V radiation, while dipicolinic acid and C a 2 + are absorbed resulting in dehydration and mineralization of the forespore. After 6 to 8 hours of development when maturation is complete, the ripened spore is liberated by mother cell lysis. 1.2 The Sporulation Transcriptional Regulatory Network. Though the morphological stages of sporulation appear simple relative to other developmental processes such as craniofacial patterning and limb development in vertebrates, sporulation still requires the coordination of hundreds of genes (Piggot and Losick, 2002). Since it is difficult i f not impossible to simultaneously consider the relationships between this many genes insightfully, one must attempt to impose manageable complexity upon confusion. As this thesis is concerned with transcriptional regulation, one profitable way to consider sporulation is as a network of transcription factors controlling this developmental process. Vegetative _ ,__ Cortex ° ^ Engulfment Growth Formation Asymmetric Forespore Coat Septation Protoplast Formation Maturation Release o o o iii H2 111 rv v v i v u spoO spoil spoil spoIII spoIV spoV spoVI Figure 1. Sporulation morphology. The main sequence of morphological events begins with the vegetative cell and ends with the release of a mature spore. Roman numerals designate the stages of sporulation characterized by electron microscopy. The italicized spo designation identifies genes whose mutation stalls sporulation at the stage immediately preceding it. For example, mutation of spoOA prevents asymmetric septum formation, while mutation of spoIIG prevents engulfment. Stage I represents cells that have replicated their genome and formed axial DNA filaments, but since this is also observed in vegetative cells and no sporulation-specific mutants have been characterized the designation has been dropped. See text for details. Adapted from Errington, 1993. 4 A schematic of the sporulation transcriptional regulatory network is presented in Fig. 2. It includes the core transcription factors directly controlling the expression of genes that drive spore formation, and the known relationships between them. The figure legend explains the use of symbols. There are several salient features. First, cross-repressive interactions between abrB, spoOA and spoOH restrain the cell from sporulating. AbrB keeps SpoOA levels low by repressing spoOH. When activated (see Introduction, section 2.2A), the initial low levels of SpoOA are sufficient to repress abrB, indirectly lifting repression of spoOH. As a H levels rise (spoOH), so too does the transcription of spoOA. Second, activated SpoOA, SpoOA~P, initiates two programs of gene expression, one specific to the forespore ending with the expression of genes such as those encoding small acid-soluble proteins (SASPs), and one specific to the mother cell ending with the expression of genes such as those for spore coat proteins. Third, sporulation is driven by a series of linked feed-forward regulatory loops. For example, SpoOA enhances its own synthesis via a positive feedback loop, and activates spoIIG. o , (spoIIG) is required for the transcription of sigK (aK) and spoIIID that drive their synthesis by a positive feedback loop as well, and activate gerE expression, the final mother cell transcription factor. SpoOA also activates spoIIA, whose product a F is required for the transcription of spoIIIG encoding a . In turn, a positive feedback loop drives up the synthesis of a , required for the expression of the gene for the final forespore-specific transcription factor, spoVT. These regulatory loops install serial programs of gene expression that correlate with some (but not all) of the morphologically defined stages. Similar controls ensure that genes involved in the other stationary phase survival strategies are expressed to the exclusion of genes implicated in alternative strategies. In fact, the regulatory network is considerably more complicated in that several of the genes implicated in the early phases of sporulation (spoOA, abrB, spoOH, codY, scoC and sinR) control genes involved in the other stationary phase survival strategies (Smith, 1993; Strauch and Hoch, 1993). Given its central role, SpoOA is the master regulator of sporulation. 2 Regulating the Master Regulator. SpoOA is a member of an enormous family of proteins called response regulators. Response regulators are usually paired with histidine kinases that collectively form a two-component sigA spoOJA spoIIG sin abrB spoOH n T IT S/?0&4_ SpoOAy 1 Coat Proteins scoC spoIIA 1 r — spoIIIG spoVT '1 SASPs Figure 2. Regulatory gene network for sporulation. Short horizontal lines from which bent arrows extend represent c/s-regulatory elements responsible for the expression of the genes named beneath the line. Genes are arranged in a temporal hierarchy; those expressed during vegetative growth are at the top, those expressed during later stages of sporulation are located progressively lower. spoOA is expressed from a vegetative promoter, spoOAv, and a stationary phase promoter, spoOAs. Regulatory inputs are represented by the coloured lines; each gene is represented by a distinct colour. The mother cell-specific gene expression program is green and ends with the production of coat proteins while the forespore-specific gene expression program is purple and ends with the production of SASPs. Regulatory inputs ending with an arrowhead act positively; regulatory inputs ending with a short horizontal line act negatively. Data taken from Piggot and Losick, 2002 and Eichenberger et al, 2004. 6 signal transduction system. As has been so presciently observed, microbes use such systems to "see their world" (Fabret et al, 1999). 2.1 Two-Component Signal Transduction Systems. The two-component system is the most common form of signal transduction by which bacteria adapt their behavior in response to environmental cues (Hoch and Silhavy, 1995; Inouye and Dutta, 2003). Typically, such systems consist of a ligand-responsive histidine kinase and a response regulator, most often a transcription factor. Binding of a specific ligand alters the enzymatic activity of the kinase potentially stimulating either autophosphorylation, phosphorylation of a conserved aspartate in the receiver domain of its cognate response regulator, or dephosphorylation of the same target (Stock et al, 2000). Phosphorylation of the response regulator by the histidine kinase controls its activity, frequently by modulating its affinity for target promoters and thereby modifying gene expression. Two-component systems are also found in some archaea, yeast, and plants; and at least in bacteria, there seems to be a correlation between genome complexity, adaptability and the number of two-component systems a particular species possesses. For example, the intracellular parasite Mycoplasma genitalium possesses no two-component signal transduction systems, while Haemophilus influenza, Thermotoga maritime, E. coli and B. subtilis possess 9, 19, 30 and 70 two-component systems respectively (Stock et al, 2000). 2.2 Activation of SpoOA. 2.2.1 The Sporulation Phosphorelay System. In addition to the classical single step phosphotransfer system, some three step phosphotransfer systems, termed phosphorelays, exist. In some systems, like the BvgA/BvgS system controlling expression of virulence genes in Bordetella pertussis(Cotter and Jones, 2003), three phosphotransfer domains are present in a single hybrid kinase. In the sporulation phosphorelay system, all phosphotransfer domains are on discrete proteins. 7 Briefly, environmental signals trigger the autophosphorylation of one of several kinases, the primary kinase being KinA. A phosphoryl group is transferred from KinA, to the isolated receiver domain SpoOF, to the histidine phosphotransferase SpoOB and ultimately to SpoOA to activate the master regulator of sporulation (Fig. 3) (Burbulys et al, 1991). The additional phosphotransfer reactions in a phosphorelay provide increased opportunities for regulating phosphate flow. In the sporulation phosphorelay, the aspartate phosphatase SpoOE family of proteins (Perego and Hoch, 1991) inactivates Spo0A~P, while RapA dephosphorylates SpoOF (Perego et al, 1996). A sophisticated peptide processing/export/import circuit controls the activity of RapA (Perego, 1997). It has been suggested that this circuit determines how long RapA can delay SpoOA activation while allowing alternative physiological events to occur (Perego, 1998). In addition the anti-kinases, Kip l (Wang et al, 1997) and Sda (Burkholder et al, 2001), inhibit the autophosphorylation of KinA. Response regulator activity can also be controlled by preventing dephosphorylation (Kato and Groisman, 2004), or by interfering with the recruitment of RNAP to the promoter (Ansaldi et al, 2004), but analogous proteins controlling Spo0A~P activity have not yet been identified. 2.2.2 Effect of Phosphorylation. Response regulators are defined by their possession of a conserved N-terminal regulatory domain, the receiver module. The receiver consists of a doubly wound a/p protein, with a central 5-stranded parallel P-sheet surrounded by 5 a-helices. In SpoOA, Asp56, located in the solvent-exposed loop between P3 and a3, is phosphorylated (Lewis et al, 1999). Highly conserved aspartates (AsplO and Aspl 1 in SpoOA) form an acidic cluster with the phosphorylated aspartate that coordinate a M g 2 + ion required for phosphotransfer. Two other residues, a threonine and an aromatic residue (Thr84 and Phel05 in SpoOA) are implicated in the phosphorylation-induced conformational change (Lewis et al, 1999; Muchova et al, 2004). Phosphorylation appears to shift the equilibrium between inactive and active conformational states of the receiver (Kern et al, 1999; Nohaile et al, 1997; Volkman et al, 2001). Structural elements are modestly re-positioned, but these changes dramatically affect topological and electrostatic features of the receiver domain. By controlling the presentation of alternate 8 rapA KinA~P KinA kinA LZ RapA SpoOF SpoOF~P SpoOF spoOF SpoOB~P SpoOB spoOB SpoOA SpoOA~P spoOA SpoOE spoOE Sporulation Delay Sporulation Killing Factors Cell Wall Biogenesis Cell Division Biofilm Formation Spore Formation Transition State Repression DNA Replication Flagellar Biosynthesis Chemotaxis Cell Metabolism Figure 3. Activation of SpoOA. SpoOA is activated by serial phosphotransfer (blue) from KinA to SpoOF to SpoOB and ultimately to SpoOA. Two phosphatases reduce phosphate flow; RapA dephosphorylates SpoOF~P while SpoOE dephosphorylates SpoOA~P. SpoOA~P has pleiotropic effects (orange). Once activated, SpoOA~P represses expression of rap A, while stimulating expression of kinA, spoOF and spoOA. At elevated concentrations, SpoOA~P also represses transcription from the sporulation-specific promoters of the spoOF and spoOA genes. Besides regulating genes involved in the phosphorelay, SpoOA~P also positively regulates genes involved in sporulation decay, sporulation killing, cell wall biogenesis, cell division, biofilm formation and other genes driving sporulation. SpoOA~P suppresses the expression of genes repressing alternate stationary phase phenomena, DNA replication, flagellar biosynthesis, chemotaxis and some aspects of cellular metabolism. Based on data from Molle et al. (2003). 9 molecular surfaces, phosphorylation controls the interaction of the receiver with alternate binding partners. This structural rearrangement switch has been exploited by bacteria to relieve inhibition of the effector domain (Grimsley et al, 1994; Huala et al, 1992; Simms et al, 1985), to stimulate dimerization (Da Re et al, 1999; Fiedler and Weiss, 1995; Lewis et al, 2002; McCleary, 1996) or oligomerization (Weiss et al, 1992; Wyman et al, 1997), or to stimulate the interaction with other proteins (Blat and Eisenbach, 1994; Welch et al, 1993) or D N A (Aiba etal, 1989; Makino et al, 1989). 3 Bacterial Gene Regulation. 3.1 Promoter Elements. Different D N A sequences within a promoter contribute to recognition by RNAP (see below). Four sequence elements may play a role in promoters recognized by B. subtilis R N A P a A or its homolog in E. coli, R N A P a 7 0 (Fig. 4). The two principal elements are the -10 and -35 hexamers, so called because they are centered about 10 and 35 base pairs upstream from the transcription start site. The -35 element is recognized by a domain of the RNAP a subunit, 04.2, and the -10 element is recognized by another domain, G2A; nucleation of D N A strand separation occurs in the -10 element. Consensus sequences have been identified and crystallographic structures of a-promoter and RNAP-promoter complexes (Campbell et al, 2002; Murakami et al, 2002a) have validated models of promoter recognition based on extensive genetic and biochemical studies. The length of the D N A separating the -10 and -35 elements is critical as well. Two other accessory sequence elements, extended -10 motifs and upstream promoter or UP elements, also influence transcription initiation. Extended -10 motifs are recognized by 03, which may compensate for absent -35 elements and permit open complex formation at relatively low temperatures (Bown et al, 1999; Burns et al, 1996; Keilty and Rosenberg, 1987; Ponnambalam et al, 1986). UP elements are AT-rich sequences upstream of the -35 element to which the carboxy-tenninal domains of the a subunits of RNAP, aCTDs, may bind (Ross et al, 1993), and consensus sequences for each UP element sub-site have been defined (Estrem et al, 1998; Estrem et al, 1999). aCTD-DNA interactions enhance both promoter recognition and subsequent steps during transcription initiation (Ross and Gourse, 2005). The sequence and 10 aNTD aCTD UP element AT-rich -35 element -10 element TTGACA TGnTATAAT Figure 4. Basic features of the promoter and RNAP-promoter interactions. Several general features of a promoter may play a role in directing transcription initiation by RNAP in the absence of accessory factors. The primary features are the transcription start site, +1 (bent arrow) and the -35 and -10 elements (white boxes); the nucleation of DNA strand separation occurs at the upstream edge of the -10 element. The consensus sequences for B. subtilis RNAPrj A are shown below the appropriate box. At some promoters a TG dinucleotide one base pair upstream of the -10 element permits DNA strand separation at low temperatures. AT-rich upstream promoter or UP elements are recognized by aCTDs and stimulate RNAP binding and the rate of conformational changes in RNAP during initiation. UP elements can be divided into proximal and distal sub-sites recognized by individual aCTDs. Non-specific aCTD-DNA interactions contribute to the same processes (Ross and Gourse, 2005). The aCTDs are connected to the aNTDs and the rest of RNAP by a long, flexible linker. RNAP core enzyme (light blue) is composed of o_PP'. The a subunit (dark blue) recognizes the -35, -10 and 11 spacing of these elements dictates the relative affinity of RNAP for a particular promoter and once bound, how rapidly RNAP can initiate transcription. 3.2 RNA Polymerase Transcription is performed by DNA-dependent R N A polymerase (RNAP) and extensive sequence homology exists between bacterial and eukaryotic RNAP subunits (Allison et al, 1985; Sweetser et al, 1987). Bacterial RNAP is an approximately 400 kDa multi-subunit, allosteric information processor that uses the free energy liberated by nucleotide polymerization to drive its forward translocation (Gelles and Landick, 1998; Mooney et al, 1998). Bacterial RNAPs are highly conserved, and high-resolution structures of bacterial RNAPs are available (Murakami et al, 2002a; Murakami et al, 2002b; Vassylyev et al, 2002; Zhang et al, 1999). The core enzyme that is capable of transcript elongation and non-specific D N A binding has a subunit composition of a 2pp'. Promoter-specific transcription initiation requires the association of a a subunit with the core enzyme to form a holoenzyme, a2pP'o\ Several dissociable subunits are sometimes isolated with RNAP holoenzyme. The co subunit apparently serves as a chaperone to assist the folding of the P' subunit (Minakhin et al, 2001). The 5 subunit unique to B. subtilis (absent in E. coli) enhances the selectivity of transcription initiation (Achberger and Whiteley, 1981; Dobinson and Spiegelman, 1987; Hyde et al., 1986; Juang and Hermann, 1994a; Juang and Helmann, 1995). The roles of the holoenzyme subunits are described below. 3.2.1 p and P' Subunits. RNAP has a crab claw-like structure in which the two largest subunits, P and P', form the "pincers". Many of the major functional features of RNAP are located within this primary channel or cleft at the base of the "pincers" (Landick, 2001; Zhang et al, 1999). The active site forms the floor of a chamber that can accommodate an 8-9 base pair R N A : D N A hybrid. At the upstream edge of this chamber a wall of protein, the P' rudder, disrupts base-pairing of the R N A : D N A hybrid, and deflects the nascent transcript into an exit channel and the D N A template through a different opening where it can re-anneal with the non-template strand (Kuznedelov et al, 2002a). At the downstream edge of this chamber, the channel continues and contains determinants responsible for binding 15 to 20 base pairs of downstream DNA. A 12 secondary channel roughly perpendicular to the primary channel allows NTPs and transcription factors access to the active site from the base of the chamber (Artsimovitch et al, 2004; Opalka et al, 2003; Perederina et al, 2004), and a groove in P can accept the non-template strand during transcription elongation. Closure of the pincers around the D N A ensures that RNAP cannot dissociate from its template during transcription. The P subunit binds the initiating nucleotides (iNTPs) and is involved in the initial polymerization steps (Jin and Turnbough, 1994; Kashlev et al, 1990; Mustaev et al, 1991), and both the P and P' subunits make extensive contacts with promoter D N A (Brodolin et al, 2000; Murakami et al, 2002a; Naryshkin et al, 2000; Severinov and Darst, 1997; Studitsky et al, 2001). A coiled-coil in the P' subunit makes extensive contacts with a (Arthur et al, 2000; Gruber et al, 2001; Owens et al, 1998; Sharp et al, 1999; Young et al, 2001), while a flap in P interacts with a to position the two domains that recognize the -35 and -10 elements (Kuznedelov et al, 2002b). 3.2.2 a Subunits. The a subunits of RNAP each consist of two independent domains connected by a flexible linker. The amino-terminal domains, aNTDs, dimerize and provide the scaffold upon which the P and p' subunits are incorporated to form the core enzyme. The carboxy-terminal domains, aCTDs, bind AT-rich UP element D N A and provide a surface accessible to DNA-bound transcription factors; both interactions strengthen RNAP-promoter interactions and may stimulate transcription. Transcription factors that bind immediately upstream from RNAP may interact with the aNTD as well. Sequence-independent interactions of aCTD with upstream promoter D N A also play a major role in promoter recognition and during the later stages of transcription initiation that occur at sites far removed from the a binding sites (Davis et al, 2005; Rao et al, 1994; Ross and Gourse, 2005). That contacts to the a subunits might affect 70 post-binding steps was foreshadowed by the discovery that an aCTD may interact with a 4, thereby bridging the upstream promoter to the core promoter elements (Chen et al, 2003; Ross etal, 2003). 13 3.2.3 o Subunit. The a subunit of RNAP has four primary functions: recognition of the core promoter elements; positioning of the enzyme's active site over the transcription start site; driving D N A strand separation; and providing an interaction surface for transcription factors. Most bacteria contain several alternative a factors controlling specialized regulons activated during specific stress conditions, growth transitions or morphological changes. Generally, organisms that must respond to a greater diversity of stresses or that can adopt alternate developmental forms contain more o factors; for example, E. coli contains 7 o factors, B. subtilis 19 a factors and Streptomyces coelicolor contains 63 a factors (Gruber and Gross, 2003; Helmann and Moran, 2002). The majority of a factors (including B. subtilis aA) are homologous to E. coli a 7 0 but an alternative group of o factors (homologous to E. coli a 5 4) lack sequence homology to a 7 0 and initiate transcription by an entirely different mechanism. Which of the alternative a factors are associated with the RNAP core enzyme at any one time is a function of their relative affinities for RNAP and the amount of active a available. A variety of mechanisms control the levels of active a but are beyond the scope of this thesis (Gruber and Gross, 2003; Ishihama, 2000). 70 Members of the a family contain up to four modules that can be further divided into subregions to which specific activities can be ascribed (Fig. 5) (Gross et al, 1998). o u is extraordinarily acidic and masks D N A binding determinants in free a (Camarero et al., 2002; Dombroski et al., 1993; Dombroski et al., 1992). It is displaced from the active site of RNAP during later stages of transcription initiation (Mekler et al, 2002) and can affect the kinetics of D N A strand separation at some promoters (Vuthoori et al, 2001; Wilson and Dombroski, 1997). A large non-conserved region separating rji.i and G1.2 in E. coli a 7 0 is not present in B. subtilis a A . Mutations in G1.2 prevent promoter melting (Wilson and Dombroski, 1997), even though oi 2 does not crosslink to promoter D N A (Naryshkin et al, 2000). Mutations in 02.1, 02.2 and 0*2.4 reduce the binding of a to RNAP core enzyme (Gruber et al, 2001; Sharp et al, 1999). Mutation analysis has implicated G2.4 in the recognition of nucleotides in the -10 element sequence, probably in a double-stranded form (Daniels et al, 1990; Kenney et al, 1989; Marr and Roberts, 1997; Siegele et al, 1989; Waldburger et al, 1990; Young et al, 14 non-conserved region 1.1 inhibition free a-DNA interaction RNAP binding A 2.1 | 2.2 I 2.3 I 2.4 nt strand binding DNA melting extended -10 -35 element •10 element abortive initiation Figure 5. Functional map of a. a structural domains and evolutionarily conserved regions are shown schematically. The thinner black bar represents the primary sequence of rj while the domain architecture is indicated by the thick black bars. Conserved regions are distinguished by colour and represented by numbered boxes. The functions of the various sub-regions are indicated by arrows. Note that B. subtilis o A lacks the non-conserved region between 1.2 and 2.1. See text for details. Figure adapted from Borukhov and Nudler (2003); Murakami and Darst (2003). 15 2004; Zuber et al, 1989). G2.3 binds to the non-template strand in a sequence-specific manner (Huang et al, 1997; Marr and Roberts, 1997) and mutations in a limited number of conserved aromatic residues in 02.3 impair D N A melting (Huang et al., 1997; Juang and Helmann, 1994b; Rong and Helmann, 1994). Basic residues in the same region have been proposed to position the promoter D N A in the proper orientation for promoter opening (Tomsic et al, 2001). A pair of aromatic residues in C2.3 are particularly important in nucleating D N A strand separation, for the recognition of the boundary between double- and single-stranded D N A , and for isomerization of RNAP between stages in the initiation process (Fenton et al., 2000; Tsujikawa et al., 2002). Amino acids in a 3 0 recognize the extended -10 element to stabilize the open complex (Barne et al, 1997; Bown et al, 1999; Voskuil and Chambliss, 2002). 03.2, an extended loop, partially blocks extension of small transcripts and appears to play a role in abortive initiation (Murakami et al, 2002b). The role of a3.i is unknown. Mutations in some residues of 04.1 impair the association of a and core enzyme (Sharp et al, 1999), and a conserved helix-turn-helix motif in 04.2 interacts directly with the -35 element (Gardella et al, 1989; Siegele et al, 1989). 0-4.2 also provides a surface accessible to transcription factors (Dove et al, 2003) and aCTDs (Ross et al, 2003) bound immediately upstream from the -35 element. 3.3 Transcription Initiation. Transcription initiation is the major regulatory step during gene expression in bacteria and minimally involves several discrete intermediates (Fig. 6). Much of what is known about 70 transcription initiation is based on studies performed with E. coli RNAPc (Browning and Busby, 2004; Murakami and Darst, 2003; Record et al, 1996), but it has been assumed that a common mechanism underlies initiation with other c factors (with the notable exception of E. coli RNAPa 5 4 ) and other bacterial RNAPs. During the first step, RNAP recognizes and binds to promoter elements to form a closed complex, RPc, so called because the D N A around the transcription start site remains fully double-stranded or "closed". During the second step, the RPc complex isomerizes to an open complex, RPo This involves massive rearrangements in RNAP that drive the separation of the D N A strands near the start site so that the promoter is 16 Figure 6. Minimal model for transcription initiation. Transcription initiation begins with the reversible binding of RNAP (R, blue) to the promoter (P) to form a closed complex, RP C . The a subunit (dark blue) directs RNAP to promoters through sequence-specific interactions with DNA bases in the -35 and -10 elements (white boxes). Sequence-specific interactions between the ccCTDs with UP elements, or non-specific interactions between the aCTDs and upstream DNA contribute to R P C formation. R P C complexes rearrange to form open complexes, R P 0 . This isomerization drives the separation of the DNA strands from the upstream edge of the -10 element past the start site to expose the bases of the template strand. In the presence of NTPs, R P Q can initiate transcript formation (red), jettisoning the rj subunit after RNAP has entered a phase of processive elongation, R E . Evidence for two closed complexes and/or two open complexes has been found at some promoters. These complexes differ in the extent to which RNAP protects the promoter from DNase I cleavage while the complexes are closed, and the size of the denatured region while the complexes are open. 17 locally single-stranded or "open". Remarkably, a fragment of E. coli RNAP comprising (3' 1 -314 and 02-3 is capable of promoter melting (Young et al, 2004). This result eliminates the possibility of promoter unwinding driving D N A strand separation, but is consistent with base-flipping of the upstream portion of the -10 element followed by the passive trapping of single-stranded D N A by 02.3. In the presence of NTPs, RNAP can begin synthesizing RNA. RNAP can repetitively initiate short, abortive transcripts. The transition from an initiating complex to an elongating complex, R P E , occurs with the cessation of abortive transcription and promoter clearance. Additional intermediates have been observed at other promoters and sometimes these additional steps are critical for the regulation of a particular gene, but the pathway presented below is a minimal overview of the process. R + P ^ R P C <-*• RPo + NTPs <-» R E The promoter sequence and extrinsic variables (for example, accessory proteins, D N A supercoiling, temperature, osmolality and small molecules) affect the rate constants of individual steps and set the thermodynamic parameters governing initiation. Transcription factors work by affecting the rates at which RNAP binds, isomerizes or enters into a processive phase of elongation; activators accelerate these rates and repressors slow them down. 3.4 Repression. Repressors reduce the number of transcripts initiated from a particular promoter. At many promoters, the mechanism of repression is relatively straightforward. Some illustrative examples are presented here to contrast with the principles underlying activation. The simplest is binding site competition or steric hindrance (Fig. 7A), the classic example being LacI repression of the lac promoter (Meuller-Hill, 1996). In such cases a high affinity binding site for the repressor overlaps with the consensus promoter elements to which RNAP binds, and the repressor prevents the interaction of RNAP with the promoter. In some cases ternary Lacl-RNAP complexes form (Choy et al, 1995; Lee and Goldfarb, 1991; Straney and Crothers, 1987), but a rigorous thermodynamic treatment of the problem was consistent with the original simple competition model (Schlax et al, 1995). At other promoters, multiple repressors bind 18 A. Binding site competition. LacI at /_cUV5 jtt B. Occlusion by looping. GalRat_a/Pl C. Interference with post-recruitment steps. SpoOA at abrB p4atA2c D. Ternary complex destabilization. Spx at hmp Spx E. Open complex destabilization. DksA and ppGpp at rrnB PI Figure 7. Simple repression mechanisms. (A) At the lac\JV5 promoter, the lac repressor (purple) binds between the -35 and -10 elements to prevent association of RNAP with the core promoter elements. (B) At the galPl promoter, GalR (azure) tetramerization forms a DNA loop. This promoter topology prevents the formation of a productive closed complex. (C) Other repressors interfere with steps following the closed complex formation. At the abrB promoter, SpoOA~P (orange) stalls transcription initiation by preventing open complex formation, while p4 (green) interacts with the aCTD of RNAP to over-stabilize RNAP at the A2c promoter, preventing promoter clearance. (D) At the hmp promoter, interactions between Spx (yellow) and the aCTD of RNAP dissociate ternary ResD(red)-RNAP-/zwp complexes by disrupting ResD-aCTD interactions. (E) At the rrnBPl promoter, DksA (black) and ppGpp rearrange the active site through the secondary channel of RNAP to further destabilize open complexes and force the dissociation of RNAP. See text for references. 19 to distant sites forming a loop that occludes RNAP from the promoter (Fig. 7B). GalR represses transcription from gal?\ by this means (Choy and Adhya, 1996). Sometimes repressors interfere with the initiation process at steps following RNAP-promoter association (Fig. 7C). For example, at the abrB gene SpoOA~P prevents D N A strand separation without displacing RNAP (Greene and Spiegelman, 1996), and at the A2c promoter the protein p4 over-stabilizes RNAP preventing promoter clearance (Monsalve et al, 1996a; Monsalve et al, 1996b). Two other examples bear mentioning since they interfere with post-binding steps but result in the displacement of RNAP from the promoter. Spx destabilizes ternary complexes of RNAP, ResD and the hmp promoter without impairing ResD-hmp complex formation (Nakano et al, 2003). Spx appears to disrupt ResD-aCTD interactions that are required to maintain KNAV-hmp association (Fig. 7D). DksA binds to RNAP and amplifies the effect of ppGpp to further reduce open complex lifetime at rrnB PI (Paul et al., 2004a; Perederina et al., 2004). 3.5 Activation. At some promoters, RNAP requires accessory factors to initiate transcription. In many cases, these accessory factors or activators appear to compensate for sub-optimal promoter sequence that prevent RNAP-promoter interaction or stall RNAP at an unproductive intermediate. Perhaps the simplest activation mechanism is recruitment in which poor RNAP binding can be improved by adhesive interactions between a DNA-bound transcription factor and RNAP. The catabolite activator protein, CAP, activates transcription from the lac operon by recruiting RNAP via specific interactions between CAP and aCTD of RNAP (Fig. 8 A) (Busby and Ebright, 1999; Lawson et al, 2004; Malan et al, 1984). In other cases, activators may elicit a conformational change in the promoter to align promoter elements. In the presence of their ligand, MerR-type activators underwind the >19 base pair spacer separating the -35 and -10 elements allowing o to make productive contacts to both (Fig. 8B) (Heldwein and Brennan, 2001; Summers, 1992). At the ilvGMEDA operon promoter, IHF binding to a remote site constrains a superhelix-induced alternative D N A structure (Fig. 8C). This supercoiling-induced D N A destabilization is transmitted to the next most labile site, the-10 element, stimulating D N A strand separation and open complex formation (Sheridan et al, 1998; Sheridan et al, 2001). Other activators, notably Xcl at P R M , stimulate transcription by increasing the transcript 20 A. Recruitment. CAP at lac CAP * D B. Conformational change of promoter DNA MerR at PT Hg(H) C. DNA structural transmission of supercoiling-induced duplex destabilization. IHF at ilvGMEDA IHF D. Stimulation of post-recruitment steps. X cl at P R M E. Pre-recruitment and DNA scanning. SoxS at zwf Figure 8. Simple Activation Mechanisms. (A) At the lac promoter, interactions between CAP (red) and the aCTD improve the association of RNAP and the promoter. (B) At Pr, MerR (green) and RNAP form an inactive R P C in the absence of mercury in part due to the 19 base pair spacing separating the core promoter elements. The binding of Hg(II) by MerR induces a conformational change which distorts the promoter DNA allowing RNAP to contact both the -35 and -10 elements to form a productive RP C . (C) At the ilvGMEDA operon, IHF (azure) binds to a destabilized DNA element. By constraining this superhelix-induced distortion, IHF forces the duplex-destabilizing strain to the -10 element, promoting R P Q formation. (D) At the A P R M promoter, Xcl (purple) stimulates the isomerization of R P C complexes to R P Q complexes. (E) SoxS (yellow) binds directly to the aCTD to direct RNAP to composite promoters with SoxS binding sites upstream of the core promoters like zwf. See text for references. 21 initiation rate without affecting the initial interaction of RNAP with the promoter (Fig. 8D) (Hawley and McClure, 1982). This is the mechanism by which SpoOA~P appears to stimulate transcription at spoIIG promoter (Bird et al, 1996). Finally, activators like SoxS and MarA may interact directly with free RNAP and so direct RNAP to a specific subset of promoters bearing composite SoxS-RNAP or MarA-RNAP binding sites (Fig. 8E) (Dangi et al, 2004; Griffith and Wolf, 2004; Martin et al, 2002; Shah and Wolf, 2004). 4 Transcription Initiation at the spoIIG Promoter. 4.1 Characteristics of the spoIIG Promoter. The spoIIG promoter controls expression of an operon encoding a pro-o protease (spoIIGA) and pro-cjE (spoIIGB); a E is required for expression of stage III genes in the mother cell during sporulation. The sequence of the spoIIG promoter is shown in Fig. 9. Although the spoIIG promoter is transcribed by R N A P o A and contains a consensus -35 element (TTGACA) and a near consensus -10 element (TATAcT) for a A , two features of the promoter are unusual. First, the transcription start site is located 2 base pairs further downstream from the -10 element than at a consensus promoter. It is not obvious what prevents RNAP from initiating at -2 (where transcription would be expected to initiate based on the distance from the -10 element), nor is it clear what regulatory consequences this preference has on transcription initiation. It may be notable that at ribosomal RNA genes in E. coli, RNAP also initiates transcription 2 extra base pairs downstream of the -10 element (Paul et al, 2004b), and that transcription of these genes and spoIIG is stimulated by the presence of initiating nucleotides, or iNTPs in vitro (Gaal et al, 1997). It has been proposed that at rrnB PI , iNTPs might drive RNAP from an inactive to an active state (Lew and Gralla, 2004a; Lew and Gralla, 2004b), but the promoter sequence imposing iNTP stimulation has not been identified. Second, instead of the optimal 17 base pair spacer between the -35 and -10 elements, 22 base pairs separate these elements at the spoIIG promoter. RNAP can tolerate some deviation in spacer length as promoters with 16 or 18 base pairs separating the two primary sequence elements are transcribed, but further deviation prevents RNAP from initiating transcription 22 -100 C T T C C T C G A C A A A T T A A G C A G A T T T C C C T G A A A A A T T G T A •< OA box 1.1 OA box 1.2 T T T T C C T C T C A A C A T T A A T T G A C A G A C T T T C C C A C A G A G C -< OA box 2.1 OA box 2.2 • • I ** • +20 T T G C T T T A T A C T T A T G A A G C A A G A A G G G G A A C A G C G T G A G - C = K = 3 C = H i Figure 9. The spoIIG promoter. (A) The spoIIG promoter sequence from -100 to +20 is shown. The -35 and -10 elements are in bold, the transcription start site (+1) is denoted by a bent arrow. Dots mark every 10 th base. Four SpoOA binding sites or OA boxes are underlined by the orange arrows. The promoter-distal OA boxes are termed the site 1 OA boxes, while the promoter-proximal OA boxes are termed the site 2 OA boxes. The sub-sites are denoted OA boxes 1.1, 1.2, 2.1 and 2.2 as indicated. Note that OA box 2.2 overlaps entirely with the -35 element. (B) A schematic of the same region of the spoIIG promoter. OA boxes are indicated by the orange arrows, the -35 element by the dashed black box, the -10 element by the black box, and the transcription start site by the bent arrow. 23 (Ayers et al, 1989; Mulligan et al, 1985; Stefano and Gralla, 1982). This would seem to imply a fairly strict spatial arrangement of 0 4 relative to 0 2 with respect to the helical phasing of the promoter elements, and would necessarily preclude RNAP from contacting the -35 and -10 elements simultaneously at spoIIG. Overlong spacing at this promoter forms the basis of the requirement for Spo0A~P in activating transcription (McLeod and Spiegelman, 2005), in no small part because a binding site for SpoOA overlaps with the -35 element. 4.2 Regulatory Inputs at the spoIIG Promoter. Four factors control transcription initiation by RNAP at spoIIG, two negatively and two positively. 4.2.1 SinR. One negative regulator is SinR, a D N A binding protein that cooperates with other transition-state regulators like AbrB and ScoC to prevent the inappropriate expression of a large number of genes during exponential growth, and so coordinates sporulation with alternative stationary phase phenomena (Strauch and Hoch, 1993) including motility (Smith, 1993), alkaline protease expression and competence development (Mandic-Mulec et al, 1992), and biofilm formation (Kearns et al, 2005). SinR prevents sporulation by repressing the spoOA, spoIIE, spoIIA and spoIIG genes. At spoIIG, SinR prevents initiation ostensibly by converting the RNAP-promoter complex into a transcriptionally inactive form. However, since SinR does not displace RNAP from the spoIIG promoter, excess Spo0A~P can overcome SinR inhibition (Cervin et al, 1998a). 4.2.2 Soj. The other negative regulator is Soj, or SpoOJA. Soj is a ParA homolog (Ogasawara and Yoshikawa, 1992) that inhibits sporulation by directly repressing transcription of the spoIIE and spoIIG promoters (Cervin et al, 1998b). Soj physically associates with SpoOA-activated spoIIE, spoIIG, spoIIA and spoOA genes in vivo to prevent their expression (Quisel and Grossman, 2000; Quisel et al., 1999), and dissociates uninitiated complexes at the spoIIG 24 promoter in vitro (Cervin et al, 1998b). The antagonistic relationship between Soj and SpoOA is complicated by the observation that Soj has no effect on SpoOA-repressed abrB gene expression (Cervin et al, 1998b). It appears that Soj has two properties: a non-specific, low affinity inhibition, and a specific high-affinity inhibition that depends on SpoOA activation (McLeod and Spiegelman, 2005). Such evidence is consistent with the idea that Soj acts on RNAP rather than directly on activated SpoOA to oppose transcription of these genes, but the means by which it promotes the dissociation of RNAP has not been addressed. 4.2.3 iNTPs. The initiating nucleotides ATP and GTP play an unexpected role in promoting transcript formation. During single-round in vitro transcription assays using the spoIIG promoter, no transcripts are observed unless RNAP and SpoOA~P are pre-incubated with iNTPs prior to heparin challenge (Bird et al., 1996) Heparin is an anionic polymer that displaces uninitiated RNAP from promoters and prevents re-binding (Pfeffer et al., 1977). RNAP-spoIIG complexes that have not synthesized at least an RNA trimer do not survive heparin challenge (Bird et al., 1996). Heparin sensitivity until relatively late stages of transcription initiation is not uncommon for B. subtilis RNAP (Rojo et al, 1993; Voskuil and Chambliss, 2002; Whipple and Sonenshein, 1992). As alluded to in section 4.1 of the Introduction, an analogous situation exists at the rrnB PI promoter in E. coli (and likely at all promoters of ribosomal RNA genes). In contrast to complexes formed at many E. coli promoters, open but uninitiated complexes formed at rrnB PI rapidly dissociate following heparin challenge. At these promoters, iNTPs have been proposed to stabilize these heparin-sensitive complexes (Gaal et al, 1997), or by increasing the number of complexes capable of initiating transcription by correcting a distortion in RNAP structure imposed by some aspect of the rrn promoter sequence (Lew and Gralla, 2004a). Compelling data have indicated that the iNTP-dependence of rrnB PI transcription underlies in part the growth-phase dependent regulation of these promoters (Murray et al, 2003). Translation has been implicated as a major cellular consumer of purine nucleotides and the intracellular concentration of ATP and GTP appears to serve as a feedback signal for homeostatic control of rRNA synthesis (Schneider et al, 2002). One might speculate that a similar mechanism could contribute to limit transcription of the spoIIG and spoIIE genes to 25 conditions during which translation is reduced and the concentration of purine nucleotides/iNTPs is high. However such a model would need to accommodate the observation that when GTP levels are elevated, the global transcriptional regulator CodY- prevents sporulation (Ratnayake-Lecamwasam et al, 2001). 4.2.4 SpoOA. This thesis is primarily concerned with the mechanism by which activated SpoOA stimulates transcription from the spoIIG promoter. SpoOA is a transcription factor that directly controls the transcription of 121 genes (Molle et al, 2003) and indirectly controls the transcription of over 12% of the B. subtilis genome (Fawcett et al, 2000). Of particular importance, Spo0A~P represses abrB, indirectly relieving repression of spoOH (Fig. 2) (Greene and Spiegelman, 1996; Strauch et al, 1990). The product of the spoOH gene, o H , is required for transcription of the phosphorelay components kinA, spoOF and the sporulation-specific spoOA promoter (Predich et al, 1992). Spo0A~P also positively regulates its own synthesis from its aH-dependent sporulation-specific promoter (Strauch et al, 1992), and the transcription of kinA (Molle et al, 2003) and spoOF (Bai et al, 1990). By stimulating the production of SpoOA and phosphorelay components, this positive feedback loop increases the intracellular concentration of phosphorylated and active SpoOA. Upon reaching a threshold concentration, Spo0A~P stimulates the transcription of the spoIIG and spoIIA operons which begin the mother cell and forespore-specific programs of gene expression (Chung et al, 1994; Fujita et al, 2005). Relative to most response regulators, native SpoOA and dozens of mutants have been intensively studied (Table 1). In the last several years four crystal structures of SpoOA domains have been published, including two of its N-terminal receiver domain, Spo0AN, (Lewis et al, 1999; Lewis et al, 2000b) and two of its C-terminal output domain, SpoOAc (Lewis et al, 2000a; Zhao et al, 2002). No crystal structure of the intact protein has been solved. SpoOAn shares the conserved receiver domain structure (Fig. 10) and conformational shifts in response to phosphorylation (Lewis et al, 1999) that stimulate dimer formation in solution (Lewis et al, 2002). The phosphotransferase SpoOB and aspartate phosphatase SpoOE interact with the receiver domain, contacting residues flanking the site of phosphorylation (Stephenson and Perego, 2002). 26 Table 1. Phenotypes Associated with spoOA Mutations. MUTATION LOCATION PHENOTYPE REFERENCE N12K pi-al loop sofl02, suppressor of spoOF (Spiegelman et al., 1990) E14K al sofl07, suppressor of spoOF (Spiegelman a/., 1990) E14A al so/115, suppressor of spoOF (Spiegelman et al., 1990) P60S P3-a3 loop sofU8, suppressor ofspoOF (Spiegelman etal, 1990) D92Y a4 sofll4, suppressor ofspoOF and spoOA9V (Spiegelman et al, 1990) Q121R a5 sofl08, suppressor of spoOF (Spiegelman et al., 1990) F105S P5 sosll8, suppressor of sofl 18 (Spo") (Spiegelman et al, 1990) D56N P3-a3 loop Spo" (Burbulys etal, 1991) P60S p3-a3 loop coil, inappropriate sporulation (Olmedo etal, 1990) A87V P4-a4 loop coil, inappropriate sporulation (Olmedo etal, 1990) Q90K p4-a4 loop coil 5, inappropriate sporulation (Olmedo etal, 1990) A(H61-D75) Aa3 sad57, phosphorylation independent (ketone/ al, 1993) A(L62-N81) A(a3-p4) sad67, phosphorylation independent (Ireton etal, 1993) A(H61-N81) A(a3-P4) sad76, phosphorylation independent (Ireton etal, 1993) A(D75) a3-P4 loop sad54, phosphorylation independent (Ireton etal, 1993) D10Q PI Spo" (Green etal, 1991) D56Q P3 Spo" (Green etal, 1991) A(E20-D28) A (a l -P2) A209, phosphorylation independent (Green etal, 1991) A(E14-E47) A (a l -P2) A267, phosphorylation independent (Greene^/., 1991) H162R aA suv4, suppressor o£spoOA9V (Perego et al, 1991) Suppressor of S250H (Schmeisser et al, 2000) L174F aB suv3, suppressor of spoOA9V (Perego etal, 1991) Suppressor of S250H (Schmeisser et al, 2000) D200N aC Fails to repress abrB or to activate spoIIA (Lewis et al, 2000a) E221A aD-aE loop Allele-specific suppression of o A (R355A) (Kumar et al, 2004b)) E231D aD Fails to repress abrB or to activate spoIIA (Lewis et al, 2000a) G227R aE Fails to activate aA-dependent promoters (Hart and Youngman, 1998) I229A aE Lower activation rjA-dependent promoters (Buckner et al, 1998) D230A aE Lower activation oA-dependent promoters (Buckner e/a/., 1998) S231F aE Suppresses oA(H359) Spo" phenotype (Buckner et al, 1998) I232A aE Lower activation oA-dependent promoters (Buckner et al, 1998) S233P aE Fails to activate oA-dependent promoters (Hatt and Youngman, 1998) F236S aE Fails to activate aA-dependent promoters (Hatt and Youngman, 1998) T239A aE Fails to activate oA-dependent promoters (Kumar et al, 2004a) V240A aE Fails to activate cA-dependent promoters (Hatt and Youngman, 1998) V240G/K265R aE Fails to activate aA-dependent promoters (Hatt and Youngman, 1998) A(226-243)/V8A A(aE) Activates spoIIA, fails to activate spoIIG (Kumar et al, 2004a) S250H aF Fails to repress abrB or to activate spoIIA (Schmeisser et al, 2000) A257V aF Represses abrB, fails to activate spoIIA (Perego etal, 1991) A257E aF Represses abrB, fails to activate spoIIA (Perego etal, 1991) D258V aF Represses abrB, fails to activate spoIIG or spoIIA (Rowe-Magnus et al, 2000) L260V aF Represses abrB, fails to activate spoIIG or spoIIA (Rowe-Magnus et al, 2000) A(I252-S267) A(aF) Spo" (Ferrari et al, 1985) 27 Figure 10. Structure of the SpoOAN receiver domain. The receiver is presented from the side (left) and from the top (right) to display the overall fold topology. SpoOAN is a member of the ap class, with a 3-layer (apcc) sandwich architecture in a Rossman fold and has a prototypical response regulator receiver domain structure. The chain is colour ramped from the N-terminus (dark blue) to the C-terminus (red) and the secondary structural features labeled (al-a5 and Pl-p5). This structure is of the SpoOAN from Bacillus stearothermophilus which is similar to that of SpoOAN from B. subtilis (98 identical residues and 24 conservative substitutions out of 122 amino acids). The figure is based on PDB file 1QMP deposited by Lewis et al. (1999) in the RCSB protein data bank (http://pdbbeta.rcsb.org/pdb/Welcome.do) and was constructed using PyMOL (Delano Scientific). 28 Figure 11. Structure of the SpoOAc activation domain. SpoOAc is a member of the mainly a class of proteins, with an orthogonal bundle architecture that lacks detailed homology with other protein structures. The chain is colour ramped from the N-terminus (dark blue) to the C-terminus (red) and the helices are labeled (aA-ocF). This structure is of SpoOAc from Bacillus stearothermophilus which is similar to that of SpoOAc from B. subtilis (62 identical residues and 6 conservative substitutions out of 68 amino acids). The aE helix and flanking loops (orange) comprise the <5A activating region. The helix-turn-helix motif includes aC and aD. Amino acids in the aD helix recognize the OA box sequence. The figure is based on PDB file 1FC3 deposited by Lewis et al. (2000) in the RCSB protein data bank (http://pdbbeta.rcsb.org/pdb/ Welcome.do) and was constructed using PyMOL (Delano Scientific). 29 Mutations in spoOA suppressing a deletion of the spoOF gene were isolated in the receiver domain (Spiegelman et al, 1990); some of these sof mutants were improved substrates for direct phosphorylation by KinA while others enhanced the stability of Sof-RNAP-spo/JG complexes (Cervin and Spiegelman, 1999). It was suggested that one sof mutant in particular (sof!14, an Asp92Tyr substitution) enhanced a natural interaction between the phosphorylated receiver of SpoOA and R N A polymerase, but the results are equally consistent with this mutation stabilizing DNA-bound SpoOA~P dimers. Members of another class of SpoOAN mutants, the sad mutants, are constitutively active deletions (Ireton et al., 1993). This has been interpreted as evidence that the receiver domain normally inhibits the output domain, but the observation that a substitution mutant in the same region acts as a null mutation complicates the issue (Cervin and Spiegelman, 2000). SpoOAc binds DNA, stimulates transcription (Grimsley et al., 1994; Rowe-Magnus and Spiegelman, 1998a) and when over-expressed in vivo supports stage II transcription (Florek et al., 2002). It binds as a dimer to a direct repeat 7 base pair consensus sequence of 5'-T G T C G A A - 3 ' (Zhao et al, 2002) and contains six a-helices in an unprecedented fold, including a helix-turn-helix motif (Fig. 11) to which mutations involved in D N A binding have been mapped (Hatt and Youngman, 2000). SpoOAc also contains a central helical bundle and a protruding mobile helix (aE) to which the majority of mutations involved in SpoOA-dependent, aA-dependent transcription map (Buckner et al, 1998; Hatt and Youngman, 1998). Genetic evidence for the interaction of SpoOA and a A has been strengthened by the identification of mutations in a A that affect only transcription from the same class of promoters (Baldus et al, 1995; Buckner et al, 1998; Schyns et al, 1997), and a corresponding suppressor of some of these mutations lies within the mobile aA-activating region (Buckner et al, 1998). Particularly compelling mutations in SpoOAc, spoOA9V(an Ala257Val substitution) as well as valine substitutions of Asp258 and Leu260, prevent activation of spoIIG and spoIIA without compromising repression of abrB (Perego et al, 1991; Rowe-Magnus et al, 2000). Suppressors of the spoOA 9 V mutation map to the opposite face of the protein; however, the tandem dimerization of SpoOAc brings the two faces into apposition (Zhao et al, 2002), suggesting that the suppressors compensate for altered packing at the dimer interface. 30 4.3 A Working Model for Activation of the spoIIG Promoter b y Spo0A~P. At the spoIIG promoter Spo0A~P binds to OA boxes located between -96 and -81 and between -53 and -37. The OA box closest to the transcription start site overlaps entirely with the -35 element, and places the downstream-most SpoOA~P molecule in a position to interact with RNAP via the protruding helix and flanking loops of SpoOA and amino acids in a A 4. Mutations in all four OA boxes reduce spoIIG expression in vivo (Baldus et al, 1994; Satola et al, 1991; Satola et al, 1992). SpoOA~P stimulates transcription initiation without improving RNAP binding by subtly changing how RNAP interacts with the promoter (Bird et al, 1996). Although RNAP cannot denature the promoter by itself, RNAP plus activated SpoOA separates the D N A strands between -14 and -3, and artificially denaturing this region bypasses the requirement for activated SpoOA (Rowe-Magnus and Spiegelman, 1998b). Thus SpoOA cooperates with RNAP to form an open complex, but precisely how it works is incompletely understood. At its outset, the specific aim of my thesis was to better understand how SpoOA and RNAP cooperated to drive open complex formation. In particular, the nucleation of D N A strand separation and the role SpoOA played in this process was unknown. More problematic still were the events between the initial binding of RNAP to the spoIIG promoter and the formation of the first stable open complex, crucial stages in the mechanism that were completely inaccessible. In general, it was hoped that by tackling these issues I could address the fundamental problem of how transcriptional activators work to stimulate post-binding steps during transcription initiation. In this thesis, I show that SpoOA~P acts prior to denaturation of the D N A strands, and has no effect on transcription from D N A templates in which the -10 element is artificially denatured to expose the non-template strand. While activated SpoOA is necessary for R N A polymerase-mediated D N A strand separation on a wild type template, I found that it did not reduce the energy required to denature the promoter. These two results suggest that SpoOA exerts its effect on RNA polymerase prior to the formation of an open complex. I present evidence for a closed intermediate complex formed by RNAP, SpoOA~P and the spoIIG promoter. To isolate the 31 effect of SpoOA~P on events prior to D N A strand separation at the spoIIG promoter I used D N A fragments that contained only promoter sequences 5' to the -10 element in electrophoretic mobility shift assays (EMSAs) . R N A P bound to these fragments readily and the RNAP-spoIIG complex recruited SpoOA~P. This complex was stabilized by Spo0A~P. In addition I investigated the effect of truncating the upstream portions of the spoIIG promoter that contain a pair of SpoOA binding sites at approximately -90 relative to the transcription start site, and found that this D N A inhibited the binding of R N A polymerase. Strikingly, SpoOA~P binding depended on the presence of the D N A upstream of the OA boxes, suggesting a role for the aCTDs in transcription at this promoter. Finally, SpoOA~P binding to the promoter played two distinct roles. Promoter-distal bound SpoOA~P appeared to increase the local concentration of SpoOA~P available to bind to promoter-proximal SpoOA binding sites, whereas promoter-proximal bound SpoOA~P stabilized and modified pre-formed R N A polymerase-^oT /G complexes. These effects account for the transcription activation of the spoIIG promoter by SpoOA~P. 32 EXPERIMENTAL PROCEDURES Standard molecular biology techniques (precipitation of D N A by ethanol and sodium acetate, extraction of protein from nucleic acids by phenol, digestion of D N A with restriction fragments, ligation of restriction fragments, sodium dodecyl sulfate polyacrylamide gel electrophoresis of proteins, separation of D N A fragments by agarose gel electrophoresis and polyacrylamide electrophoresis) were performed as described in Molecular Cloning: A Laboratory Manual (Sambrook etal, 1989). 1 Manipulation of DNA and Oligonucleotides. 1.1 Preparation of spoIIG and IIG17 Promoter Fragments. The spoIIG and IIG17 templates were isolated from the plasmids pUCIIGtrpA (Satola et al., 1991) and pUCIIGl 7trpA (gift of Brett McLeod) respectively as 415 base pair fragments by digestion with PvuII and BamHl. A l l plasmids used in this thesis and their sources are presented in Table 2. These restriction fragments contained the spoIIG promoter sequence from -100 to +134 and included promoter-distal and promoter-proximal SpoOA binding sites, but lacked the trpA terminator. The promoter fragment from p\JCIIG17trpA differed from that from pUCIIGtrpA in that 17 base pairs rather than 22 base pairs separated the -35 and -10 elements. Both fragments gave rise to run-off transcripts of 131 base pairs. The fragments were isolated by electrophoresis, recovered using a QIAquick gel extraction kit (Qiagen), and stored in 10 mM HEPES (pH 7.9), 20 m M potassium acetate, and 0.1 m M EDTA at 4 °C. The concentration of template fragments was determined by measuring absorbance at 260 nM. 1.2 Heteroduplex Templates. Heteroduplex templates were created essentially as described earlier (Rowe-Magnus and Spiegelman, 1998b). 33 Table 2. Plasmids Used in this Thesis. PLASMID DESCRIPTION REFERENCE or SOURCE +/-pUCIIGtrpA pUCIIG17trpA pGEM-T pBluescriptSK pSS3 p S K / / G + pSKJ/G7V pSK//G2 + /" pSK//G3 + / " pSKZ/Go^7" pSKIIGf-pMCOA pTMOA pHisOAC pETOFHis pUCIIG2.1Up pUCIIG2.1Down 240 bp spoIIG promoter fragment in pUC19. pUCIIGtrpA with 17 bp spacer. PCR cloning vector. Phagemid vector for ssDNA production. AluI-BamHI spoIIG fragment with 4 bp mutated in pGEM-T. Hindlll-BamHI spoIIG fragment in pBluescriptSK47". Hindlll-BamHI spoIIG fragment with 12 bp mutation in pBluescriptSK+ /\ Hindlll-BamHI spoIIG fragment with 8 bp mutation in pBluescriptSK+/". Hindlll-BamHI spoIIG fragment with 4 bp mutation in pBluescriptSK+/". Hindlll-BamHI spoIIG fragment with 6 bp mutation in pBluescriptSK+/". Hindlll-BamHI spoIIG fragment with 2 bp mutation in pBluescriptSK+ /\ pET16b-based SpoOA over-expression plasmid. pET16b-based SpoOAc over-expression plasmid. pET16b-based SpoOAc-His over-expression plasmid. pET20b-based SpoOF-His over-expression plasmid. pUCIIGtrpA with near consensus 2.1 OA box. pUCIIGtrpA with 2.1 OA box null. (Satola, etal, 1991) (McLeod, et al, 2005) Promega Stratagene This work. (Rowe-Magnus et al, 1998a) (Rowe-Magnus et al, 1998a) This work. This work. This work. This work. (Cervin et al, 1999) (Zhao et al, 2002) This work. (Tzeng etal, 1997) This work. This work. 34 1.2.1 Construction of pSKSIIGT and pSK3I/G\ The plasmids pSK3/ /G + and pSK3//G" were created in several steps. First, a PCR product was generated using Taq D N A polymerase (Invitrogen), the template pUCIIGtrpA, a mutagenic primer, SS3, complementary to the non-template strand, and a downstream primer, IIG2X, that anneals to the template strand adjacent to the BamHI site. A l l primers used in this thesis and their sequences are presented in Table 3. The primer SS3 contained nucleotides that were identical to the template strand from -14 to -11. The PCR product was cloned into the pGEM-T vector (Promega) to create plasmid pSS3, and the insert was verified by sequencing. The Hindll l-Alul fragment was isolated from pUCIIGtrpA and ligated to the AluI-BamHI fragment from pSS3, and the ligation mixture used as a template for PCR by using IIG2X and an upstream primer, IIGA, that anneals to the non-coding strand adjacent to the Hindlll site. The resulting product was ligated into pGEM-T to create pSS3//G. The E. coli strain DH5a was used for all transformations and plasmid preparations were performed as described by Sambrook et al. (Sambrook et al, 1989). A l l strains used in this thesis and their origins are presented in Table 4. The plasmid pSS3/JG was digested with.Hindlll and BamHI, and the promoter-bearing fragment ligated into pBluescript S K + and pBluescript SK" (Stratagene) digested with the same enzymes to create pSK3/ /G + and pSK3/JG\ 1.2.2 Construction of pSKM/G* and pSKHIG^. The pSK/JG plasmid was used as the template in a PCR to create pSK677G and pSK77/G . Reactions were composed of primer pairs IIG6NT and IIG6T or IIG7NT and IIG7T, along with 2.5 m M dNTPs, lx Pfu Turbo reaction buffer, and 1 unit of Pfu Turbo D N A polymerase (Stratagene) and subjected to 18 cycles of 30 s at 95 °C, 1 min at 55 °C, and 6 min at 68 °C. The PCR products were treated with 10 units of Dpnl for 1 h and ethanol precipitated after the addition of 100 ng salmon sperm DNA. The D N A was re-suspended in water and used to transform E. coli DH5a. Mutations in plasmids recovered from transformants were verified by sequencing. Table 3. Primers Used in this Thesis. PRIMER SEQUENCE (5 ' to 3') SS3 G A G C T T G C T T T A T A C T T A T G A A G C IIG2X G G G G A T C C T C T C G A G T C A EGA A A G C T T A T C G A C A A A T T A A IIG6NT C C C A C A G A G C T T G C T T A T A T G A T A T G A A G C A A G A A G G G IIG6T C C C T T C T T G C T T C A T A T C A T A T A A G C A A G C T C T G T G G G IIG7NT C C C A C A G A G C T T G C T T A T T A C T T A T G A A G C A A G A A G G G IIG7T C C C T T C T T G C T T C A T A A G T A A T A A G C A A G C T C T G T G G G HisOACNT G G G G A G G A A G A A A C A T G C A C C A C C A C C A C C A C C A C G A G A G C A G C C A G C C HisOA CT GGCTGGCTGCTCTCGTGGTGGTGGTGGTGGTGCATGTTTCTTCC TCCCC M13R C A G G A A A C A G C T A T G A C C +50R T T T T C A C A T C T G A C T C C T T T C -J5R C T G C A A G C T C T G T G G G A A A G T C T G -19R A A G C T C T G T G G G A A A G T C T G T -23R T C T G T G G G A A A G T C T G T C A A T -27R T G G G A A A G T C T G T C A A T T A A T -31R A A A G T C T G T C A A T T A A T G T T G -35R T C T G T C A A T T A A T G T T G A G A G -80F G A T T T C C C T G A A A A A T T G T A T T -69F A A A T T G T A T T T T C C T C T C A A C A -58F T T C C T C T C A A C A T T A A T T G A C A IDownF A A G C T T A T C C A G A A A T T A A G G A G A T T T C C C T G A -15RK033 C T G C A A G C T C T G T G G G A A A C A G T G T C A A T T A A T 2.1 UpNT A A T T G T A T T T T C C T C T C G A C A T T A A T T G A C A G A C T T 2.1 UpT A A G T C T G T C A A T T A A T G T C G A G A G G A A A A T A C A A T T 2. JDownNT A A T T G T A T T T T C C T C A G A A G A T T A A T T G A C A G A C 2. IDownT G T C T G T C A A T T A A T C T T C T G A G G A A A A T A C A A T T Underlined bases are mutated relative to the spoIIG sequence, except for HisOACNT and His OA CT where the underlined bases have been introduced into the pTMOA plasmid. Table 4. Bacterial Strains Used in this Work. STRAIN G E N O T Y P E R E F E R E N C E or SOURCE E. coli DH5a [hsdRJ 7(rK'mK+) supE44 thi-1 rexAl gyrA (Nat) Invitrogen relAl A(lacZYA-argF) U169 JM101 [F' trad36proA+ proB+ lacf lacZbMl5/supE thi Invitrogen A(lac-proAB)] BL21(DE3) pLysS F" ompThsdSB (r_~mB~) gal dcm (DE3) pLysS Invitrogen (CamR) BL21 Star (DE3) F" ompT hsdS& (rB"mB") gal dcm rne!31 (DE3) Invitrogen B. subtilis JH642 tprC2 phe-1 J. Hoch, Scripps 37 1.2.3 Purification Single-stranded Plasmid DNA The wild type spoIIG promoter had previously been cloned into pBluescript S K + and pBluescript SIC to create pSK/7G + and pSKY/G\ The plasmids p S K l / / G + , pSKl/ /G", pSK2//G + , and pSK2//G" (called pSK12//G + , p S K l 2/7(7, pSK877G+ and pSK877G" earlier) had been constructed previously (Rowe-Magnus and Spiegelman, 1998b). The pSK + /" series of plasmids were transformed into E. coli J M 1 0 1 . Single colonies were picked from a freshly streaked plate and used to inoculate 100 pi of Luria-Bertani media supplemented with 100 pg/ml ampicillin in a 1.7 ml microfuge tube. The microculture was shaken at 180 rpm for 1 h at 37 °C before infection with 2 pi of M13K07 helper phage (~2 x 109 viral particles). Infected cultures were shaken at 180 rpm for 2 h at 37 °C before inoculation into 500 ml of Luria-Bertani media supplemented with 100 pg/ml ampicillin and 70 pg/ml kanamycin. Cultures were grown for 16 to 20 h at 150 rpm and 37 °C. Cells were removed by centrifugation (12,000 x g for 30 min), and 0.25 volumes of 20% PEGgooo/3.5 M sodium chloride was added to the supernatant. The solution was mixed gently by inversion and left overnight on ice. Phage particles were collected by centrifugation (12,000 x g for 10 min) and then re-suspended in 10.0 ml TE (10 mM Tris (pH 7.9), 0.1 m M EDTA). The phage particle suspension was phenol extracted until all protein was removed, and the single-stranded D N A ethanol precipitated. Single-stranded D N A was dissolved in 0.5 ml TE and its concentration determined by measuring absorbance at 260 nM. 1.2.4 Production of Heteroduplex Templates. To produce locally single-stranded promoters, 25 pg of complementary single-stranded DNAs were mixed and adjusted to 0.2 N sodium hydroxide in 50 pi. Samples were held at 37 °C for 15 min and then neutralized with hydrochloric acid. After 15 min at 37 °C. the annealed heteroduplex plasmids were precipitated with ethanol, re-suspended in 60 pi of lx PvuII buffer and digested with 30 units of PvuII and 30 units of BamHI. Fragments containing the heteroduplex promoters were isolated by electrophoresis, recovered using a QIAquick gel extraction kit (Qiagen), and stored in 10 mM HEPES (pH 7.9), 20 mM potassium acetate, and 38 0.1 m M EDTA at 4 °C. The concentration of template fragments was determined by measuring absorbance at 260 nM. 1.3 End-labeling of Restriction Fragments. The template strand of the spoIIG promoter was labeled by digesting 10 pg of the pUCIIGtrpA plasmid with 10 units of BamHI. The D N A was phenol extracted, ethanol precipitated, and dissolved in 20 pi lx Calf Intestinal Alkaline Phosphatase buffer (CIAP; Boehringer-Mannheim) with 20 units of CIAP to dephosphorylate the protruding 5' ends of the BamHI-linearized pUCIIGtrpA plasmid. After 90 min at 37 °C, the sample was brought to 100 pi and the enzyme inactivated by adding SDS and EDTA to final concentrations of 0.5% and 5 mM, respectively. The sample was phenol extracted 3x, the organic phase back-extracted with TE, and the D N A precipitated by adding 2 volumes of ethanol and 0.1 volumes of 3 M sodium acetate (pH 7.0) to prevent the precipitation of EDTA under acidic conditions. The D N A was re-suspended in 20 pi of lx Kinase Forward Buffer (50 m M Tris-Hydrochloride (pH 7.6), 10 m M magnesium chloride, 4 mM dithiothreitol (DTT), 0.1 m M spermidine and 0.1 m M EDTA; (Sambrook et al., 1989), including 575 pCi [y 3 2P]ATP (7000 Ci/mmol; GE Healthcare) and 5 units of T4 kinase. The labeling reaction was allowed to proceed for 30 min, and then the sample was digested for 2 hours with 30 units of PvuII in a 60 pi volume of lx PvuII buffer. Labeled fragments were separated by electrophoresis through a 5% non-denaturing polyacrylamide gel. The s/w//G-containing D N A fragments were located by autoradiography, and the gel slice containing the fragment excised. Gel slices were forced through a hole (punched with a 21 gauge hypodermic needle) in the bottom of a 0.7 ml microfuge tube, and collected by centrifugation (13,000 x g, 1 min) into a 1.7 ml microfuge tube into which the smaller microfuge tube had been nestled. The crushed gel fragments were suspended in 500 pi of acrylamide elution buffer (500 m M ammonium acetate, 10 mM magnesium acetate, 10 m M Tris (pH 7.9) and 1 mM EDTA), and the D N A eluted passively overnight at 37 °C. The resulting suspension was filtered through a 0.22 pm syringe-driven filter and the D N A was ethanol precipitated twice. The labeled spoIIG promoter fragments were re-suspended in 40 pi of 10 mM HEPES (pH 8.0), 20 mM potassium acetate and 0.1 mM EDTA, and their activity estimated by measuring the emitted Cerenkov radiation. 39 1.4 Oligonucleotide Labeling. Primers were ethanol precipitated at least twice to prevent inhibition of T4 kinase activity by residual ammonium ions. Quantitative labeling of primers was achieved by composing reactions of 20 pmol oligonucleotide, 1150 uCi [y 3 2P]ATP (7000 Ci/mmol; GE Healthcare) and 5 units of T4 kinase in lx Kinase Forward Buffer for a final volume of 20 pi. Reactions were incubated at 37 °C for 1 hour, brought to 200 pi with distilled, deionized water, phenol extracted once, and then chloroform extracted once. [y 3 2P]ATP was removed from the radiolabeled primers by ethanol precipitation. One pi 10 ug/ul glycogen was added, as were 2 pi 1 M magnesium chloride, 100 pi 7.5 M ammonium acetate and 1 ml ethanol and the samples were held at -70 °C overnight or on crushed dry ice for at least 1 hour. Samples were centrifuged (13,000 x g, 30 min), the supernatant decanted, and the precipitated primer washed by the addition of 300 pi 70% ethanol. The samples were centrifuged as above, the supernatant decanted and the radiolabeled primers air dried, re-suspended in 50 pi TE and stored at 4 °C. 1.5 Preparation of Labeled DNA Fragments. Radiolabeled oligonucleotides were used to create PCR products for use in DNase I footprinting and electrophoretic mobility shift assays. A l l fragments used in this thesis and the primers and templates used to prepare them are listed in Table 5. PCR were composed with 10 pmol labeled primer, 15 pmol unlabeled primer, 0.5 pmol template DNA, 2.5 mM dNTPs and 1 unit Pfu D N A polymerase in 50 pi of lx Pfu reaction buffer. The reactions were split into 25 pi aliquots and subjected to 30 cycles of 95 °C for 30 s, 55 °C for 1 min, and 68 °C for 1 min in a Biometra T Gradient fhermocycler (Montreal Biotech). Labeled fragments were separated by electrophoresis through a 5% non-denaturing polyacrylamide gel, located by autoradiography and excised. Gel slices were crushed as described as above, suspended in 500 pi of acrylamide elution buffer (500 mM ammonium acetate, 10 mM magnesium acetate, 10 mM Tris (pH 7.9) and 1 mM EDTA), and eluted passively overnight at 37 °C. The resulting suspension was filtered through a 0.22 um syringe-driven filter and the D N A was ethanol precipitated twice. Table 5. Primers Used to Prepare Radiolabeled Fragments Used in this Thesis. FRAGMENT UPSTREAM DOWNSTREAM TEMPLATE PRIMER PRIMER spoIIG (EMSA) IIGA IIG2X pUCIIGtrpA TUG (EMSA) IIGA -15R pUCIIGtrpA spoIIG (DNase) M13R +50R pUCIIGtrpA TUG (DNase) M13R -15R pUCIIGtrpA TIIG(-\9) IIGA -19R pUCIIGtrpA TIIG(-23) IIGA -23R pUCIIGtrpA TIIG{-21) IIGA -27R pUCIIGtrpA TIIG{-3\) IIGA -31R pUCIIGtrpA TIIG(-35) IIGA -35R pUCIIGtrpA (S0)TIIG -80F -15R pUCIIGtrpA (-69)TIIG -69F -15R pUCIIGtrpA (-58)TIIG -58F -15R pUCIIGtrpA TIIGlDown IDownF -15R pUCIIGtrpA TIIG2 IIGA -15RK033 pUCIIGtrpA TIIG2.1Up IIGA -15R pUCIIG2.1Up TIIG2.1Down IIGA -15R pUCIIG2.1Down 41 The labeled spoIIG promoter fragments were re-suspended in 40 pi of 10 m M HEPES (pH 8.0), 20 m M potassium acetate and 0.1 m M EDTA, and their activity estimated by Cerenkov radiation emitted. 2 Protein Purification. 2.1 SpoOA. The pET16b-based plasmid, pMCOA (Cervin and Spiegelman, 1999), was used to overexpress SpoOA in freshly transformed BL21(DE3) pLysS E. coli cells. An overnight culture (5% volume/volume) was used to inoculate 4 liters of Luria-Bertani broth supplemented with 100 pg/ml ampicillin and 34 pg/ml chloramphenicol. Cultures were grown at 30 °C until an OD600 of 0.9 was reached, at which point isopropyl-p-D-thiogalactopyranoside (IPTG) was added to a final concentration of 0.5 mM. Cultures were grown at 30 °C for 4.5 hours until the cells were harvested by centrifugation (12,000 x g, 15 min) and frozen on crushed dry ice. Cell pellets from 4 liters of culture were thawed on ice in 80 ml of Buffer A S P O O A (20 m M Tris (pH 8.3), 150 m M potassium chloride, 10 m M magnesium chloride, 1 mM EDTA, 1 mM phenyl methyl sulfonyl fluoride (PMSF) and 1 mM DTT) in the presence of 1 mg/ml lysozyme. Cell suspensions were sonicated on ice (1 Ox for 30 s at power level 3 using an Ultrasonic Processor X L ; Misonix) and cleared by centrifugation (12,000 x g, 30 min). Protein in the supernatant was precipitated by the addition of solid ammonium sulfate to 20% (weight/volume). After stirring to dissolve the ammonium sulfate, the precipitated proteins were removed by centrifugation (27,000 x g, 30 min) and the pellet discarded. The supernatant was adjusted to 50% ammonium sulfate by adding solid ammonium sulfate, centrifugation was repeated (27,000 x g, 30 min), and the pellet retained. The SpoOA-containing protein pellet was dissolved with 20 ml of Buffer B S P O O A (20 m M sodium phosphate (pH 8.0), 150 m M sodium chloride, 1 mM EDTA and 1 mM PMSF) and dialyzed against 2 liters of Buffer B S P O O A - Dialysates were filtered through 0.45 pm Millex filters (Millipore) and loaded onto a 10 ml heparin-agarose column (Sigma) at 1 ml/min. After washing off unbound protein, SpoOA was eluted with a 250-800 mM sodium chloride linear gradient in Buffer B S P O O A - Fractions were analyzed by SDS-PAGE, 42 and the SpoOA-containing fractions pooled and dialyzed against 2 liters of Buffer B S P O O A -Dialysates were diluted 3x with Buffer B S P O O A lacking sodium chloride and loaded onto a 10 ml D N A cellulose column at 1 ml/min. After washing off unbound protein, SpoOA was eluted with a 50-800 m M sodium chloride gradient in Buffer B S P O O A - Fractions were analyzed by SDS-PAGE, and the SpoOA-containing fractions pooled and dialyzed overnight against 2 liters of Buffer B B S P O O A - Dialysates were concentrated to 1 -2 mg/ml using Centricon Plus-80 ultrafiltration devices (Amicon Bioseparations; Millipore). Glycerol and DTT were added to 30% and 0.1 m M respectively, and aliquots of SpoOA stored at -70 °C. Protein concentrations were estimated by absorbance at 280 nm using a S S P O O A = 16,270 M^cm" 1 . 2.2 SpoOAc. The pET16b-based plasmid, pTMOA (Zhao et al, 2002) was used as template in a PCR along with the primer pair HisOACNT and HisOACT to create a plasmid pHisOAC that overexpresses N-terminally His-tagged SpoOAc. Mutagenesis was performed essentially as in Experimental Procedures 1.2.2. pHisOAC was transformed into BL21 Star (DE3) E. coli cells (Invitrogen). An overnight culture (5% volume/volume) was used to inoculate 4 liters of Luria-Bertani broth supplemented with 100 pg/ml ampicillin. Cultures were grown at 37 °C until an OD600 of 0.6 was reached, at which point IPTG was added to a final concentration of 1 mM. Cultures were grown at 30 °C for 4 hours. The cells were harvested by centrifugation (12,000 x g, 15 min), and the cell pellets frozen on crushed dry ice. Cell pellets were thawed on ice in 50 ml of Buffer A S P O O A C (20 mM Tris (pH 6.8), 1 m M EDTA, 1 mM phenyl methyl sulfonyl fluoride (PMSF) and 10 mM P-mercaptoethanol) in the presence of 1 mg/ml lysozyme. Cell suspensions were sonicated on ice as in Introduction, section 2.1, cleared by centrifugation (12,000 x g, 15 min) and the supernatant loaded directly onto a 10 ml heparin-agarose column (Sigma) at 1 ml/min. The column was washed with Buffer A S P O O A C until the absorbance at 280 nm dropped below 0.05, and then Spo0A c was eluted with a 150-800 mM sodium chloride gradient in Buffer A S P O O A C - Fractions were analyzed by SDS-PAGE, and the SpoOA-containing fractions pooled and dialyzed against 2 liters of Buffer B S P O O A C (10 m M Tris (pH 6.8), 300 m M sodium chloride and 0.5 m M PMSF). Spo0A c was further purified by 43 Ni-NTA (Qiagen) batch elution exactly as per the manufacturer's instructions. Fractions were analyzed by SDS-PAGE, and SpoOAc-containing fractions pooled and dialyzed for 2 hours against 2 liters of Buffer B S P O O A C to remove the imidazole. SpoOAc was concentrated to 20 mg/ml using an Amicon Ultra 15 centrifugal filter device with a molecular weight cut-off of 5000 kDa (Millipore), aliquotted and stored at -20 °C. Protein concentrations were estimated by absorbance at 280 nm using a S S P O O A C = 12,090 IVf'cm"1. 2.3 SpoOF. The pET20b-based plasmid, here called pETOFFfis (Tzeng and Hoch, 1997) was used to overexpress C-terminally His-tagged SpoOF. BL21(DE3) pLysS E. coli cells were transformed with pETOFHis and overnight cultures (5% volume/volume) were used to inoculate 4 liters of Luria-Bertani media supplemented with 100 pg/ml ampicillin. Cultures were grown until the OD 6 0o reached 0.6 at which point IPTG was added to a final concentration of 0.4 mM. Cultures were grown at 30 °C for 4 hours, when the cells were harvested by centrifugation (12,000 x g, 15 min) and frozen on crushed dry ice. Cell pellets were thawed on ice in 20 ml of Buffer A S p o oF (10 mM HEPES (pH 7.9), 10 mM magnesium acetate, 20 m M potassium acetate, 300 mM sodium chloride, 10 mM imidazole and 50 pg/ml PMSF) plus 1 mg/ml lysozyme and sonicated as in Introduction, section 2.1. Cell suspensions were cleared by centrifugation (12,000 x g, 15 min), and purified by Ni -NTA (Qiagen) batch elution as per the manufacturer's instructions. Fractions were analyzed by SDS-PAGE, and SpoOF-containing fractions pooled and dialyzed for 2 hours against 2 liters of Buffer AspooF to remove the imidazole. Protein concentrations were determined by Bradford assay exactly as per the manufacturer's instructions. Fractions were aliquotted and stored at -20 °C. 2.4 RNA Polymerase. RNA polymerase was purified from 25 liters (25 g cell pellet) of Bacillus subtilis JH642 cells harvested in mid-log phase growth in Luria-Bertani broth + 0.1% dextrose and frozen on crushed dry ice. The following procedure is a modification of that previously published 44 (Dobinson and Spiegelman, 1987). Cell pellets were thawed on ice in 35 ml of Buffer A R N A P (10 mM Tris (pH 7.9), 1 m M EDTA, 10 mM magnesium chloride, 1 m M p-mercaptoefhanol and 50 pg/ml PMSF) supplemented with 5 ml 2 M Tris (pH 7.9) and 250 mg lysozyme. Cell suspensions were split in two and each portion sonicated (8x for 30 s at power level 6 using an Ultrasonic Processor X L ; Misonix), and cleared by centrifugation (39,000 x g, 15 min). Three Dextran/PEGgooo phase separations with increasing concentrations of sodium chloride were performed to reduce the amount of contaminating protein exactly as described previously. The final PEG phase was diluted 1:1 with Buffer A R N A p . Ammonium sulfate (final concentration 0.165 g/ml) was then added and the solution centrifuged as described earlier. The volume of the aqueous phase was measured and the amount of ammonium sulfate needed to raise the concentration to 0.4 g/ml was prepared. After dilution of the aqueous phase with 1.5 volumes of Buffer A R N A P the solid ammonium sulfate was added to bring the solution to 0.4 mg/ml. After stirring, the protein suspension was centrifuged (39,000 x g, 15 min) and the supernatant removed. Pellets were dissolved in a minimal volume of Buffer B R N A P (10 m M Tris (pH 7.9), 1 mM EDTA, 500 mM sodium chloride, 10 mM magnesium chloride, 10% glycerol, 20 m M P-mercaptoethanol and 50 pg/ml PMSF), and proteins separated on a 2.7 cm x 110 cm Biogel column (BioRad) overnight. RNAP-containing fractions determined by assaying for activity as described earlier, were pooled and dialyzed twice for a total of 3 hr against 2 liters of Buffer C R N A P (10 m M HEPES (pH 7.9), 1 m M EDTA, 15% glycerol, 10 m M magnesium chloride, 20 mM P-mercaptoethanol and 50 pg/ml PMSF). Dialysates were loaded onto a heparin-agarose column (Sigma), washed until the absorbance at 280 nM fell below 0.1, and eluted with a 100-600 m M sodium chloride gradient in Buffer C R N A P - RNAP-containing fractions from the heparin-agarose column (determined by activity assay) were pooled and dialyzed against 2 liters of Buffer C R N A P for 2 hours and loaded onto a DNA-cellulose column. The DNA-cellulose column was washed with 50 mM sodium chloride in Buffer C R N A P until the absorbance at 280 nM fell below 0.1, and RNAP was eluted with 600 m M sodium chloride in Buffer C R N A P -RNAP-containing fractions determined by activity assay were pooled, concentrated in an Amicon Centriplus YM-50 ultrafiltration device (Millipore) as per the manufacturer's instructions. Glycerol was added to 30%, and the RNAP was aliquotted and stored at -70 °C. 45 3 Activation of SpoOA. SpoOA was phosphorylated by reconstituting the phosphorelay reaction in vitro (Bird et al., 1993). Reactions containing 8.0 pM SpoOA, 2.0 p M SpoOB, 2.0 p M SpoOF, 0.2 pM KinA and 1 m M ATP in 10 m M HEPES (pH 7.9) and 20 m M potassium acetate were composed and allowed to incubate at room temperature for 2 hours. KinA was the gift of Barbara Turner and SpoOB of J. A. Hoch (Scripps Research Institute). 40 pi aliquots of Spo0A~P were stored for up to two weeks at -20 °C. 4 In vitro Transcription. Standard reactions were composed of an 8 pi initiation mix containing 1 pi 1 Ox transcription buffer (100 mM HEPES (pH 8.0), 100 m M magnesium acetate, 1 m M DTT, 1 mg/ml acetylated bovine serum albumin and 200 mM potassium acetate), 2 pi 20 nM promoter DNA, 2 pi of 4 m M Spo0A~P, 400 nM ATP, 50 pM GTP and 3 pCi of [a 3 2P]GTP (800 Ci/mmol; PerkinElmer Life Sciences). The reaction tubes were incubated at the indicated temperature for 90 s before the addition of 1 pi containing RNAP. After the indicated initiation period, the complexes formed were challenged with 1 pi of elongation mix (100 pg/ml heparin, 400 nM UTP and 400 nM CTP) to allow run-off transcription. The reaction was stopped after 5 min with 5 pi of a saturated urea solution in 0.5x TBE (45 mM Trizma base, 45 mM boric acid and 1 mM EDTA), and the transcripts separated by electrophoresis over a 6% polyacrylamide gel (40 acylamide:lbis-acrylamide) containing 7 M urea in 0.5x TBE. The transcripts were detected by autoradiography using Kodak X A R film overnight at -20 °C, and promoter activity quantified on a Phosphorlmager SI (Molecular Dynamics; GE Healthcare) using ImageQuant 5.2 software. The percentage of template transcribed was calculated by dividing the moles of 131 base pair spoIIG transcript produced by the moles of template added to the reaction. The amount of transcript produced was calculated from Cerenkov radiation in an excised gel slice containing IT the transcript, the number of G residues per transcript, and the specific activity of the [a P]GTP in the reaction. 46 The apparent van't Hoff enthalpy reported represents the average value taken after substitution of the temperature values (7) and fractional relative transcription (F) at each non-zero data point into the formula AH =R\n((F]) - l)/((T)'1- (Tm)']) where the Tm or transition temperature is the temperature at which transcription is half-maximal (Grimes et al., 1991). 5 Potassium Permanganate Footprinting. Reactions containing 2 nM radiolabeled promoter fragments (1 x 105 cpm) in 20 pi of lx transcription buffer were composed; when present, the concentration of RNAP was 10 nM, SpoOA~P or SpoOAc was 800 nM and NTPs were 0.4 mM. Reactions were held at 37 °C for 2 min before the addition of 1 pi of freshly prepared 200 mM potassium permanganate. After 2 min further, the permanganate modification reaction was terminated with the addition of 50 pi stop solution (1.5 M P-mercaptoethanol, 420 m M sodium acetate, 0.1 m M EDTA and 30 ng/pl salmon sperm DNA). Samples were extracted once with phenol and the D N A precipitated by the addition of 3 volumes of 95% ethanol. Samples were held on crushed dry ice until solid, and the precipitate collected by bench-top centrifugation (14,000 x g, 30 min). Precipitates were dissolved in 100 pi 1 M piperidine and the samples were incubated for 30 min at 90 °C. After cooling the modified DNA, the D N A was precipitated by the addition of 10 volumes of butanol and collected by centrifugation (14,000 x g, 30 min). The butanol supernatant was removed, and the pellet dissolved in 150 pi of 1% SDS and butanol precipitated again. After collecting the pellet by centrifugation and removing the supernatant, the D N A was lyophilized for 30 min in a SpeedVac concentrator (Savant). Cleaved D N A fragments were dissolved in formamide loading buffer (10 mM EDTA (pH 8.0) and 1 mg/ml xylene cyanol in 80% formamide), denatured at 90 °C for 90 s, and separated by electrophoresis through a 5% polyacrylamide gel containing 7 M urea in 0.5x TBE. Equal numbers of counts were loaded in all lanes. Gels were dried and labeled D N A fragments were detected by autoradiography using Kodak X A R film overnight at -20 °C, and band intensity quantified on a Phosphorlmager SI (Molecular Dynamics; GE Healthcare) using ImageQuant 5.2 software. 47 6 DNase I Footprinting. Reactions containing radiolabeled promoter fragments (2 nM; 2.5 x 105 cpm) and 5 pi 1 Ox transcription buffer were composed in 45 pi aliquots. Samples were allowed to equilibrate for 2 min at 37 °C before 5 pi of diluted DNase I (5 x 10 ~3 units; Invitrogen) was added. Digestion was terminated after 15 s by the addition of 50 pi stop solution (40 m M EDTA (pH 8.0) and 300 m M sodium acetate) and 100 pi phenol. Samples were phenol extracted, and 1 pg of glycogen and 2.5 volumes of 95% ethanol added to the aqueous phase. Samples were held on crushed dry ice until solid, and the D N A was collected by centrifugation (14,000 x g, 30 min). The supernatant was removed and 150 pi of 70% ethanol added. Samples were held on crushed dry ice until solid, centrifuged as before and the supernatant removed. Samples were air-dried, re-suspended in 3 pi formamide loading buffer and separated by denaturing electrophoresis. Equal numbers of counts were loaded in all lanes. Gels were dried and labeled D N A fragments were detected by autoradiography using Kodak BioMax MS film (Kodak; GE Healthcare) overnight at -70 °C. 7 Electrophoretic Mobility Shift. Reactions containing 2.5 nM radiolabeled fragments (5 x 104 cpm) in 10 pi of lx transcription buffer were composed; the concentrations of RNAP, Spo0A~P or Spo0A c and iNTPs used are indicated in the figure legends. Samples were equilibrated at 37 °C for 2 min 20 s before the addition of 3.3 pi of either heparin loading buffer (0.1 pg/pl heparin, 20% glycerol and lx transcription buffer) or D N A loading buffer (0.3 pg/ul salmon sperm DNA, 20% glycerol and lx transcription buffer), and were immediately loaded onto running (20 mA) 4.5% non-denaturing polyacrylamide gels. Slab gels (145 mm x 90 mm x 1 mm) were prepared with 1 x Tris-Acetate (40 mM Trizma base and 40 m M acetic acid), using a 40 acylamide: 1 bis-acrylamide ratio and included 2% glycerol. Electrophoresis was continued for 2 hr for assays performed with D N A fragments greater than 100 base pairs, and 1 hr 15 min for assays performed with D N A fragments less than 100 base pairs. It was determined during the course of this work that it is critical to keep the electrophoresis time to a minimum to avoid dissociation of the complexes within the gel. Gels were dried and labeled D N A fragments detected by 48 autoradiography using Kodak X A R film overnight at -20 °C. Band intensity was quantified on a Phosphorlmager SI (Molecular Dynamics; GE Healthcare) using ImageQuant 5.2 software. Fractional binding was calculated as the ratio of Phosphorlmager units from bands representing specific complexes to the total Phosphorlmager units per lane. To estimate the relative stability of RNAP-promoter (RP) and RNAP-SpoOA~P-promoter (RPOA) complexes one master reaction was composed and incubated at 37 °C for 2 min. Immediately prior to the addition of unlabeled competitor DNA, a 9 pi aliquot from the master reaction was removed and loaded onto the running gel; this sample served to estimate the pre-challenge level of complex formation. After complexes were challenged with 25 nM unlabeled competitor, 10 pi aliquots were removed at the indicated times, added to 3.3 pi of D N A loading buffer and immediately loaded onto the running gel. Samples were prepared in which RNAP was added after the competitor allowing estimation of the levels of binding in the presence of competitor. The fraction of complex remaining was calculated by dividing the normalized fractional binding at the indicated times by the fractional binding immediately prior to the addition of unlabeled competitor. Normalized fractional binding was calculated by subtracting the levels of fractional binding in the presence of competitor from the fractional binding at the indicated times. Dissociation rates constants, kd, were determined from the first-order decay equation, 6 = 6>maxe~(Ad)(. 49 RESULTS 1 RNA Polymerase Purification. Modifications to the existing protocol for the purification of R N A P a A from B. subtilis were made to increase enzymatic activity. The preliminary purification steps, up to and including the first chromatography step that separated R N A P a A from smaller proteins or complexes on the basis of size, were identical to that described (Dobinson and Spiegelman, 1987). Biogel fractions (6 ml) were collected overnight at 0.4 ml/min (Fig. 12A). A small peak of protein elutes around fraction 36, while the concentration of protein steadily increases from fractions 48 to 70. RNAP activity was found between fractions 50 to 66. In lieu of concentrating polymerase activity on a D N A cellulose column followed by a preparative ultracentrifugation step over a 15-30% glycerol gradient, a heparin-agarose column was used to eliminate a persistent, contaminating nuclease (Fig. 12B). Two ml fractions were collected at 1 ml/min using a 100 to 600 mM sodium chloride gradient. Nuclease activity eluted within fractions 12 to 28, while RNAP eluted between fractions 30 to 42. RNAP activity was further purified by passage over a DNA-cellulose column at 1 ml/min collecting 2 ml fractions (Fig. 12C). RNAP activity co-eluted with the bulk of the protein; both A280 and RNAP activity peaked in fraction 35. Fractions 34 through 38 were pooled and the volume reduced from 10 to 2.5 ml. The concentrated RNAP was diluted with 1.1 ml of glycerol and stored in aliquots at -70 °C. The resulting preparation had an active RNAP concentration of 800 nM as judged in a single-round transcription assay with increasing concentrations of template DNA. 2 Characterization of Late Events in Transcription Initiation at the spoIIG Promoter. The focus of this work was to determine precisely how SpoOA stimulates transcription at the spoIIG promoter. Previous work had demonstrated that RNAP required activated SpoOA to form an open complex, and suggested that one avenue by which the role of SpoOA in D N A strand separation might be probed was through the use of a series of heteroduplex templates (Rowe-Magnus and Spiegelman, 1998b). Heteroduplex templates contain identical rather than complementary sequence at experimenter-defined base pairs, and have been previously identify the exploited to test the effects of D N A melting-deficient mutants in o A (Aiyar et al, 1994b), 50 40000 15000r 30000 20000 ' Activity ise Activity 10000 -10000 RNAI Nuclea 5000-60000 45000 g Fraction Fraction 1.00 0.75 I 0.501-w < 0.25 C. 0 . 0 0 l ^ i ^ » 29 70000 52500 -»-» 35000 \PAc 17500 0 40 Fraction Figure 12. RNAP purification. (A) Fractions from a Biogel size exclusion column were assayed for protein by A 2 8 0 (open squares) and for RNAP activity (closed circles). Absorbance values are on the left axis and RNAP activity on the right axis (cpm 3H-UTP incorporated into trichloroacetic acid precipitable material as described in Dobinson and Spiegelman, 1987). (B) Fractions eluted from a heparin-agarose column with a 100 to 600 nM sodium chloride gradient were assayed for nuclease activity (open circles) and RNAP activity (closed circles). Nuclease activity is on the left axis and RNAP activity on the right axis. Nuclease activity was followed by exposing 12,500 cpm of 32P-end labeled DNA to gradient fractions for 15 min and determining the loss of precipitable material. (C) Fractions eluted from a DNA cellulose column were assayed for protein by A 2 g 0 (open squares) and for RNAP activity (closed circles). 51 and to identify the transitions during transcription initiation upon which NtrC (Wedel and Kustu, 1995) and a-UP element interactions exert their stimulatory effects (Fredrick and Helmann, 1997). Here I took advantage of this system to investigate the effects of sequence (as single-stranded DNA) and size of the artificially denatured regions. By controlling these parameters I constructed templates that mimicked intermediates that may form during open complex formation, a relatively late event during transcription initiation. The rationale behind their construction follows. Two classes of heteroduplex templates were created: those with artificially denatured regions that contained non-template strand sequence on both strands, denoted NT, and those with artificially denatured regions that contained template strand sequence on both strands, denoted T. Since D N A in open complexes melts uni-directionally from the upstream edge (Chen and Helmann, 1997; Rowe-Magnus and Spiegelman, 1998b) and the upstream edge of the open complexes at spoIIG extends from -14, the heteroduplex template series shared a common upstream boundary at -14. And since the first stable open intermediate observed at spoIIG contains 12 base pairs of single-stranded D N A our heteroduplex template series included D N A fragments in which 2, 4, 6, 8 or 12 base pairs were artificially denatured. The heteroduplex templates were named according to the strand of origin of the sequence on both strands, the upstream edge of the artificially denatured region, and the downstream edge of the artificially denatured region. For example, NT14/13 contained the non-template strand sequence on both strands from -14 to -13 leading to a 2 base pair denatured region, while T14/3 contained the template strand sequence on both strands from -14 to -3 leading to a 12 base pair denatured region. These D N A constructs were used in an in vitro transcription assay to determine template utilization, SpoOA~P-dependence, temperature dependence, and relative rates of initiation-competent complex formation, and were compared to data obtained using a pair of fully double-stranded promoters, the wild type spoIIG promoter and a variant, IIG17. At the IIG17 promoter, the spacer separating the -35 and -10 elements has been reduced to a consensus 17 base pairs instead of the 22 base pairs found at the wild type promoter. A schematic of the various templates used for transcription assays in this study are presented in Fig. 13. Several transcripts were observed by denaturing polyacrylamide gel electrophoresis of the products from in vitro transcription reactions using some of these templates. In this thesis, 52 spoIIG IIG17 NT14/3 NT14/7 NT14/9 NT14/11 NT14/13 T14/3 T14/7 T14/11 • - < T Z = > • - < r = : > •-<r::;:::::> • - < J z : z : z z > • - < r r r : z : > D - C Z = > TATACTTATGAA TATACTTATGAA TATACTTA TATACTTA TATACT TATACT TATA TATA TA O TA ATATGAATACTT ATATGAATACTT ATATGAAT ATATGAAT ATAT ATAT Figure 13. In vitro transcription assay templates. Promoters used contain spoIIG DNA sequence from -100 to +134; only the region between -60 and +10 are schematized here. OA boxes are represented by the orange arrows, the -10 elements by the black outlined boxes and the -35 element by the dashed black outlined boxes. Transcription start sites are marked by arrows, and local regions of single-stranded DNA are indicated by the bubbles. Heteroduplex promoters contain either the non-template (NT) or template (7) sequence on both strands and are named to reflect the limits of the single-stranded regions. The sequence of the non-template strand (top) and the template strand (bottom) are shown beside the single-stranded regions. 53 only the major transcript, which comprised >70% of the total number of transcripts generated, is discussed. Previous primer extension analysis using R N A derived from NT14/7 and NT14/3 had determined that only the major transcript represents initiation at the position observed on the wild type promoter in the presence of SpoOA~P (Rowe-Magnus and Spiegelman, 1998b). 2.1 Levels of in vitro Transcription from Heteroduplex Templates. RNAP alone directed only a low level of transcription from a template created by annealing fully wild type spoIIG D N A strands (Table 6), whereas transcription was increased 6-50x for all the heteroduplex templates tested. Two general trends were evident. First, for promoter fragments with artificially denatured regions of equivalent size, RNAP was able to use templates containing non-template sequence as well or more effectively than templates containing template sequence. This is consistent with the importance of contacts between oA2.3 and the non-template strand sequence (Huang et al., 1997). Second, in general the amount of transcription from artificially denatured NT promoter fragments in the absence of SpoOA~P increased with the amount of single-stranded DNA. RNAP used 0.32 of the template with the largest denatured region, NT14/3 and only 0.04 of the template with the smallest denatured region, NT14/13. The exception, NT 14/7, also gave rise to a transcript that utilized the opposite strand of the heteroduplex (not shown), that would be predicted to obstruct RNAP binding and reduce the amount of the appropriate transcript generated. This correlation likely reflected improved formation of RNAP-promoter complexes on promoter fragments with larger artificially denatured regions, but the idea was not pursued further. 2.2 Effects of SpoOA~P on in vitro Transcription from Heteroduplex Templates. Because the influence of SpoOA on transcription is obscured by the variation in promoter strength, I calculated the fold-stimulation of transcription by SpoOA~P as the most meaningful comparison between heteroduplexes. SpoOA~P stimulated the level of transcription from the wild type spoIIG promoter 18x (Table 6). SpoOA~P stimulated transcription from templates with the two smallest pre-denatured regions, NT 14/13 and NT14/11, by 3.5x and 2.8x, respectively. By comparison, transcription from IIG17 and from the templates with the three largest pre-denatured regions, NT14/9, NT14/7 and NT14/3, was independent of SpoOA~P. 54 Table 6. In vitro Transcription: Template Utilization. PERCENT TEMPLATE TEMPLATE TRANSCRIBED 3 FOLD-STIMULATION BY SpoOA~P No SpoOA~P SpoOA~P spoIIG 0.6 ±0 .3 13 ± 2 18 ± 2 IIG17 13 ±2 12 ± 6 ~1 NT14/13 4 ± 1 16 ± 6 3.5 ±0 .7 NT14/11 5 ± 1 15 ± 6 2.8 ±0 .8 NT14/9 18 ± 4 18 ± 5 ~1 NT 14/7 11 ± 2 11 ± 4 ~1 NT 14/3 32 ± 8 30 ± 9 ~1 T14/11 7 ± 1 8 ± 1 ~1 T14/7 6 ± 1 6 ± 1 ~1 T14/3 6 ± 1 7 ± 1 ~1 aThe percentage of templates transcribed was calculated by dividing the moles of 131-bp spoIIG transcript produced by the moles of template added to the reaction. The transcript produced was calculated from Cerenkov radiation in an excised gel slice containing the transcript, the number 32 of G residues per transcript, and the specific activity of the [a P]-GTP in the reaction. The error represents the standard deviation from four experiments. 55 This sharp transition to SpoOA~P-dependence as the single-stranded region shrank from 6 to 4 base pairs (NT14/9 to NT14/11) indicated that the effect of SpoOA~P on open complex formation was limited to stages preceding the complete exposure of the non-template strand of the -10 element. Heteroduplexes containing the template strand sequence on both stands exhibited reduced levels of transcription and were not affected by the presence of SpoOA~P. This suggested that the single-stranded regions do not simply increase template flexibility that might help to align the -35 element and the transcriptional start site with the appropriate regions in RNAP. Moreover, the reduced levels of transcription are consistent with the interpretation that interactions between aA2.3 and single-stranded D N A from the non-template strand of the -10 element were essential for holoenzyme-directed transcription. The significance of the interaction between the non-template stand sequence of the -10 element and aAfrom B. subtilis (Qiu and Helmann, 1999) and a 7 0 of E. coli (Marr and Roberts, 1997; Roberts and Roberts, 1996) has been well-established during transcription initiation. Since transcripts from the T class of heteroduplex promoter fragments were SpoOA~P-independent and previous work has demonstrated that such artificially denatured regions were sufficient to permit initiation by RNAP core enzyme lacking a a subunit (Aiyar et al., 1994a), the T class of heteroduplexes were not further characterized. 2.3 Structural Probing of Open Complexes with Potassium Permanganate. The extent of D N A melting in response to RNAP, activated SpoOA, and combinations of ' nucleotides permitting the synthesis of R N A dimers, trimers and 11-mers was probed with potassium permanganate (KJVin04). KMn04 oxidizes pyrimidine residues at their 5,6 double bond to give pyrimidine glycol residues (Hayatsu and Ukita, 1967). Thymine is preferentially oxidized relative to cytidine, and the mechanism behind this preferential reactivity is believed to arise from an out-of-plane attack on the 5,6 double bond of the thymine ring. Although the ring remains intact, treatment of the pyrimidine glycol with a strong base leads to ring opening and cleavage of the phosphodiester backbone (Hayatsu and Ukita, 1967; Iida and Hayatsu, 1971; Rubin and Schmid, 1980). Since the 5,6 double bond of thymine is only accessible to KMn04 in the context of single-stranded or distorted DNA, KMn04 reactivity at thymine residues serves as a sensitive indicator of base pairing disruptions. The technique has been widely adopted to 56 follow D N A strand separation during open complex formation since its introduction (Sasse-Dwightand Gralla, 1989). I compared the position and extent of D N A strand separation on two heteroduplex templates to those formed at spoIIG under identical conditions. When present, SpoOAc was used instead of Spo0A~P as the isolated D N A binding domain is active in the absence of phosphorylation (Rowe-Magnus and Spiegelman, 1998a). This permitted the elimination of ATP (the initiating nucleotide at spoIIG) during the pre-initiation stage of the assay that might confound our assessment of D N A strand separation during early stages of transcription initiation. Consistent with previous reports (Rowe-Magnus and Spiegelman, 1998a), neither RNAP nor SpoOA c alone induced KMnCXt reactivity of thymines on a wild type template (Fig. 14A). When added together, SpoOAc and RNAP induced D N A strand separation between -13 and -3 as indicated by KMnCU reactivity of thymines on the template strand at -13, -11, -7, -4 and -3. Inclusion of ATP or pppApA permit the formation of an RNA dimer or occupation of the two nucleotide binding sites in the active site, but had no discernible effect on either the level of reactivity of individual thymines or the extent of DNA strand separation. With the inclusion of pppApA and GTP which allow the synthesis of an RNA trimer, the thymine at +2 became reactive and KMnC»4 reactivity at -13 was enhanced. In the presence of ATP and GTP, the denatured region extended to +13, though the thymines upstream of the transcription start site were virtually inaccessible. The lack of reactivity upon addition of iNTPs suggested that RNAP melted the promoter, initiated transcription, and translocated downstream more rapidly than KMnC>4 could oxidize upstream thymines. In all cases, KMnC^ reactivity was dependent on the simultaneous presence of RNAP and SpoOAc. Formation of an open complex at the spoIIG promoter required both RNAP and activated SpoOA. NT14/11 was chosen as representative of partially SpoOA-dependent heteroduplex promoter fragments. In the absence of any protein, thymines at -14 and -12 were accessible to KMnCU as a consequence of the 4 base pair pre-denatured region (Fig. 14B). In contrast to the effect of RNAP on spoIIG, pre-denaturing the promoter from -14 to -11 permitted RNAP to extend the denatured region to -3. The addition of SpoOAc enhanced the KMnCU reactivity of thymines at -12, -7, -4 and -3 suggesting that SpoOA c increased the number of open complexes formed but not the extent of the denatured region. As at spoIIG, inclusion of pppApA or ATP had 57 RNAP - + + + + + + + + + + SpoOAC - . + . + . + . + . + pppApA - - - + + ATP + + . . . . pppApA + GTP + + - -ATP + GTP + + spoIIG 1.+2 NT14/11 NT14/3 Figure 14. DNA strand separation at representative promoters. Transcription reactions containing 1 nM spoIIG (A), NT 14/11 (B) or NT 14/3 (C) radiolabeled on the template strand, 10 nM RNAP, 800 nM SpoOAC and the initiating nucleotides indicated above each lane were treated with potassium permanganate as described in the Experimental Procedures. Modified DNA was cleaved with piperidine and the cleavage products were analyzed on an 8% denaturing polyacrylamide gel. The gel was dried and exposed to film. Accessible thymines are indicated to the right side of each film. 58 negligible effects on KMnCv reactivity, while the addition of pppApA and GTP extended the denatured region past +1. Addition of ATP and GTP allowed RNAP to extend the denatured region to +13, accompanied by a striking reduction in KMnCU reactivity of thymines between -14 and -3. The near complete inaccessibility of thymines at -14 and -12 that cannot base pair suggests that they are buried in R N A polymerase. SpoOAc restored the KMn04 reactivity of thymines at -14 and -12 in the presence of ATP and GTP, though the reactivity of thymines at -7, -4 and -3 was reduced relative to the levels observed with other nucleotide combinations in the presence of SpoOAc . In all cases, SpoOAc enhanced KMnCU reactivity. The other heteroduplex template investigated was NT14/3, chosen as representative of SpoOA~P-independent heteroduplex promoter fragments. In the absence of any protein, thymines at -14, -12, -9, -8 and -6 were accessible to KMnC^ as a result of the 12 base pair pre-denatured region (Fig. 14C). Inclusion of RNAP had no effect on the size of the denatured region though it did change the relative reactivity of individual thymines; in particular, KMnC>4 reactivity was reduced at -14, enhanced at -12 and -6 and unchanged at -9 and -8. Though there was some variability between total counts within single lanes, the extent of the denatured region and the relative reactivity of individual thymines were not affected by the addition of pppApA or ATP. Unlike open complexes formed at spoIIG and NT14/11 however, thymines at +1 and +2 were not accessible to KMnC>4 under conditions that permit trimer formation. Inclusion of ATP and GTP permitted the extension of the denatured region to +13 without any concomitant reduction in the KMnCU reactivity of thymines upstream of+1. SpoOAc had no effect on the size of the denatured region or the relative reactivity of individual thymines. It is not clear why thymines at +1 and +2 were inaccessible in complexes formed on NT14/3 under conditions that permit trimer formation. 2.4 Temperature-Dependence of Transcription. As SpoOA~P is required to form an open complex at spoIIG (Rowe-Magnus and Spiegelman, 1998b) and because separation of the DNA strands is widely believed to be an energy intensive process (deHaseth and Helmann, 1995; Helmann and deHaseth, 1999), I reasoned that SpoOA~P might facilitate open complex formation by reducing an energetic barrier to transcription. It seemed possible then that lower reaction temperatures might enhance the dependence of 59 transcription on SpoOA~P even on templates with denatured regions. Denaturing additional base pairs could also require additional energetic input so that transcription from templates containing smaller pre-denatured regions might be more sensitive to reduced temperatures. Consequently, I investigated the effect of SpoOA~P on the temperature dependence of transcription from each of these templates. Without SpoOA~P, transcription from the fully duplexed wild type spoIIG promoter was low at all temperatures, although a slight increase in transcription with elevated temperature could be detected (Fig. 15A). Even in the presence of SpoOA~P, below 22 °C the level of transcription from the wild type spoIIG promoter was less than 0.04 of that observed at 37 °C. The amount of transcription increased dramatically between 27 and 37 °C, and the transition temperature (Tm), at which point transcription was half-maximal, was about 30 °C (Table 7). This value was higher than expected from studies with E. coli RNAP where the Tm is typically 25 °C under similar reaction conditions (Nakanishi et al., 1975). Extensive experience in the lab has shown that transcription from the spoIIG promoter in the presence of Spo0A~P is reduced at temperatures greater than 37 °C (data not shown), presumably due to the instability of RNAP. Therefore, transcription at temperatures above 37 °C for the other templates was not tested and the amount of transcription at 37 °C was taken as the maximal value. I had anticipated that reducing the overlong spacing at the spoIIG promoter would both obviate the requirement for SpoOA~P and, because of the appropriate phasing of the conserved promoter elements, be less energetically demanding. Although transcription from IIG17 was independent of SpoOA~P (Fig. 15B), the thermal profile of transcription from IIG17 was indistinguishable from spoIIG, both in terms of its response to temperature (the shape of the curve) and in its Tm (also 30 °C; Table 7). SpoOA~P had no effect on the temperature dependence of transcription from IIG17. Thus the unusual temperature-dependence of transcription of the spoIIG promoter was due to a characteristic of the interaction of RNAP with promoter sequences, and was not a consequence of the activity of SpoOA~P. In contrast to the low levels of transcription from the fully double-stranded promoters at reduced temperatures, appreciable amounts of transcription from NT14/3 were observed at temperatures as low as 12 °C (Fig. 15G). The amount of transcription increased gradually (about 0.2 every 60 Table 7. In vitro Transcription: Temperature-Dependence and Acquisition of Heparin-Resistance. TIME TO RATE TEMPLATE Tm AH3pp t x n a RP, M A X STIMULATION (°Q (kcalmoT1) (s) BY SpoOA~Pb spoIIG 30 42 60 25 ± 6 IIG17 30 42 60 1 NT14/13 28 33 30 3.5 ± 0.2 NT14/11 28 33 30 4.9 ±1 .6 NT14/9 24 24 <10 1 NT14/7 24 24 n.d.c n.d. NT14/3 20 13 <10 1 a The apparent van't Hoff enthalpy reported represents the average value taken after substitution of the temperature values (T in K) and fractional relative transcription (F) at each non-zero data point into the formula AH =R\n((Fi) - \)/((T)~'- (Tm)']) where the Tm or transition temperature is the temperature at which transcription is half-maximal (Grimes et al., 1991). The errors are estimated to be ± 30% of the cited value. bAverages and standard deviation from three experiments. cnot done. 61 5 10 15 20 25 30 35 40 Temperature (°C) 10 15 20 25 30 35 40 Temperature (°C) 10 15 20 25 30 35 40 Temperature (°C) 10 15 20 25 30 35 40 Temperature (°C) 10 15 20 25 30 35 40 Temperature (°C) 10 15 20 25 30 35 40 Temperature (°C) 10 15 20 25 30 35 40 Temperature (°C) Figure 15. Effect of SpoOA~P on the temperature dependence of transcription. Quantification of the primary products of in vitro transcription reactions. Reactions were performed using spoIIG (A), IIGI7(B), NT14/13 (C), NT14/11 (D), NT14/9 (E), NT14/7 (F) and NT 14/3 (G) as the template. The levels of transcription from the templates were plotted relative to the level of transcription in the presence of Spo0A~P at 37 °C. The reactions containing either 800 nM Spo0A~P (closed circles) or an equivalent buffer lacking SpoOA (open circles), 4 nM DNA template, and the initiating nucleotides ATP and GTP in lx transcription buffer were prepared on ice. The reactions were incubated at the indicated temperature for 2 min, and then transcription was initiated with the addition of lpl (400 fmol) RNAP. After 2 min, a single round of transcription was permitted by the addition of heparin, UTP and CTP. The transcription products were separated by electrophoresis, and the relative amounts of transcription were quantified as described in "Materials and Methods". The graphs represent the averages from four independent experiments. Errors were less than 10% of the plotted values and are omitted for clarity. At 42 °C, transcription from the spoIIG promoter in the presence of Spo0A~P was less than that at 37 °C (data not shown), presumably due to the instability of RNAP. Therefore, the amount of transcription at 37 °C was taken as the maximal value for transcription and the transition temperatures for each template listed in Table 7 are estimated based on the relative transcription at 37 °C. 62 5 °) between 12 and 22 °C before leveling off and increasing only about 0.1 every 5 0 between 27 0 and 37 °C. Notably, artificially denaturing the promoter between -14 and -3 reduced the Tm of transcription for this template to about 20 °C. Spo0A~P appeared to have no effect; the temperature dependence of transcription in the presence or absence of SpoOA~P appeared identical. The remaining templates fell into two classes on the basis of shared transition temperatures and Spo0A~P dependence. Although there were modest differences in the relative amounts of transcription at any given temperature, within experimental uncertainty, NT 14/9 and NT 14/7 exhibited a common Tm of 23-24 °C and were Spo0A~P independent (Fig. 15D and E). In comparison, the Tm values of NT14/13 and NT14/11 were significantly higher (28 °C), slightly less than that observed at fully double-stranded promoters (Fig. 15B and C). Critically, inclusion of SpoOA~P did not shift the Tm, indicating that the activator had no effect on the energetics of the overall transcription reaction from these templates. SpoOA~P appeared to increase the amount of transcript formed but not the thermal energy required to initiate transcription from these promoters. This indicates that Spo0A~P affects an energy-independent step in the transcription reaction; this contradicts any models in which Spo0A~P and RNAP cooperate to denature the D N A strands which required energy. 2.5 Temperature-Dependence of Open Complex Formation. In an attempt to gauge the amount of open complex formed as a function of temperature, KMn04 footprinting was performed on the spoIIG promoter in the presence of RNAP and SpoOA~P. Accessibility of thymine residues to K M n 0 4 was probed at temperatures between 7 and 37 °C (Fig. 16). Under our experimental conditions, no difference in KMn04 activity was observed with reduced temperatures (not shown). As previously described, incubation of RNAP, Spo0A~P and spoIIG resulted in the exposure of template strand thymines at -13, -11, -7, -4 and -3 at 37 °C (Fig. 16). Reducing the temperature to 32 °C decreased the accessibility of thymines (sum of all reactive bands) to 0.74 of that observed at 37 °C. Ternary complex formation at 27 °C reduced thymine reactivity to 0.14 of the maximum level, while the fraction of complexes containing single-stranded D N A at temperatures less than 22 °C was negligible. The Tm of open complex formation was 30 °C, identical to the Tm of transcription, suggesting 63 A. Temperature (°C) Figure 16. Effect of temperature on open complex formation. (A) Reactions containing 1 nM spoIIG radiolabeled on the template strand, 10 nM RNAP and 800 nM SpoOAC (when present) were composed and incubated at the indicated temperatures. Complexes were probed with potassium permanganate as described in the Experimental Procedures. Modified DNA was cleaved with piperidine and separated on an 8% denaturing polyacrylamide gel. The gel was dried and exposed to film overnight. (B) A representative gel was exposed to a phosphor screen for 2 hours and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. The fraction of open complex was calculated by dividing the total number of units measured by the phosphorimager between -13 and -3 at each temperature by the total number of units measured by the phosphorimager between -13 and -3 at 37 °C. 64 that D N A strand separation (or some combination of steps preceding it) was the energetic barrier to transcription at spoIIG. 2.6 Relative Rates of Acquisition of Heparin-Resistance. Heparin resistance has been used to discriminate between stable and unstable RNAP-promoter complexes. At the spoIIG promoter, heparin resistance requires the synthesis of an pppApApGRNA trimer and thus represents a relatively late stage in the initiation reaction (Bird et al, 1996). In practice, inclusion of SpoOA~P, ATP, GTP and the spoIIG promoter results in the formation of an 11 -mer RNA. As a means of investigating the effect of denatured base pairs on the formation of initiated complexes at the spoIIG promoter, I examined the rates of acquisition of heparin-resistant complexes on the series of pre-denatured templates. In the absence of SpoOA~P, the formation of heparin-resistant initiated complexes at spoIIG was negligible regardless of the incubation time (Fig. 17A). When SpoOA~P was present the formation of initiated complexes increased steadily for the first 15 s. By 30 s, 0.6 of the maximal number of initiated complexes had formed, and the proportion of resistant complexes increased to just less than 0.9 of the maximal level within the first 120 s. The formation of initiated complexes at the IIG17 promoter occurred with an initial rate 1.5-fold faster than that at the wild type spoIIG in the presence of Spo0A~P (Fig. 17B; Table7). Although inclusion of Spo0A~P resulted in a small increase in the amount of initiated complexes formed, this effect was within experimental uncertainty. The heteroduplex templates tested fell into two distinct categories. Initiated complexes formed extremely rapidly on NT 14/9 (Fig. 17E) and NT14/3 (Fig. 17F). Maximal levels of initiated complex were formed in less than 10 s in the presence or absence of Spo0A~P. By comparison, heparin-resistant complexes formed more slowly on NT14/13 (Fig. 17C) and NT14/11 (Fig. 17D) than on templates with larger denatured regions but more rapidly than on the fully double-stranded promoters; maximal levels of initiated complex formation took about 30 s on NT14/13 and NT14/11. The effects of Spo0A~P at NT14/13 and NT14/11 were two-fold. First, although RNAP alone was able to isomerize to a heparin-resistant form on NT14/13 and NT14/11, Spo0A~P stimulated the initial rate of this transition about 4-fold in each case. Second, 65 0 20 40 60 80 100 120 0 20 40 60 80 100 120 0 20 40 60 80 100 120 Time (s) &•—•—•—•—i—i—i u 6—i—i—i—i——i—i u 4 i i i i i I 0 20 40 60 80 100 120 0 20 40 60 80 100120 0 20 40 60 80 100120 Time (s) Figure 17. Time course of initiated complex formation. Quantification of the primary products of in vitro transcription reactions. Reactions were performed using spoIIG (A), IIG17 (B), NT14/13 (C), NT14/11 (D), NT14/9 (E), NT14/3 (F) as the template. The transcription levels indicative of the amount of initiated complex formed at various times was plotted relative to the transcription levels in the presence of Spo0A~P after 120 s for each template. The transcription level indicative of the amount of initiated complex formed at at various times at the other templates was plotted relative to the transcription levels in the absence of Spo0A~P after 120 s. The reactions containing either 800 nM Spo0A~P (closed circles) or an equivalent buffer lacking SpoOA (open circles), 4 nM DNA template, and the initiating nucleotides ATP and GTP in lx transcription buffer were prepared on ice. The reactions were incubated at 37°C for 2 min, and then transcription was initiated with the addition of lpl (400 fmol) RNAP. At the indicated times, samples were withdrawn and added to a lpl mixture containing heparin, UTP and CTP. Elongation was allowed to proceed for 5 min before the reactions were terminated, and the transcription products were separated by electrophoresis. Relative amounts of transcription were quantified as described in "Experimental Procedures". The graphs represent the averages from three independent experiments. Errors were less than 8% of the plotted values and are omitted for clarity. The time required to acheive maximal formation of initiated complexes and the estimated effect of Spo0A~P in stimulating the rate of this process is listed in Table 6. 66 SpoOA~P increased the total number of complexes that acquired heparin-resistance on the heteroduplex templates containing the smaller denatured regions. 3 Characterization of Early Events in Transcription Initiation at the spoIIG Promoter. Earlier work has shown that the RNAP-spoIIG promoter complex cannot efficiently initiate transcription on a linear D N A template (Bird et al., 1993; Bird et al., 1996). SpoOA~P is required, in part by acting in concert with RNAP, to permit formation of an open complex at this promoter (Rowe-Magnus and Spiegelman, 1998b). However, the results of the previous section reveal that the activator does not reduce the energetic barrier to D N A melting (Seredick and Spiegelman, 2004). Since RNAP binds to the spoIIG promoter relatively efficiently (Bird et al, 1996), this suggested that SpoOA~P and RNAP must cooperate to form an intermediate complex preceding formation of an open complex. I sought evidence for this intermediate and characterized some of the factors leading to its assembly. 3.1 Evidence for Low Temperature Intermediates at the spoIIG Promoter. Protein-DNA complex formation was assayed using electrophoretic mobility shift assays (EMSAs) performed at 37°C, a temperature known to be permissive for open complex formation (Fig. 16). Consistent with earlier reports (Bird et al, 1996; Rowe-Magnus and Spiegelman, 1998a; Satola et al, 1991), at 37 °C SpoOA~P does not bind well to the spoIIG promoter while RNAP formed two complexes as judged by differences in electrophoretic mobility (Fig. 18A). These complexes differed in the extent to which upstream D N A wraps around RNAP (G.B.S., unpublished observation). Inclusion of SpoOA~P and RNAP increased the amount of D N A fragment bound by RNAP, and resulted in a complex with lower electrophoretic mobility (supershifted the complexes). A l l complexes formed survived challenge with an excess of non-specific D N A ; however only initiated complexes (those formed in the presence of iNTPs) were resistant to heparin challenge (Fig. 18B). As mentioned earlier, heparin-resistant complexes at the spoIIG promoter require the formation of at least a pppApApG RNA trimer. Previous analyses of RNAP-spoIIG promoter complex formation by E M S A have been limited because low levels of complexes were formed (for example, see Cervin et al., 1998b). Using the modified RNAP purification protocol and adjusting the electrophoretic conditions (see 67 spoIIG -C=K=> C=>-C=> RNAP - - + + + + SpoOA-P . + . + _ + ATP & GTP - . . _ + + Figure 18. SpoOA~P and RNAP form a low temperature intermediate complex at the spoIIG promoter. A schematic of the DNA template used is shown at the top of the figure. The fragment contains the spoIIG gene sequence from -100 to +134 and contains four OA boxes (orange arrows), a -35 element that overlaps with the promoter-proximal OA box (dashed black box), a -10 element (black box) and the transcription start site (bent arrow). The same symbols are used in the following figures. Reactions containing radiolabeled Hindlll-BamHI fragments of the spoIIG promoter (shown at the top) and the indicated components were incubated at 37 or 7 °C. After three minutes, loading buffer containing heparin or DNA was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 2 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film. When present, [SpoOA~P] was 800 nM, [RNAP] was 40 nM and [iNTPs] were 0.4 nM. Arrows to the right of the panels indicate KNAP-spoIIG (RP) and KNAP-spoIIG-Spo0A~P (RPOA) complexes. 68 Experimental Procedures, section 7), this limitation could be overcome (for example, see Fig. 18A). To identify potential closed intermediates, identical assays were performed at 7 °C, a temperature that does not permit D N A melting to occur (Fig. 16). The pattern of complexes formed at 7 °C and stable to D N A challenge were qualitatively similar to those formed at 37 °C. Again, SpoOA~P binding to the spoIIG promoter fragment was not significant while RNAP formed two complexes (Fig. 18C). The complex formed in the presence of RNAP and SpoOA~P was supershifted relative to that formed by RNAP alone, and the addition of RNAP and SpoOA~P resulted in stable binding of nearly all of the labeled fragment. As expected, at 7 °C none of the complexes survived heparin challenge, and were therefore uninitiated complexes (Fig 14D). These results supported the idea RNAP and SpoOA~P formed a closed intermediate at the spoIIG promoter at 7 °C. 3.2 Evidence for a Closed RNAP-s/w/ZG'-SpoOA-P Intermediate. To ensure that the complex observed at 7 °C was an intermediate during transcription initiation and not an off-pathway, low temperature artifact, I assayed the ability of complexes, pre-formed at 7 °C, to initiate transcription when shifted to 37 °C (Fig. 19). RPOA complexes were formed at 7 or 37 °C, and iNTPs plus excess unlabeled spoIIG promoter fragments were added to permit initiation and to prevent re-association of RNAP that has released the labeled spoIIG promoter fragments, respectively. Samples were returned to either 7 or 37 °C and were challenged with heparin. Complexes were analyzed by E M S A . Only those complexes that remain bound to radiolabeled templates and initiated transcription survive heparin challenge and are visible. At 37 °C, RPOA complexes capable of initiating transcription (as judged by the acquisition of heparin-resistance) were formed on 0.51 of the labeled template in the presence of iNTPs and a 10-fold excess of unlabeled spoIIG competitor (Fig. 19, lane 2). Inclusion of unlabeled competitor prior to the addition of RNAP reduced the fraction of labeled template utilized by initiated complexes to 0.11 (Fig. 19, lane 3). This reduction demonstrated the competitor's effectiveness in sequestering RNAP. At 7 °C, less than 0.02 of the labeled template was found in initiated complexes regardless of when the competitor was added to the reaction (Fig. 19, lanes 4 and 5). When initiating nucleotides and a 10-fold excess of competitor were added to ternary complexes formed at 7 °C, increasing the temperature to 37 °C allowed initiated 69 spoIIG RNAP . + + + + + + SpoOA~P + + + + + + + competitor - - + - + . + binding T (°C) 37 37 37 7 7 7 7 I ATP + GTP or competitor + ATP + GTP I initiation T (°C) 37 37 37 7 7 37 37 Figure 19. RPOA complexes formed at low temperatures can initiate transcription if brought to 37 °C. Reactions were composed with 4 nM radiolabeled spoIIG promoter DNA, 10 nM RNAP and 800 nM SpoOA~P and allowed to equilibrate at the indicated binding temperature. After 2 min, ATP and GTP or ATP, GTP and 40 nM unlabeled spoIIG promoter DNA were added. The reaction tubes were then returned to baths at the indicated initiation temperatures. After 2.5 min, loading buffer containing heparin was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 2 hr at 20 mA to separate initiated complexes from free DNA. Gels were dried and exposed to film. 70 complex formation on 0.35 of the labeled template (Fig. 19, lane 6). By comparison, inclusion of unlabeled competitor reduced the utilization of labeled template by initiated complexes to 0.08 (Fig. 19, lane 7). Thus, at 7 °C RNAP initiates transcription very inefficiently, but 0.35/0.51 or 0.69 of closed RPOA complexes formed at 7 °C initiated transcription when brought to temperatures that permit D N A strand separation. This result is consistent with the idea that a closed RPOA complex at 7 °C is an authentic intermediate during transcription initiation at the spoIIG promoter. 3.3 Complex Formation on a Truncated Derivative of the spoIIG Promoter. The improvement in ability to detect RPOA and RP complexes by E M S A (previous section) provided the means to explore RNAP-Spo0A~P interactions at stages preceding open complex formation for the first time. Coupled with the finding in Results, section 2.4, that Spo0A~P stimulated transcription without affecting any process energetically linked to open complex formation, the E M S A results indicated that the critical interaction between RNAP and Spo0A~P involved steps at the binding of the two proteins to the promoter. This binding is paradoxical since one of the two SpoOA binding site overlaps the -35 site, and i f Spo0A~P binds as a dimer, the dimer should compete for that region with RNAP. One interaction model would be that Spo0A~P displaces RNAP, while potentially holding RNAP near the D N A while the polymerase searches for downstream sequences including the -10 element. Since it was possible to observe these RPOA complexes by electrophoretic mobility shift assays, I attempted to separate the interaction of Spo0A~P and RNAP with upstream promoter sequences from the interaction of RNAP with downstream promoter sequences. As a first step, the interaction of Spo0A~P and RNAP was examined using a truncated spoIIG promoter fragment that contained the D N A sequences from -100 to -15, TIIG. The lack of sequences downstream of-15 would eliminate contributions from the interaction of RNAP with the -10 element and the transition to open complex formation that occurs largely independently of Spo0A~P (Results, section 2.1-2.5). It seemed likely that RNAP would not require downstream sequence elements for initial binding as the -35 and -10 elements at the spoIIG promoter are separated by an additional one-half helical turn. Consequently, they are presented out of phase with one another, and so are probably contacted separately. 71 spoIIG TIIG RNAP SpoOA~P SpoOAc RPOA RPOA, Figure 20. RNAP and SpoOA~P form a complex on a truncated promoter derivative, TIIG. The TIIG promoter fragment contains the spoIIG promoter sequence from -100 to -15 and lacks the -10 element and all DNA downstream. Reactions were composed with 2.5 nM radiolabeled spoIIG promoter DNA, 10 nM RNAP and 800 nM Spo0A~P or SpoOAC, and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film; a single representative film is shown. 72 The complexes formed by RNAP and SpoOA~P on TIIG (sequence from -100 to -15) were qualitatively similar to those formed on the full-length fragment (-100 to +134) as measured by E M S A (Fig. 20). Neither Spo0A~P nor the isolated D N A binding domain of SpoOA, Spo0A c, were able to bind to the fragment (lanes 2 and 3). RNAP bound efficiently to TIIG in the absence of SpoOA, forming two complexes that differed in electrophoretic mobility, as with the full-length fragment (lane 4). This confirms the idea that RNAP can make initial contacts with upstream D N A sequences at this promoter in the absence of the -10 region. The inclusion of Spo0A~P increased the fraction of bound D N A and supershifted the complexes (lane 5). This is the first direct evidence that Spo0A~P enhances RNAP-promoter interactions, and that Spo0A~P does not work by stimulating RNAP to exchange contacts around the -35 element for contacts around the -10 element. The replacement of Spo0A~P with SpoOAc resulted in a supershifted complex of slightly greater mobility than that formed by RNAP and full-length Spo0A~P (compare lanes 5 and 6). This is consistent with the interpretation that SpoOA is part of the intermediate closed complex, and does not simply modify RP. 3.4 DNase I Footprinting of the spoIIG Promoter and the TIIG Promoter Fragment. DNase I footprinting pattern of complexes formed on spoIIG (Fig. 21) and TIIG (Fig. 22) promoter fragments were analyzed to compare complex formation on the truncated fragment with that on the full-length promoter. At spoIIG, Spo0A~P alone protected regions from DNase I cleavage located from -105 to -80 and from -55 to -35 that matched the positions of the OA boxes. This is consistent with previous reports (Rowe-Magnus and Spiegelman, 1998a; Satola et al, 1991; Satola et al, 1992). The addition of RNAP resulted in protection of a reactive site near -90 and complete protection between -60 and -10 except for reactive sites at -54, -45 and -23. Partial protection of reactive sites was observed between -10 to +4. By comparison, E. coli RNAP protects lacUV5 promoter D N A from -55 to +15 with hypersensitive sites centered around -37 and -23 (Kovacic, 1987), and protects the XPR promoter from -57 to +20 with reactive sites at -48 and -38 (Craig et al, 1995). The simultaneous presence of Spo0A~P and RNAP gave additive zones of protection from -100 downstream to +8 with reductions in DNase I cleavage that could be clearly attributed to either Spo0A~P or RNAP on the basis of their individual footprints. Specifically, reductions in DNase I cleavage at -100, -96, -83, -82 -61, -54 and -45 in RPOA relative to RP could be attributed to Spo0A~P, and compared to P0A, 73 A. RNAP - - + + + B. SpoOA~P - + . + + ApA + GTP - - - - + P POA RP RPOA RPOAj Figure 21. SpoOA~P re-models RNAP protection of the spoIIG promoter. (A) A DNA fragment containing the spoIIG promoter sequence between -100 and +50 radiolabeled on the template strand was incubated with combinations of 800 nM Spo0A~P, 80 nM RNAP and 0.4 mM ApA and GTP as indicated. After 2 min, complexes were probed with DNase I for 15 s before digestion was halted by the addition of an equal volume of 40 mM EDTA and 300 mM NaAc and two volumes of phenol. Samples were prepared as described in Experimental Procedures and the DNA separated on an 8% denaturing polyacrylamide gel. The gel was dried and exposed to film overnight. (B) A schematic of the spoIIG promoter fragment, P, the regions protected by Spo0A~P, POA, in orange; the regions protected by RNAP, RP, in blue; the regions protected by RNAP and Spo0A~P, RPOA, in light green; and the regions protected by initiated complexes (RNAP, Spo0A~P and ApA and GTP), RPOAj in dark green, are aligned with the scanned film. 74 A . RNAP SpoOA~P -100 + + B. P POA RP RPOA -50 OA box OA box 1.2 OA box 2.1 OA box 2.2/ -35 element Figure 22. SpoOA~P and RNAP form a specific complex on a TIIG promoter fragment. (A) A DNA fragment containing the spoIIG promoter sequence between -100 and -15 radiolabeled on the template strand was incubated with combinations of 800 nM Spo0A~P and 80 nM RNAP as indicated. After 2 min, complexes were probed with DNase I for 15 s before digestion was halted by the addition of an equal volume of 40 mM EDTA and 300 mM NaAc and two volumes of phenol. Samples were prepared as described in Experimental Procedures and the DNA separated on an 10% denaturing polyacrylamide gel. The gel was dried and exposed to film overnight. (B) A schematic of the TIIG promoter fragment, P, the regions protected by Spo0A~P, POA, in orange; the regions protected by RNAP, RP, in blue; the regions protected by RNAP and Spo0A~P, RPOA, in green, are aligned with the scanned film. 75 reductions in DNase I cleavage at -61, -20, -15, -13, -9, -7, -5, -4, +5, +7 and +8 in RPOA, could be attributed to RNAP. In addition, protection between -10 and +8 was enhanced in the presence of SpoOA~P. The individual footprints of either Spo0A~P or RNAP could not account for protection from cleavage at -23 by RPOA on spoIIG; the base was accessible to DNase I in presence of either protein alone, but not when a ternary complex was formed suggesting that SpoOA~P modified RP complexes. The DNase I footprint of initiated complexes (RNAP, SpoOA~P, pppApA and GTP) was indistinguishable to that of uninitiated complexes. SpoOA~P protected the same two regions of the TUG fragments as it protected on the fragment containing the full-length spoIIG promoter. RNAP protected the TUG promoter fragment from -62 to the 3' end of the fragment except for a reactive site at -54 and a hypersensitive site at -45. RNAP also protected an isolated site at -91. Together, RNAP and SpoOA~P protected the promoter from -108 to the 3' end of the fragment, except for a single reactive site at -47 that was present in the lanes including Spo0A~P, but not in the RNAP-77/G complex. At spoIIG, the same pattern of protection at -47 was observed but was not as prominent in part because SpoOA~P partially protected the site. In spite of these slight variations in the cleavage patterns between spoIIG and 77/G, Spo0A~P and RNAP appear to bind to identical positions on the two fragments. 3.5 Characterization of Complex Assembly on TUG. 3.5.1 RNAP Titration. The observation that the RPOA complexes formed on the full-length spoIIG promoter and the truncated TUG promoter fragment provided a tool to characterize assembly of this closed complex. Complex formation as a function of RNAP concentration was compared in the presence or absence of SpoOA~P (Fig. 23). RNAP alone bound efficiently, linearly up to 10 nM with fractional binding saturating at 0.45 at 40 nM RNAP. In the presence of Spo0A~P, complex formation never saturated reaching a maximum value of 0.69 at 80 nM RNAP. At all inputs, Spo0A~P increased the amount of D N A bound. Critically, Spo0A~P did not reduce the concentration at which RNAP binding saturated. This finding implied that Spo0A~P did not recruit RNAP to TUG. 76 TUG SpoOA~P + + + + + + + 0 10 20 30 40 50 60 70 80 [RNAP] (nM) Figure 23. Complex assembly on TUG- RNAP titration. Binding reactions containing 2.5 nM radiolabeled 77/G DNA, 0 to 80 nM RNAP and 800 nM Spo0A~P were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film overnight or phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. Fractional binding was calculated by dividing the Phosphorlmager units representing specific complexes by the total Phosphorlmager units per lane. Representative data from one of three experiments is plotted; RP (open circles), RPOA (closed circles). 77 A. TIIG — C = K = Z 3 C=3—0=3 RNAP + + + + + + [SpoOA~P] (nM) [SpoOA~P] (nM) Figure 24. Complex assembly on TIIG - SpoOA~P titration. (A) Binding reactions containing 2.5 nM radiolabeled TIIG DNA, 0 to 800 nM SpoOA~P and 40 nM RNAP where indicated were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film overnight or phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. (B) Spo0A~P titration at several RNAP concentrations. Fractional binding was calculated by dividing the Phosphorlmager units representing shifted complexes (no RNAP) or supershifted complexes (RNAP present) by the total Phosphorlmager units per lane. Representative data from a single set of titrations at 0 nM RNAP (open circles), 10 nM RNAP (hatched circles), 20 nM RNAP (grey circles) and 40 nM RNAP (closed circles) are shown. (C) Complex formation at 20 nM RNAP. Representative data from one of three experiments is shown; POA (closed squares), RP (open circles), RPOA (closed circles). 78 3.5.2 SpoOA~P Titration. Since SpoOA~P-containing complexes could be discriminated from those that lack SpoOA~P, it was possible to follow formation of supershifted RPOA complexes as a function of SpoOA~P concentration at several concentrations of RNAP (Fig. 24B). In the absence of RNAP, even 800 nM SpoOA~P (a 320-fold molar excess over the labeled D N A probe) shifted less than 0.04 of the labeled fragment. Increasing amounts of RNAP led to an increased fraction of supershifted complex at each concentration of SpoOA~P. That is, more RNAP allowed the incorporation of more Spo0A~P into supershifted RPOA complexes. At 10 and 20 nM RNAP, increasing amounts of Spo0A~P led to the formation of increasing amounts of supershifted complex, while Spo0A~P binding saturated in the presence of 40 nM RNAP at about 600 nM Spo0A~P. In the presence of 40 nM RNAP, only the supershifted complex was observed at 600 and 800 nM Spo0A~P, and half-maximal supershifted complex formation occurred at 200 nM Spo0A~P. These results suggest that RNAP recruits Spo0A~P to form the supershifted complex on this truncated fragment. 3.6 Definition of the Minimal Promoter Fragment Necessary for Complex Formation. 3.6.1 Downstream Truncations. The observation that Spo0A~P did not displace RNAP from TUG prompted an investigation of the 3' limit of D N A essential for the formation of RP and RPOA. EMS As were performed using a nested set of promoter fragments that extended from -100 downstream with 3' termini between -35 to -19 (Fig. 25A; the template with a 3' end at -15 corresponds to TUG). RP and RPOA complexes were able to form on each of the templates tested (Fig. 25B). At each promoter fragment in this series Spo0A~P enhanced RNAP binding. Quantification of the fraction of labeled template incorporated into RNAP-containing complexes indicated no systematic trends in the efficiency of RP or RPOA complex formation with diminishing downstream D N A (Fig. 25C). This data suggested that contacts to D N A downstream of -35 played a minimal role in the assembly of RP or RPOA complexes on promoter fragments lacking -10 elements. Conversely, these results indicate that the major contributions to RP and RPOA formation are made by sequence elements lying between -100 and -35. 79 3' terminus TUG — C = K = 1 C=3—0=3 - ! 5 77/GX-19) —<=K=zn :>-<rZZ3 -19 77/6X-23) — ( = K = D C=Z>^ZZ3 -23 77/G(-27) — C = K = 3 C = K i = 3 -27 77/G(-35)—C= — 1ZZ> -35 B. RNAP + + + + + + + + + + SpoOA~P + - + - + - + - + C. 0.8 :g 0.6 g s •a 0.4 & & & 0.2 0.0 - iti n ri, I H I II l l l l l l l l rh ^ r S > r 6 > ^ ^ <6 Figure 25. Complex formation on 3' nested spoIIG promoter fragments. (A) Promoter fragments used in this assay contained spoIIG promoter sequence from -100 to the 3' terminus indicated. (B) Binding reactions containing 2.5 nM radiolabeled DNA were incubated with 20 nM RNAP and when present, 800 nM Spo0A~P were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film overnight or phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. (C) Fractional binding was calculated by dividing the Phosphorlmager units incorporated within shifted complexes by the total Phosphorlmager units per lane. Data shown represent the average and standard deviation from three independent experiments; RP complexes (grey), RPOA complexes (white). 8 0 A. 5' terminus TIIG -15 —<=_>£___ (-80)77/G -80 ( 1-(-69)77/G -69 ^ = D _ H j I Z > (-58)r//G -58 —C=3—c;::.> Figure 26. Complex formation on 5' nested spoIIG promoter fragments. (A) Promoter fragments used in this assay contained spoIIG promoter sequence from the 5' terminus indicated to -15. (B) Binding reactions containing 2.5 nM radiolabeled DNA, 20 nM RNAP and when present 800 nM Spo0A~P, were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film overnight or phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. (C) Fractional binding was calculated by dividing the Phosphorlmager units representing shifted complexes by the total Phosphorlmager units per lane. Data shown represent the average and standard deviation from three independent experiments; RP complexes (grey), RPOA complexes (white). 81 3.6.2 Upstream Truncations. The results of the previous section raise the question of how RNAP binds so efficiently in the presence of SpoOA~P, under conditions that would appear to prevent any contacts between c A and the promoter fragment. EMSAs were performed using a nested set of promoter fragments with 5' termini at -100 (TUG), -80, -69 and -58 extending to a common downstream end at -15 (Fig. 26A). At 20 nM RNAP, RP complexes form on 0.46 of labeled TUG templates. Addition of 800 nM Spo0A~P supershifted the RNAP-containing complexes and increased the fractional binding of promoter fragment to 0.72 (Fig. 26B and C). On the -80 to -15 promoter fragment RP formed very efficiently, incorporating 0.66 of the labeled template at 20 nM RNAP. While Spo0A~P clearly bound to the RP complex as evidenced by the reduction in electrophoretic mobility, it did not significantly strengthen the binding of RNAP to the fragment. In comparison, RP formation on the -69 to -15 fragment was reduced to a fractional binding of 0.26 (1.8-2.5x less relative to the two longer fragments), and no RPOA was formed (indicated by the absence of a supershifted complex). RP formation on the -58 to -15 fragment was further reduced to a fractional binding of 0.07 to the labeled D N A (6.6-1 lx less relative to the two longest fragments) and RPOA complexes did not form. These results are consistent with the interpretation that RNAP makes contact with the D N A between -80 and -58, and that the region between -69 and -58 is a critical portion of the promoter for the assembly of RP complexes. The data also indicated that Spo0A~P binding to the promoter via recruitment by RNAP (the formation of RPOA complexes) minimally required the D N A between -80 and -69 despite the fact that Spo0A~P clearly does not bind to that region by itself (Fig. 21 and 22). 3.7 Characterization of Complex Assembly on (-80)77/(7. 3.7.1 RNAP Titration. The observation that deletion of the promoter from -100 to -80 (that removes the upstream OA boxes) improved RNAP binding relative to complex formation on TUG prompted a more thorough investigation of complex assembly on this fragment. Complex formation on (-80)77/(7 promoter fragments as a function of RNAP concentration was compared in the presence or absence of Spo0A~P (Fig. 27). RNAP alone bound efficiently, linearly up to 20 nM with fractional binding saturating at a fractional binding of 0.75 at 40 nM RNAP. In the presence of 8 2 (-80)77/G RNAP SpoOA~P RPOA RP 0 10 20 30 40 50 60 70 80 [RNAP] (nM) Figure 27. Complex assembly on (-80) TZ/G - RNAP titration. Binding reactions containing 2.5 nM radiolabeled (-80)77/G DNA, 0 to 80 nM RNAP and 800 nM Spo0A~P were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film or phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. Fractional binding was calculated by dividing the Phosphorlmager units representing specific complexes by the total Phosphorlmager units per lane. Representative data from one experiment of three is plotted; RP (open circles), RPOA (closed circles). 83 SpoOA~P, RPOA formation also saturated having reached a maximum fractional binding of 0.78 at 40 nM RNAP. As with the TIIG D N A fragment, the presence of SpoOA~P did not reduce the concentration at which RNAP binding saturated and consequently did not recruit RNAP to the promoter fragment. However, relative to RP and RPOA formation on the TIIG D N A fragment, there were two significant differences. First, Spo0A~P did not increase the amount of D N A incorporated into RPOA complexes relative to RP complexes at identical RNAP inputs on this promoter fragment. This was a consequence of increased RP complex formation on the (-80)TIIG promoter fragment relative to the TIIG promoter fragment (compare Fig. 27 to Fig. 23), and suggested that the D N A between -100 and -80 inhibits RP formation. Secondly, using the (-80)TIIG promoter fragment in the presence of 800 nM Spo0A~P, some RP complex remained at all RNAP concentrations (Fig. 27). In comparison, 800 nM Spo0A~P was sufficient to convert all RP complexes formed on the TIIG promoter fragment to RPOA complexes (Fig. 24). 3.7.2 Spo0A~P Titration. One potential hypothesis to explain the residual persistence of RP complexes in the presence of Spo0A~P on the (-80)TIIG fragment is that the upstream OA boxes removed act to increase the concentration of Spo0A~P in the immediate vicinity of the promoter. This could facilitate the occupation of the lower affinity OA boxes downstream. Such a proposal is consistent with the known affinities of Spo0A~P for the two pairs of binding sites; site 1 OA boxes are occupied before site 2 boxes (Rowe-Magnus and Spiegelman, 1998a; Satola et al., 1991; Satola et al., 1992). This hypothesis predicts that more Spo0A~P would be required to form RPOA complexes on the (-80)77/G promoter fragment, compared to formation on the TIIG promoter fragment. An E M S A was performed exploring the formation of RPOA on the (-80)777G promoter fragment as a function of Spo0A~P concentration. In the absence of RNAP, no Spo0A~P bound to the promoter fragment (Fig. 28) even after prolonged overexposure (not shown). When RNAP was present, increasing amounts of Spo0A~P led to the formation of increasing amounts of RPOA, and correspondingly less RP complexes, supporting the idea that the two complexes are mutually exclusive and that Spo0A~P converts RP complexes into RPOA complexes. There were three differences compared to the Spo0A~P binding titration on TIIG promoter fragment (Fig. 24C). First, substantial amounts of RPOA were not formed until a concentration of 400 nM Spo0A~P compared to 200 nM Spo0A~P at the 77/G promoter 8 4 (SO)TIIG 0 100 200 300 400 500 600 700 800 [Spo0A~P] (nM) Figure 28. Complex assembly on (-80) TIIG - SpoOA~P titration. Binding reactions containing 2.5 nM radiolabeled (-80)77JG DNA, 0 to 800 nM Spo0A~P and 20 nM RNAP when present, were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hours at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film or phosphor screens for 2 hours and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. Fractional binding was calculated by dividing the units representing shifted complexes (RP) or supershifted complexes (RPOA) by the total counts per lane. Representative films of Spo0A~P binding titrations at 0 and 20 nM RNAP are shown. Representative data from one experiment of three is shown; POA (closed squares), RP (open circles), RPOA (closed circles). 85 fragment. Second, the formation of RPOA never saturated and some RP complex remained over the range of SpoOA~P concentrations tested. Last, 500 nM SpoOA~P was required to achieve half-maximal RPOA formation on the (S0)TIIG promoter fragment. This was 2.5x more SpoOA~P than was required to achieve half-maximal RPOA formation on the TIIG promoter fragment, a remarkable effect considering that the removal of the D N A between -100 and -80 would not be expected to impair Spo0A~P-DNA interactions at the promoter-proximal OA boxes or Spo0A~P-oA interactions. This requirement for elevated concentrations of Spo0A~P to form RPOA in the absence of upstream OA boxes was suggestive of a role for the site 1.1 and 1.2 OA boxes in increasing the local concentration of Spo0A~P at spoIIG. 3.8 Complex Formation on Variants of the TIIG Promoter Fragment. 3.8.1 Complex Formation on TIIGlDown. While the promoter-distal OA boxes seemed to play a role in increasing the local concentration of Spo0A~P to enhance the formation of RPOA, the results of the RNAP titration on (-80)77/G (Fig. 26) suggested that they might serve an additional function. Specifically, the D N A between -100 and -80 might impair RNAP binding. Spo0A~P could prevent that inhibition by occupying that region and occluding RNAP contact. Alternately, Spo0A~P bound to the promoter-proximal OA boxes could re-model RNAP contacts, preventing interaction of RNAP with the D N A between -100 and -80. In effect, promoter-proximal bound Spo0A~P could enhance RNAP binding by counteracting the negative influence of the D N A between -100 and -80. Removal of upstream D N A has been observed to improve the initial RNAP-promoter interaction at lacUV5 (Ross and Gourse, 2005) and X P R (Davis et al, 2005). Since the effect of upstream D N A at lacUV5 is dependent on the presence of the aCTD (Ross and Gourse, 2005), the implication is that rapidly re-equilibrating non-specific contacts between the aCTDs and upstream D N A imposed additional energetic costs that compromise the establishment of contacts with the core promoter (Davis et al, 2005). Since eliminating Spo0A~P binding upstream without removing the D N A would distinguish whether Spo0A~P bound to the promoter-proximal OA boxes was sufficient to stimulate RNAP binding, a template, TIIGlDown, was created in which predicted specific contacts between Spo0A~P and the D N A were eliminated based on the crystal structure of SpoOAc bound to OA boxes (Zhao et al, 2002). 86 Figure 29. Complex assembly on TIIGlDown. The TIIGlDown DNA fragment is a modified TUG DNA fragment containing the spoIIG promoter sequence between -100 and -15, except that the promoter-distal OA boxes have been eliminated. Binding reactions containing 2.5 nM radiolabeled TIIGlDown DNA, 0 to 80 nM RNAP and 800 nM Spo0A~P when indicated, were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to film or phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. Fractional binding was calculated by dividing the Phosphorlmager units representing specific complexes by the total Phosphorlmager units per lane. Representative data from one experiment of three is plotted; RP (open circles), RPOA (closed circles). 87 An E M S A was performed with different concentrations of RNAP in the presence or absence of SpoOA~P to test this idea (Fig. 29). While slightly less RP complex was formed at lower RNAP concentrations, no significant differences in the formation of RPOA were observed at TIIGlDown relative to TUG. SpoOA~P did not appear to bind to the mutated promoter-distal OA boxes; no SpoOA~P-DNA complexes were observed in the absence of RNAP according to the E M S A data, and the Rf value of RP0A7y/G/Dow/i was intermediate between that of KPwciDown and RPOA77/G (0.23 for RPOATHGIDOW,, compared to 0.20 for RPOA77 /G and 0.27 for RPTIIGIDOWH = R P T O G ) - RNAP binding to TIIGlDown was enhanced solely in consequence of interactions between SpoOA~P and the promoter-proximal OA boxes. As at (S0)TIIG (Fig. 26) some RP complex remained in the presence of 800 nM Spo0A~P. This is consistent with the earlier assertion that the promoter-distal OA boxes acted to increase the local concentration of Spo0A~P, but did not appear to be involved in the interaction of RNAP and the upstream DNA. A complex with lower mobility was occasionally observed on TIIGlDown, but it was not investigated since it represented only a small proportion of the shifted template. 3.8.2 Complex Formation on the TIIG2.1Up and THG2.1Down Promoter Fragments. The promoter-proximal tandem SpoOA binding sites overlap the -35 sequence at the spoIIG promoter (see Fig. 9). The binding site consists of two OA boxes, 2.1 (located from -53 to -47 inclusive) and 2.2 (located from -43 to -37 inclusive). The 2.1 OA box was mutated away from (TIIG2.1Down), and towards (TIIG2.1Up) the predicted optimal OA box consensus sequence (Zhao et al, 2002). E M S A titration experiments were performed using these promoter fragments to examine the effect of mutations in the 2.1 OA box on complex formation. The 2.2 OA box was not mutated as it overlaps with the -35 element and most mutations in this region severely reduce transcription in vivo (Satola et al, 1991). Spo0A~P binding to the TIIG2.1 Up promoter fragment was increased relative to binding to the wild type promoter (compare Fig. 30A to Fig. 24C). Two complexes were observed. Remembering that the D N A fragment contains two SpoOA binding sites each containing a pair of OA boxes, the complexes likely correspond to the occupation of a single binding site by one Spo0A~P dimer, and then the occupation of both OA box pairs by two Spo0A~P dimers. The fact that this fragment formed these two complexes and that the wild type TUG promoter 88 [SpoOA~P] (nM) [SpoOA~P] (nM) Figure 30. Complex assembly on TIIG2.1Up and TIIG2. IDown. (A) The TIIG2.1Up DNA fragment is a modified TIIG DNA fragment containing the spoIIG promoter sequence between -100 and -15, except that the sequence of the OA box immediately upstream of the -35 element has been altered towards consensus (thicker orange arrow). Binding reactions containing 2.5 nM radiolabeled TIIG2.1Up DNA, 0 to 800 nM Spo0A~P and 20 nM RNAP where indicated, were composed and allowed to equilibrate at 37 °C. After 2 min, loading buffer containing DNA to eliminate non-specific binding was added and the samples immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 rnA to separate complexes from free DNA. Gels were dried and exposed to film overnight or phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. Fractional binding was calculated by dividing the Phosphorlmager units representing specific complexes by the total Phosphorlmager units per lane. Representative data from one of three experiments using 20 nM RNAP is shown; POA (closed squares), RP (open circles), RPOA (closed circles). (B) (A) The TIIG2. IDown DNA fragment is a modified TIIG DNA fragment containing the spoIIG promoter sequence between -100 and -15, except that the sequence of the OA box immediately upstream of the -35 element has been altered away from consensus (thinner orange arrow). The assay was performed as described in (A). Representative data from a single experiment using 20 nM RNAP is shown; POA (closed squares), RP (open circles), RPOA (closed circles). 89 fragment does not, suggests that the first complex observed utilized the site 2 OA boxes. As anticipated, the base substitutions in TIIG2.1 Up had little effect on RP complex formation, as the fractional binding was about 0.4 for both labeled TIIG and TIIG2.1Up in the absence of SpoOA~P (compare Fig. 30A to Fig. 24C). With the TIIG2.1 Up promoter fragment, only supershifted complexes were observed at SpoOA~P concentrations of 200 nM or greater. Half-maximal formation of RPOA occurred at 100 nM Spo0A~P, half the concentration necessary for supershifted complex formation with the wild type TIIG promoter fragment. In addition, at 800 nM Spo0A~P the RP0A/RP ratio for complexes formed at TIIG2.1Up was 1.8 compared to an RP0A/RP ratio for complexes formed at TIIG of 1.25. Thus, improving the binding of Spo0A~P to the promoter fragment improved the formation of RPOA complexes. This is consistent with in vivo data in which a mutation introduced in the 2.1 OA box enhanced spoIIG transcription (Baldus et al, 1994). As expected, Spo0A~P did not bind to a TIIG2.1Down promoter fragment (Fig. 30B). The characteristic streaking of the free D N A in the presence of Spo0A~P that was attributed to unstable interactions between Spo0A~P and the wild type promoter fragment were absent on TIIG2.1Down. This indicated that complexes between TIIG2. IDown and Spo0A~P were too unstable to enter the gel. The binding of RNAP alone to the labeled fragment was reduced relative to binding to TIIG (fractional binding of 0.32 for the TIIG2.1Down promoter fragment as compared to 0.4 for the TIIG promoter fragment). The maximum amount of supershifted complex formed in the presence of Spo0A~P was much reduced relative to that formed on the wild type promoter fragment (fractional binding of 0.36 for the TIIG2.1Down promoter fragment compared to 0.59 for the TIIG promoter fragment). In the case of the TIIG2.1Down fragment, some of the RP complex remained, even at 800 nM Spo0A~P. With the TIIG2. IDown promoter fragment, half-maximal formation of the supershifted complex occurred at 500 nM Spo0A~P, 2.5x more than the amount required for the TIIG promoter fragment. Although RNAP still recruited Spo0A~P to the TIIG2. IDown promoter fragment to form RPOA, the RP0A/RP ratio was reduced to 1.13 compared to an RP0A/RP ratio of 1.25 for complexes formed at TIIG. Together these results suggested that although the contacts between Spo0A~P and the 2.1 OA box were not essential to recruit Spo0A~P to RP complexes, SpoOA~P-2.1 OA box interactions contributed to the formation of RPOA complexes. 90 3.9 Relative Stabilities of Complexes Formed on TIIG, TIIGllUp and TIIG2.IDown. Transcription activators that recruit RNAP stimulate transcription by enhancing or stabilizing the initial binding of RNAP to specific promoters. Alternatively, activators that stimulate a post-recruitment step (such as XcT) do so by stabilizing an intermediate complex during transcription initiation (Dove et al, 2000). Previous results indicated that SpoOA~P activates transcription by stabilizing an intermediate complex. Formally however, SpoOA~P could equally well stimulate transcription by destabilizing the initial RP complex if that complex acted as a kinetic barrier to initiation. Because the binding of RNAP to the spoIIG promoter was fast and survived gel electrophoresis, I considered the possibility that this latter mechanism was responsible for the stimulatory effect of SpoOA~P. A means to distinguish stabilization of an intermediate from destabilization of the initial complex would be to follow the decay of RP and RPOA complexes formed at TIIG, TIIG2.1Up and TIIG2. IDown. If SpoOA~P acts to destabilize RNAP-promoter complexes, RPOA complexes should decay faster than RP complexes. RPOA complexes formed with the TIIG D N A fragments were more stable than RP complexes (Fig. 31 A). Specifically, in the absence of SpoOA, RNAP dissociated from the wild type TIIG fragment with a rate of (3.7 ± 0.6) x 10"3 s"1, while RPOA complexes decayed with a rate of (1.9 ± 0.5) x 10"3 s"1. Spo0A~P stabilized the interaction of RNAP with the TIIG promoter fragment by a factor of 2. Similar results were obtained for the complexes formed on the TIIG2.1Up D N A fragment except that RPOA complexes were 4x more stable than RP complexes (compare Fig. 31B with 31 A). RP complexes formed with the TIIG2.1 Up D N A fragment decayed at a rate of (3.7 ± 0.6) x 10"3 s"1 identical to that of RP complexes formed with the TIIG D N A 3 1 fragments, while KP0A-TIIG2.1Up complexes dissociated with a rate of (1.0 ± 0.6) x 10" s" . By comparison, the complexes formed on the TIIG2. IDown D N A fragment were much less stable than those formed on the TIIG D N A fragment (compare Fig. 31C with 31 A). Both RP and RPOA complexes dissociated with rates of (20 ± 8) x 10"3. Improving Spo0A~P binding to OA box 2.1 enhanced the stabilizing effect of Spo0A~P on RNAP-promoter complexes, while reducing/preventing Spo0A~P binding to OA box 2.1 completely eliminated Spo0A~P-mediated stabilization. These data are highly supportive of a model in which Spo0A~P acts by stabilizing an intermediate during the transcription activation process at the spoIIG promoter. This 91 A. TUG B. TIIG2.1Up TIIG2.1Down 300 600 Time (s) Figure 31. Complex stability on TUG, THG2.1Up and TIIG2.1Down. Binding reactions containing 2.5 nM radiolabeled TUG (A), TIIG2.1Up (B), and TIIG2.1Down (C) DNA, 10 nM RNAP and 800 nM Spo0A~P when present, were composed and allowed to equilibrate at 37 °C. After 2 min, a 20-fold excess of unlabeled competitor promoter fragments was added. Samples were removed at the indicated time intervals after the addition of competitor and added to loading buffer containing DNA to eliminate non-specific binding. Samples were immediately loaded onto a running 4.5% non-denaturing polyacrylamide gel. Electrophoresis was continued for 1.25 hr at 20 mA to separate complexes from free DNA. Gels were dried and exposed to phosphor screens for 2 hr and then scanned on a Phosphorlmager SI using ImageQuant 5.2 software. The fraction of complex remaining was calculated by dividing the fractional binding at the indicated times by the fractional binding immediately prior to the addition of unlabeled competitor. Rate constants indicated in the text are the average of four experiments, but the data shown is from a single representative set of experiments using 20 nM RNAP; RP (open circles), RPOA (closed circles). 92 complex appears to form by recruitment of SpoOA~P to the D N A by the RP complex and is dependent on the binding of SpoOA~P to the 2.1 OA box. 93 DISCUSSION These experiments have addressed how SpoOA~P stimulates transcription from the spoIIG promoter, an essential step in the commitment to sporulation. In the Discussion, I wi l l first point out technical advances that allowed new information to be uncovered, and then present a refined model for transcription initiation at spoIIG, including caveats based on the work collected in this thesis. Transcription initiation at spoIIG wi l l be compared to that at other wel l-studied promoters, primarily performed using E. coli R N A P , and the differences in how R N A P s from B. subtilis and E. coli initiate transcription wi l l be discussed. The mechanism by which SpoOA~P stimulates transcription initiation is compared to other bacterial activators, and general principles underlying prokaryotic transcription wi l l be presented. 1 Technical Advances. The principal technical contributions of work presented in this thesis were improvements in R N A P purification, radiolabeling of D N A and the E M S A system. 1.1 RNA Polymerase Purification. The previous R N A P purification made use of PEG/Dextran phase separations, ammonium sulfate precipitations, size exclusion chromatography, D N A affinity chromatography, a preparative glycerol gradient ultracentrifugation and (sometimes) heparin affinity chromatography. The new R N A P purification scheme eliminated the glycerol gradient where losses of up to 50% of R N A P activity were typical, and rearranged the order of chromatographic steps to separate R N A P from other proteins on the basis of size exclusion, then heparin affinity and finally by affinity for D N A . Gradient elution rather than step elution of R N A P from a 32 heparin-agarose column eliminated a contaminating nuclease that removed P from radiolabeled D N A in E M S A and footprinting assays. On the basis of its 3' -> 5' nuclease activity, molecular mass and the similarity of purification schemes, the offending nuclease has been tentatively identified as A d d A B , the R e c B C D homolog in B. subtilis (Chedin et al, 1998; Shevelev and Hubscher, 2002). The addition of an ultrafiltration step to concentrate R N A P activity further and storage of the enzyme at -70 °C were minor modifications but significantly 94 improved the purification. Relative to a typical RNAP preparation using the previous protocol, this purification scheme increased the activity more than 5x, tripled the yield and at least doubled the lifetime of the enzyme. This dramatically improved all assays and permitted the use of a broader, more meaningful range of RNAP concentrations for binding studies. 1.2 Radiolabeled Template Synthesis. End labeling of primers and PCR synthesis of radiolabeled D N A fragments instead of end-labeling restriction fragments increased the specific activity of templates and permitted the generation of fragments of multiple sizes and sequences. Obviously, D N A sequence dependence could be easily assessed since mutagenesis was simple and rapidly performed. As well, using PCR products freed me from the limitations imposed by useful restriction sites and enabled a thorough investigation of complex formation on various truncated fragments that proved insightful. 1.3 Alterations in E M S A Techniques. During the course of this study many E M S A experiments were attempted. Earlier attempts were plagued by low levels of complex detection despite the fact that transcription assays showed high levels of template usage. Dissociation within the gel was problematic but believed the lesser of two evils; it had been necessary to run EMSAs for three hours to resolve complexes formed on the 234 base pair Hindlll-BamHI fragments used. Even then, RPOA complexes could not be resolved from RP complexes. To overcome these limitations, 1 used smaller D N A fragments that had two principal advantages. First, the samples could be electrophoresed for one-third the time reducing the amount of dissociation within the gel and providing a more accurate measurement of RNAP affinity for spoIIG. Secondly, it permitted the resolution of RPOA complexes from RP complexes and therefore a direct assessment of the early stages in transcription initiation. In sum, these improvements converted the E M S A as performed in our lab from a qualitative and relatively uninformative assay to a quantitative, highly reproducible, and penetrating one. 95 There is a discrepancy in SpoOA~P binding as assayed by DNase I footprinting and EMSAs. SpoOA~P clearly protects the spoIIG promoter from DNase I digestion, while complexes formed between spoIIG promoter fragments and SpoOA~P are scarcely observed (compare Fig. 20 to Fig. 21). I believe that this reflects the intrinsic instability of Spo0A~P-5po/ /G complexes and differences in how the assays are performed. I subjected protein-DNA complexes to DNase 1 digestion for 15 s after a binding equilibrium had been reached. Dissociation of complexes over 15s liberates free D N A that may be nicked, resulting in an underestimate of the extent of protection. By comparison, protein-DNA complexes assayed by gel shift take at least 90 s to enter the gel, and continue to decay throughout the duration that the E M S A is run (though at a reduced rate because of the caging effect within the polyacrylamide matrix). I believe this instability is an essential part of the mechanism by which Spo0A~P stimulates transcription at the spoIIG promoter and is consistent with kinetic data demonstrating that SpoOA~P stimulates isomerization rather than the initial binding of RNAP (Bird et al., 1996). It would have been preferable to explore complex formation using an equilibrium assay like fluorescence anisotropy or a filter binding assay that is more nearly an instantaneous sampling of equilibrium binding. However, practical considerations limit the usefulness of fluorescence anisotropy to interactions with D N A fragments up to about 45 base pairs, and neither technique can resolve between RP and RPOA complexes. 2 A Model for Transcription Initiation at the spoIIG Promoter. The principal results of this thesis were that SpoOA~P activates transcription from the spoIIG promoter by stimulating a rate-limiting transition between the initial interaction of RNAP and the promoter, and the complete exposure of the non-template strand of the -10 element. SpoOA~P was necessary for RNAP-mediated D N A strand separation but played no role in the energetics of D N A melting. In other words, promoter denaturation was predominantly driven by interactions between RNAP and the promoter. Evidence was presented that SpoOA~P and RNAP cooperated to form a closed intermediate that preceded promoter melting, and that RNAP recruited SpoOA~P to this closed intermediate. Roles for specific regions of the spoIIG promoter in complex formation were defined. 96 In conjunction with earlier work (Bird et al, 1996; Rowe-Magnus and Spiegelman, 1998b), I propose that transcription initiation proceeds through the following minimal series of intermediates at this promoter. R + P <-» R P C + OA <-> RPOAc <-> RP0A O i + pppApA + GTP <-> RPOA, + NTPs -> R E + OA A pictorial representation of this model is presented in Fig. 32. Initially RNAP binds to spoIIG to form RPc- This complex completely protects the D N A from DNase I digestion between -60 to -10 and partially protects the D N A between -10 to +4 (Fig. 21). The D N A located between -80 and -35 appeared to be critical for RPc formation (Fig. 25 and Fig. 26). While the D N A between -80 and -58 is located further upstream than canonical UP elements (Gourse et al, 2000), it is reminiscent of the UP elements contacted by E. coli a subunit CTDs. It seems possible that at least one aCTD binds specifically between -67 and -57 at the spoIIG promoter. This region shares 9 of 11 positions with the consensus UP element distal subsite defined in E. coli (Estrem et al, 1999) and removal of this D N A dramatically reduced RNAP binding (Fig. 26). By analogy with UP element usage in E. coli, the other aCTD may bind non-specifically downstream of -57, although the reduction in RNAP binding with the removal of D N A between -80 and -68 suggests that one aCTD may occupy this region at least part of the time. In E. coli it has been proposed (Ross et al, 2003; Typas and Hengge, 2005) that an equilibrium might exist whereby both a subunit CTDs might share a single subsite, or one aCTD may bind to each subsite. I propose that contacts between the aCTDs and the DNA, and between a \ and the -35 element, likely position RNAP on the promoter. This binding is tight and rapid, and in this intermediate formed on a linear template, a \ blocks access to OA box 2.2 and oA2 cannot reach the -10 element. In the next stage, the bulk of RNAP can slide along the D N A although the aCTDs remain bound upstream and tether RNAP to the promoter. If RNAP moves sufficiently far downstream, OA box 2.2 is revealed and the combination of Spo0A~P-DNA interactions plus Spo0A~P-aA interactions defined genetically (Baldus et al, 1995; Buckner et al, 1998; Hatt and Youngman, 1998; Schyns et al, 1997) recruit Spo0A~P to the promoter to form R P 0 A c (Fig. 24). Spo0A~P-DNA interactions are critical as mutations in OA box 2.1 affect both the amount (Fig. 97 R P O A Q I Figure 32. A model for transcription initiation at the spoIIG promoter. In the first step, RNAP (R) binds reversibly to the spoIIG promoter (P) primarily via aCTD contacts to the DNA between -80 and -59 to form RP C . RNAP cannot productively engage downstream promoter elements. SpoOA~P binds to site 1 OA boxes to increase the local concentration of SpoOA~P. In the second step, RNAP diffuses downstream while maintaining aCTD-DNA contact. This exposes the weak OA boxes at site 2. Interactions between RNAP and SpoOA~P recruit SpoOA~P to the site 2 OA boxes to form RP0A c . From this downstream-shifted position, RNAP can contact the downstream promoter elements. SpoOA~P prevents RNAP from sliding back and SpoOA~P-o4 contacts stabilize RNAP during the formation of the first open complex, RPOA 0 1 , in which the DNA strands are separated between -14 and -3. In the presence of pppApA and GTP, RNAP can initiate transcription to form a RNA trimer (red) exposing the transcription start site; the promoter is denatured between -14 and +2. In the presence of all four NTPs, RNAP can enter a phase of processive elongation, RP E , and jettison the a subunit. 98 30) and stability (Fig. 31) of RPOA complexes. Interactions between a A 4 and the sequence immediately downstream of the -35 element may play a role in the formation of RPOAc as has been suggested (Kumar et al., 2004b). It seems likely that the ovSpoOA-P interface at the spoIIG promoter functionally substitutes for specific -35 element-o"4 interactions that are observed at typical promoters. Sliding of RNAP leads to protection of the promoter downstream to cover the transcription start site. From this position, RNAP has access to the -10 element. Recruitment of Spo0A~P by RNAP affords the stabilization of a kinetic intermediate long enough to allow the energy-dependent step required for D N A strand separation. Therefore the role of SpoOA~P is to prevent decay of this downstream-shifted RPOAc intermediate back to the upstream RPc form. Re-positioning accounts for the observation that Spo0A~P is essential for open complex formation (Rowe-Magnus and Spiegelman, 1998b) without influencing the energetics of promoter melting (Seredick and Spiegelman, 2004). Two open complexes form at this promoter. The first open complex, RPOAoi, contains single-stranded D N A from -14 to -3 (Rowe-Magnus and Spiegelman, 1998b). Spo0A~P is required to maintain the position of RNAP until the -10 element is fully single-stranded (the transition from SpoOA~P-dependent transcription to SpoOA~P-independent transcription occurs as the single-stranded region of heteroduplex templates expands to include the entire -10 element; Table 1), at which point contact with the exposed bases of the non-template strand positions RNAP (Fredrick and Helmann, 1997; Qiu and Helmann, 1999). The effect of SpoOA during exposure of the non-template strand of the -10 element is evident in Fig. 14B. The enhanced accessibility of thymines to permanganate in the presence of SpoOAc supports the idea that Spo0A~P stabilizes RNAP even after promoter melting has nucleated. A second open complex, RPOAi, is observed with the inclusion of pppApA and GTP and contains single-stranded D N A from -14 to +2 with the formation of an RNA trimer (Rowe-Magnus and Spiegelman, 1998a). While no direct evidence was obtained to support its existence, it seems intuitive that a second open complex, R P O A 0 2 , exists in which the D N A has been melted past +1 permitting selection of the initiating nucleotides and phosphodiester bond formation. The fact that this complex cannot be detected could be explained if R P O A 0 2 collapses to RPOAoi so rapidly as to preclude permanganate modification of thymines at +1 and +2. Inclusion of the remaining NTPs allows extension of the nascent transcript as is easily observed by inclusion of ATP and GTP. 99 3 Caveats of the Proposed Model. 3.1 The Role of a-UP Element Interactions in B. subtilis. In comparison to the E. coli a subunit (Gourse et al, 2000), relatively little is known about the a subunit in B. subtilis and its role in the recognition of UP elements. Based on deletion analyses, early reports noted that AT-rich sequences between -70 and -40 at the spoVG promoter were required for high levels of activity in vivo and in vitro (Banner et al, 1983; Frisby and Zuber, 1991), and that sequences upstream of the -35 element were essential for high expression of the autolysin gene cwlB (Kuroda and Sekiguchi, 1993). A thorough study at the flagellin promoter (P/,ag), demonstrated that isolated a 2 bound to an AT-rich region between -70 and -32, and that this interaction stimulated transcription by increasing the affinity of RNAP for the promoter (Fredrick et al, 1995). Similarly, AT-rich sequences upstream of the strong R N A P o A -dependent A2c, A2b and C2 promoters of the lytic B. subtilis bacteriophage cp29 are necessary for high level expression, and act as docking sites for aCTD (Camacho and Salas, 2004). While UP element subsites generally lie one and two helical turns upstream of the -35 element (Camacho and Salas, 2004; Fredrick et al, 1995; Gourse et al, 2000), the subsites at the A2b promoter lie two and three helical turns upstream of the -35 element (Camacho and Salas, 2004), an arrangement analogous to that found at the spoIIG promoter. This plasticity in promoter architecture is believed to be permitted by the long, flexible linker connecting the aCTD to the N-terminal domain of the subunit (Blatter et al, 1994; Jeon et al, 1997). In E. coli at least, RNAP can interact with activators at different locations or with displaced UP element subsites as long as the appropriate helical phasing is maintained (Belyaeva et al, 1998; Blatter et al, 1994; Meng et al, 2001; Murakami et al, 1997; Newlands et al, 1992). Protein sequences of bacterial aCTDs are highly similar (Ebright and Busby, 1995; Gaal et al, 1996; Murakami et al, 1996), and it is reasonable to assume they would adopt similar structures and recognize similar D N A sequences. The residues in the aCTD that are critical for interaction with UP elements in E. coli (Gaal et al., 1996) are nearly identical in B. subtilis (Mencia et al., 1996) and reside within a pair of helix-hairpin-helix motifs that interact with UP element D N A in and across the minor groove (Ross et al, 2001; Yasuno et al, 2001). It seems likely that the 100 enrichment of AT-rich sequences between -80 and -35 noted in an informatic analysis (Helmann, 1995) represents a common feature of B. subtilis oA-dependent promoters. Moreover, the idea that the region between the promoter-distal and promoter-proximal OA boxes is recognized by the RNAP a subunit CTD is lent support by the near identity of the spoIIG promoter sequence with the UP element distal subsite consensus sequence in E. coli. 3.2 The Role of aCTDs in Transcription Initiation at the spoIIG Promoter. A proposed role for aCTD interaction with D N A between the OA boxes and the importance of this region in the formation of RP and RPOA (Fig. 26) seemingly contradicts earlier work in the laboratory that demonstrated that aCTDs were not necessary for SpoOA~P-stimulated transcription from the spoIIG promoter (Rowe-Magnus et al, 1998). That these results are reconcilable is due in part by the hypotheses being tested and the interpretative biases those expectations introduce, and in part by the way in which the experiments were performed. The work presented in this thesis was initiated to determine the contribution of specific regions and sequences in the spoIIG promoter in the formation of early intermediates during transcription. The proposal that aCTD contacts are essential to RP and RPOA complex formation was based on experiments performed on promoter fragments lacking all D N A downstream of-15. While this experimental design was deliberately chosen to limit our analysis to stages of transcription initiation preceding open complex formation, by eliminating the potential for contacts with the -10 element and transcription start site, it is reasonable to assume that these conditions exaggerated the relative importance of upstream contacts. However, the overlong spacing of conserved promoter elements, poor protection of downstream D N A in the absence of SpoOA~P (Fig. 21) and remarkably efficient formation of RP and RPOA complexes on TIIG (Figs. 18 and 19) argue that the early stages of transcription initiation are dominated by contacts between RNAP and SpoOA~P with promoter D N A between -100 and -15. In contrast, as many transcriptional activators require the a subunit of RNAP to function (Busby and Ebright, 1997) the earlier work (Rowe-Magnus et al, 1998) was undertaken to determine whether there was an essential interaction between Spo0A~P and the aCTDs. The primary 101 finding from the earlier work was that SpoOA~P could stimulate transcription independently of the aCTDs. This does not imply that the aCTD are unimportant in transcription from spoIIG. Although SpoOA~P stimulated transcription by RNAP reconstituted with mutant a subunits lacking either the C-terminal 15 or 59 amino acids (RNAPA15a and RNAPA59a) to the same extent as the wild type enzyme, the amount of transcript produced by RNAPA15a and RNAPA59a was reduced to less than 0.3 of that of the wild type RNAP. In fact, the defect in initiation as measured by gel shift was substantially worse. In the paper it was pointed out that the reduced transcription could be due to the need for aCTD during initiation, or possibly to lower specific activity resulting from the reconstitution of RNAP bearing mutant a subunits. As alluded to, the nature of the assays used in the previous work to seek evidence for SpoOA~P-aCTD interactions would also tend to underestimate the contribution of aCTD-DNA interactions in transcription. The single-round in vitro transcription assay with the spoIIG promoter as a template relies on the pre-incubation of RPOA complexes with ATP and GTP that permit initiation. In the absence of initiating NTPs during pre-incubation, no transcripts are produced, presumably because open, uninitiated complexes formed at spoIIG are exceedingly unstable to heparin challenge. (Heparin must be included to prevent multiple rounds of initiation.) a-DNA interactions act by enhancing the initial binding of RNAP to the promoter as well as stimulating RNAP isomerizations leading to open complex formation (Fredrick et al, 1995; Gourse et al, 2000; Ross and Gourse, 2005). Even though RNAPA15a and RNAPA59a might bind and isomerize slowly, the prolonged incubation period in the presence of iNTPs would be expected to minimize the differences between wild type and mutant RNAPs. The same rationale applies to KMnCU footprinting that directly assesses the formation of a late intermediate preceding transcript production. Finally, it is worth noting that the data in this thesis suggests that the role of the aCTD is to enhance RNAP binding and that Spo0A~P is recruited after that binding; the main conclusions in the two works are consistent. One might also consider the possibility that Spo0A~P stimulates transcription by an entirely different mechanism when faced with RNAP lacking aCTD. In lieu of upstream D N A interactions that seemed to be dominant for RP and RPOA formation on TIIIG, RNAP lacking the aCTDs and Spo0A~P might bind cooperatively to the promoter. In this scenario, the RPc 102 complex would be much less stable (or bypassed entirely) with Spo0A~P and RNAP binding simultaneously to the spoIIG promoter. SpoOA~P-aA interactions could still compensate for a mis-aligned -35 element until RNAP is able to denature the promoter and engage the non-template strand of the -10 element. Recruitment of RNAP to promoters with poor -35 elements by D N A bound activators interacting with 0 4 has been proposed for UhpA at uhpT, PhoB at pstS and MotA at T4 middle promoters (Chen and Kadner, 2000; Makino et al, 1996; Pande et al, 2002). The situation at spoIIG could be comparable to the binding of E. coli RNAP to the aidB or ada promoters via its aCTDs, which also results in the formation of a closed complex. At both the aidB and ada promoters, specific contacts between Ada bound immediately upstream of the -35 element and a 7 0 4 are necessary to drive transcription (Landini et al., 1998; Landini and Volkert, 1995). It should also be pointed out that the results obtained with mutant RNAP reconstituted with truncated aCTDs (Rowe-Magnus et al, 1998) did not preclude the possibility of interactions between Spo0A~P and the aCTDs, only that in the context of the in vitro assays, potential Spo0A~P-aCTD interactions are less important than those made between Spo0A~P and o A 4. A number of activators interact with both a and a including FNR, the bacteriophage p Mor protein, and CAP (Rhodius and Busby, 1998). In fact a sophisticated series of genetic screens employing semi-synthetic promoters, positive control mutants and suppressors was necessary to uncover the interaction between CAP and a 7 0 in E. coli (Bell et al., 1990; Rhodius and Busby, 2000a; Rhodius and Busby, 2000b; Williams et al., 1991; Williams et al., 1996). An equally sophisticated series of genetic experiments was required to demonstrate that RNAP and CAP formed a continuous nucleoprotein structure (aCTD-CAP-CAP-aCTD-c 7 0 4 RNAP) at the lac promoter whose integrity was necessary to enable RNAP to rapidly initiate transcription (Chen et al, 2003). It is tempting to speculate that Spo0A~P might act as a molecular spacer inserting between the proximal a subunit and a A 4 R N A P and so form an analogous extended nucleoprotein structure at the spoIIG promoter. Given the position of the promoter-distal OA boxes and the DNase I protection pattern (Fig. 21) it is possible that a continuous nucleoprotein structure comprising SpoOA~P-SpoOA~P-aCTD-aCTD-SpoOA~P-SpoOA~P-rjA 4RNAP forms to stimulate the rate at which RNAP can initiate transcription at spoIIG. Definitive evidence of the involvement of aCTD-DNA interactions in spoIIG transcription would require a thorough 103 analysis of kinetic quantities known to be affected by aCTD using aCTD mutants or reconstituted RNAP with truncated aCTD. Alternatively, wholesale substitution of the D N A sequence between the OA boxes could provide evidence for aCTD-UP element-like interactions as mutation of single base pairs in this region had little effect on transcription in vivo (Satola et al, 1991; Satola et al, 1992). 3.3 In vivo Concentration of RNAP and the Recruitment of SpoOA. It has been asserted that the most crucial factor in microbial gene regulation is the limited supply of RNAP (Browning and Busby, 2004). It is estimated that -2000 RNAP molecules are present in a growing cell of E. coli (Ishihama, 2000) or about half that of the number of genes. Much of the cell's RNAP is devoted towards the production of stable RNAs (ribosomal and transfer) required for translation, while some of the remainder is actively transcribing other genes or bound non-specifically to the genome. In proportion to the total number of promoters to which RNAP could bind, relatively little free polymerase is available. That being the case, a mechanism for Spo0A~P activation at the spoIIG promoter that relies on a promoter-bound but non-productive RNAP and polymerase-mediated recruitment of a transcription factor might seem somewhat unlikely to accurately reflect the situation in vivo. Several considerations mitigate potential difficulties in applying the in vitro model to in vivo conditions. First, sporulation is a post-exponential phase response to conditions in the environment that do not support rapid growth. The stringent response regulator ppGpp(p) downregulates rRNA synthesis, freeing up more R N A P c A for transcription of stage II sporulation genes, including spoIIG. In support of this notion, there is evidence that RNAP in B. subtilis is not limited during a growth rate shift up (Webb et al, 1982), implying that free RNAP may be available during the transition to a stationary phase of growth when the spoIIG gene is induced. Second, Spo0A~P binds much more poorly than RNAP to the spoIIG promoter. The for SpoOAc at spoIIG was recently estimated to be 1700 nM and shifted only a small fraction of the D N A (Fujita et al, 2005) while the Kd for RNAP at TUG was less than 10 nM. Activation of the spoIIG promoter requires that a positive feedback loop drive up the level of phosphorylated SpoOA (Fujita and Sadaie, 1998; Molle et al, 2003; Strauch et al, 1992; Strauch et al, 1993) and spoIIG responds to a higher level of Spo0A~P than many other 104 promoters (Chung et al., 1994; Fujita et al., 2005). Thus, the relative in vivo concentrations and Kd values argue strongly that RNAP binds to the promoter first, and that the RP complex recruits Spo0A~P. Third, a precedent for RNAP-mediated recruitment of transcription factors exists. In in vitro experiments, RNAP recruits and re-positions the phage-encoded repressor, p4, at the A2c promoter of the lytic B. subtilis bacteriophage q>29 (Monsalve et al., 1998). Last, the proposed model does not suggest that RNAP is trapped at spoIIG. The instability of RP and RPOA complexes formed at spoIIG relative to other promoters hints at the idea that RNAP "samples" the spoIIG promoter, transiently forming RPc and dissociating in the absence of Spo0A~P. If Spo0A~P is present, RP is modified to form R P 0 A c and then, under appropriate conditions, the ternary complex may rearrange, melting the promoter to form RPOAo- If iNTPs are available, RNAP will initiate transcription and may enter a phase of processive elongation. If iNTPs are not available, R P 0 A o collapses to RPOAc and rapidly dissociates. Thus the intrinsic instability of the complexes formed at spoIIG serve as integrative checkpoints where biochemical signals may be interpreted, influencing the progression of RNAP through successive stages in initiation. 3.4 Downstream Re-positioning of RNAP. Several observations have led to the proposal that the core of RNAP moves downstream during the transition between RPc and RPOAc to gain access to the -10 element. First, 0 4 of RNAP and Spo0A~P recognize similar sequences and have overlapping binding sites at spoIIG since OA box 2.2 occupies the same base pairs as the -35 element. Additionally, the spoIIG promoter is unusual in that 22 base pairs (instead of the optimal 17 base pairs) separate the -35 and -10 elements. The extra half-helical turn should prevent RNAP from making simultaneous contacts to both regions. In support of this idea, DNase I footprinting of the spoIIG promoter (Bird et al., 1996) revealed that RNAP only partially protects the promoter near the -10 element and transcription start site, while protection of this region was extended and enhanced in the presence of Spo0A~P. This is consistent with the idea that Spo0A~P acts to help RNAP overcome the extra half-helical turn within the spacer DNA. In fact, the stimulatory effect of Spo0A~P at the spoIIG promoter was entirely dependent on the overlong spacing of the conserved promoter elements. Reducing the spacer from 22 to 17 base pairs renders transcription independent of Spo0A~P (McLeod and Spiegelman, 2005). Consensus spacing, 105 and presumably simultaneous contact of the -35 and -10 elements by RNAP, bypasses the requirement for activated SpoOA. It should be pointed out that a low level of transcription did occur in the absence of SpoOA so that RNAP itself was capable of rearranging to allow initiation. However, such a process must occur either very slowly, or at very low frequency in the absence of SpoOA. While the data suggested that SpoOA~P acts by re-modeling the RP complex, it did not discriminate between SpoOA~P-assisted displacement of RNAP downstream and SpoOA~P-assisted conformational rearrangements in RNAP. Several facts favor a downstream displacement model for the activation of transcription at the spoIIG promoter. Crystal structures of oA4 from Thermus aquaticus complexed to a consensus -35 element and SpoOAc bound to the OA boxes from the abrB promoter have been determined since this work was initiated (Campbell et al, 2002; Zhao et al, 2002). As a A 4 / c 7 0 4 from B. subtilis, T. aquaticus and E. coli are highly conserved and recognize identical sequences within the -35 element I can reasonably predict how Spo0A~P and RNAP might bind to the spoIIG promoter. Inspection of the D N A contacts made in the two structures reveals that Spo0A~P and G A 4 2 are expected to make extensively overlapping and in some cases identical contacts with specific features of the D N A in this region (Fig. 33), precluding co-occupation of the same region of DNA. This is unlike the case of BvgA, a transcription factor that contacts the opposite helical face of the promoter as does RNAP (Boucher et al, 2003). Displacement of RNAP from the -35 element mediated by the mutually exclusive binding of Spo0A c and a A 4 2 would permit RNAP movement downstream to gain access to the -10 element. Closer inspection of the DNase I footprints (Figs. 17 and 18) provides further evidence consistent with a displacement mechanism. Spo0A~P uniformly protected the region between -55 and -35 from DNase I cleavage, while reactive sites at -45 and -55 remained when RNAP alone was present. When both proteins were present, the protection of this region was uniform; that is, protection of this region was Spo0A~P-like rather than RNAP-like. This data suggests that Spo0A~P , not RNAP, was bound to the -35 region of the D N A in RPOA complex. The available evidence indicated that the downstream displacement of RNAP was not likely to be driven by Spo0A~P. Spo0A~P bound very poorly relative to RNAP (Fig. 18), and therefore could not compete with RNAP for the -35 element/OA box. In fact, efficient binding of Figure 33. Overlapping aA and SpoOA~P contacts at the spoIIG -35 element. A schematic representation of the spoIIG promoter from -46 to -34 is shown including contacts predicted to be made by o A alone (blue), SpoOA~P alone (orange) and by both C J a and SpoOA~P (orange and blue). Bases to which all specific contacts are made in the two structures are identical at the spoIIG promoter despite differences in the sequence of oligonucleotides used in each of the crystal structures. Predictions of a A were based on the structure from Campbell et al, 2002; and predictions of SpoOA~P contacts were based on Mol A-Box 1 interactions from Zhao et al, 2002. 107 SpoOA~P depended upon the presence of RNAP (Fig. 24). This suggests that RNAP must slide downstream off the -35 element independently of any contributions from Spo0A~P and that when the binding pocket presented by both OA boxes and oA4.2 becomes exposed and properly aligned, SpoOA~P binds. Though RNAP must move to permit Spo0A~P binding, such a mechanism presents RNAP with the challenge of maintaining contact with the promoter after relinquishing the -35 element, but before establishing contacts with the -10 element. A potential solution to the structural rearrangements could lie in the apparent paradox that Spo0A~P recruitment (RPOA formation) required D N A upstream from the SpoOA binding sites. If RNAP was tightly bound to the D N A upstream of the core promoter, it would be able to transiently release the -35 region without dissociation, even though it has not located the -10 element. The recruitment of Spo0A~P stabilized RNAP that had released the -35 element while at the same time serving as a molecular spacer to position the aA 2.3 domain responsible for promoter melting. Moreover, recruitment of SpoOA~P by RNAP implies a strict temporal sequence of events. RNAP binding, primarily to sequences upstream of the -35 element, is a necessary prerequisite to the involvement of SpoOA~P and the efficient establishment of contacts near the -10 element and start site. This model helps to explain why some transcription occurs on linear templates even in the absence of SpoOA~P. RNAP can slide downstream while maintaining contacts to the upstream DNA, recognize and melt the -10 element, extend the denatured region past the transcription start site and initiate transcription, all entirely independently of SpoOA. It is likely that an intermediate state in which RNAP contacts both the region of D N A upstream of the -35 element and the -10 element simultaneously would be strained and highly prone to return to a state in which only the upstream portions of the promoter were occupied. Insertion of a Spo0A~P dimer at the site 2 OA boxes would stabilize the displaced form of RNAP and enhance the efficiency at which steps subsequent to downstream movement occur. This would enhance the rate at which RNAP can make the transition to an initiated complex. I propose that Spo0A~P maintains the position of RNAP over the -10 element, and stabilizes it until the -10 element is completely denatured and contacts with the non-template strand are established. 108 Direct evidence for this downstream shift was not obtained in this work despite considerable effort. High-resolution footprinting techniques using hydroxyl radicals, even when the reaction was performed on complexes isolated from free D N A by non-denaturing polyacrylamide gel electrophoresis, proved unsuccessful. Exonuclease III footprinting and DMS footprinting were similarly uninformative. The ineffectiveness of these techniques may be related to the instability of uninitiated complexes at the spoIIG promoter. While contacts to the upstream portion of the promoter are relatively well-defined and provide stability to RP and RPOA complexes, prior to initiation the downstream boundaries may be particularly labile. In principle, one could follow this movement by tethering a D N A cleavage agent to a single specific amino acid side chain of a protein (Meares et al, 2003). The reagent p-bromoacetamidobenzyl-EDTA-Fe (FeBABE) has one reactive group facilitating covalent attachment to cysteine side chains, whereas a second group coordinates an Fe(II) atom. Under appropriate conditions, Fe(II) can participate in the generation of hydroxyl radicals that attack deoxyribose units, resulting in D N A strand scission (Tullius and Dombroski, 1986). Since the diffusion of hydroxyl radicals is limited, the D N A is cleaved only in the immediate vicinity of the conjugated probe. Mutants a A proteins with cysteine-substituted residues in aA4 or a A 2 could be produced (sigA lacks cysteine residues), derivatized and reconstituted with RNAP core enzyme. Activation of the cleavage reagent in RP and RPOA complexes would reveal whether RNAP shifted downstream en masse, or whether Spo0A~P stimulated the extension of o A 2 without a concomitant shift in oA4. In conjunction, FeBABE could be incorporated into the HTH D N A binding motif of SpoOA (requiring the mutation of Cys7 and Cys41 in the N -terminus of SpoOA). While the working assumption has been that RNAP uses the -35 and then slides downstream to the -10 element, calling the spacer between the conserved promoter elements at spoIIG "overlong" may be something of a misnomer. RNAP clearly used the spoIIG -35 element when the spacer was shortened, since IIG17 was a fairly strong promoter (Table 6). However, at the wild type promoter RNAP needs to treat either the -35 element or -10 element as absent and since it uses the -10 element (Kenney et al, 1988; Kenney et al, 1989) it is possible that RNAP may largely ignore the -35 element. Genetic evidence that R N A P a A uses the -35 element at the spoIIG promoter has not been forthcoming despite the attempt (Kenney et al, 109 1989). An analogous experiments in E. coli in which an Arg588His mutation in o 4 2 suppressed a T T G A C A ->TTTACA substitution in the -35 element was successful (Gardella et al., 1989). Although SpoOA~P and a A 4 recognize nearly identical sequences and mutations in the -35 element would be expected to compromise the binding of both SpoOA~P and a A 4 2 , perhaps this might be tested by generating complementary suppressors in both SpoOA and oA. Both proteins make base-specific contacts to the 3 l d and 5 t h bases in the -35 element (the underlined bases in the T T G A C A sequence) that could be exploited by a genetic suppression approach. 3.5 SpoOA Recruitment and oA-Dependent Mutants in SpoOA. As binding of SpoOA~P alone to the spoIIG promoter is so weak, the model presented above suggests that the sum of protein-DNA and protein-protein interactions recruits SpoOA~P to the promoter-proximal OA boxes. The role of the SpoOA~P-oA (Baldus et al., 1995; Buckner et al., 1998; Hatt and Youngman, 1998; Schyns et al., 1997) interface in recruiting SpoOA~P has not been addressed in this thesis. However, the fact that RPOA still forms (SpoOA~P still binds) when the 2.1 OA box has been mutated away from consensus suggests that protein-protein interactions play an important (and perhaps primary) role in the binding of SpoOA~P immediately upstream of RNAP. However, the methodology described in this work provides a means by which mutations in SpoOA that specifically impair aA-SpoOA~P interaction can be investigated. Previously there was no means of in vitro analysis of such mutants since the only techniques available (DNase I protection and in vitro transcription) were uninfonnative. As a result, there was no means to discriminate at which stage of initiation such mutant SpoOA proteins were defective. The E M S A system using the truncated D N A templates described in this thesis would permit a quantitative investigation of the ability of SpoOA mutants to promote the formation of RPOA, the earliest stage in transcription initiation at the spoIIG promoter at which SpoOA~P acts. One would predict that SpoOA mutants defective in oA-dependent transcription may not form RPOA complexes, implying that RNAP cannot recruit them to the promoter, or that such complexes may form, but will not be stabilized by the binding of SpoOA~P. Such studies would allow identification of specific residues on SpoOA that were involved in binding and 110 stabilization. For example, i f the interactions of residues in the aE helix that specifically hinder RNAPcA-dependent transcription are required for SpoOA~P recruitment, such mutants would not allow the formation of RPOA complexes. On the other hand, i f the interaction stabilizes RPOA such mutants would form at high SpoOA~P concentrations but would be no more stable than RP complexes. In addition, the analysis of such mutants would clarify the role of the promoter-distal and promoter-proximal OA boxes; loss-of-function mutants near the aE helix in SpoOA would be expected to affect only processes occurring at the promoter-proximal OA box. 3.6 SpoOA Stimulation and DNA Topology. One unresolved problem in understanding how SpoOA~P regulates the spoIIG promoter is that while SpoOA~P is required for high level transcription from linear D N A templates (Bird et al, 1993), it is largely dispensable for transcription from supercoiled plasmids (Bird, 1994), yet essential for activation in vivo (Satola et al., 1991) where the promoter would be expected to be supercoiled. Reconciling the disparate requirements for SpoOA~P under these conditions requires a more thorough understanding of how supercoiling affects transcription initiation at the spoIIG promoter. While such studies have not been carried out, a few general points frame the problem. D N A in cells assumes a negatively supercoiled topology and the extent of supercoiling depends upon the relative activities of multiple topoisomerases and the binding constraints imposed by abundant nucleoid-associated proteins (Travers and Muskhelishvili, 2005). Control over topoisomerase activity is complex and difficult to study since opposing enzymes such as gyrases and topoisomerases act to balance each other's activity. For instance, the activity of D N A gyrase is controlled by the cellular energy charge that is tightly coupled to cellular growth rates (Hatfield and Benham, 2002), as is the production of the nucleoid-associated proteins HU, H -NS, IHF, Lrp, Dps, Fis, and CAP in E. coli (Travers and Muskhelishvili, 2005). Supercoiling affects the regulation of 7% of the genes in E. coli, some profoundly (Peter et al, 2004). In vitro it has been demonstrated that negative supercoils enhance the association of RNAP with rrnB PI (Leirmo and Gourse, 1991), as well as the amount (Ohlsen and Gralla, 1992) and rate of open complex formation (Revyakin et al, 2004) at that promoter. The general principle underlying the effect of supercoiling on transcription initiation is that negative superhelicity I l l facilitates the formation of RNAP-promoter complexes in which the D N A is writhed around the enzyme and that this torsional energy is transmitted through the enzyme to drive D N A strand separation at the AT-rich, and therefore relatively unstable -10 element (Travers and Muskhelishvili, 1998). Unlike E. coli, B. subtilis lacks CAP, Fis and H-NS- and IHF-like proteins. The B. subtilis homologue of Hu, HBsu is essential (Micka and Marahiel, 1992) and affects growth and nucleoid structure (Kohler and Marahiel, 1997). Detailed studies of the Bacillus Irp and dps homologies have not been carried out. In vitro, supercoiling bypasses the requirement for SpoOA~P for high level transcription from the spoIIG promoter. Two promoter alterations of linear templates presented in this thesis also bypassed the requirement for SpoOA~P and suggest mechanisms by which supercoiling may promote SpoOA-independent transcription of spoIIG in vitro. First, reducing the spacer length eliminated the dependence of transcription on Spo0A~P. The introduction of a negative supercoil may act similarly to reduce the effective distance or angular rotation of mis-aligned promoter elements. Second, RNAP efficiently transcribed templates with artificially denatured -10 elements. Theoretical considerations have led to the prediction that supercoiling enhances the transient opening of D N A (Vologodskii et al., 1979) and negative supercoiling predisposes torsionally labile AT-rich D N A to spontaneously denature (Drew et al., 1985; Spassky et al, 1988) that may permit transcription from spoIIG on supercoiled plasmids in a manner exactly analogous to that from the heteroduplex templates. Alternately, RNAP binding may drive conformational transitions of the D N A on supercoiled plasmids that promote denaturation of the -10 element. It is possible that the contacts RNAP makes upstream at the spoIIG promoter constrain the D N A in such a fashion as to predispose the least stable segment of the D N A in the immediate vicinity, the -10 element, to denature and thus relieve torsional strain. Such a phenomenon, termed supercoiling-induced duplex destabilization, operates at the E. coli ilv P Q (Sheridan et al, 1998) and /euF(Opel et al, 2004) promoters. In vivo however, since SpoOA is essential for transcription of the spoIIG operon, it is likely that it overcomes the repressive effects of at least one of the negative regulators of spoIIG transcription. 112 3.7 SinR and Soj. Two negative regulators act directly to oppose transcription initiation at the spoIIG promoter. One, Soj (also known as SpoOJA), is a ParA homolog involved in chromosome partitioning during sporulation (Ireton et al., 1994; Sharpe and Errington, 1996). Soj has been proposed to physically associate with the spoIIE, spoIIA, spoIIG and spoOA promoters on the basis of elevated frequencies of crosslinking to these promoters in vivo (Quisel and Grossman, 2000; Quisel et al., 1999). In early in vitro studies, Soj appeared to dissociate uninitiated RPOA complexes formed at the spoIIG promoter (Cervin et al., 1998b). Since Soj inhibited transcription of the SpoOA-independent IIG17 promoter much less efficiently than the SpoOA~P-dependent spoIIG promoter, it appears that Soj interferes with a SpoOA-specific function during initiation (McLeod and Spiegelman, 2005). The other negative regulator, SinR, is a small tetrameric D N A binding protein (Lewis et al., 1998) that binds to and represses transcription of a number of genes, including the sporulation-specific spoIIE, spoil A, spoIIG and spoOA promoters (Mandic-Mulec et al., 1995; Mandic-Mulec et al., 1992). SinR represses transcription from linear spoIIG templates without displacing RNAP, while Spo0A~P can overcome the repressive effect of SinR (Cervin et al., 1998a). Both Soj and SinR are produced during vegetative growth in Bacillus, and both proteins have antagonists that block their activity and are required for sporulation. It seems likely that in vivo Spo0A~P must operate in the presence of these negative effectors. One hypothesis of the mechanism of Soj action is that Soj inhibits transcription initiation by accelerating the dissociation of open complexes. A similar mechanism has been proposed for DksA, a transcription factor essential for the stringent response in E. coli. DksA binds directly to RNAP to reduce the half-life of open complexes formed at rrnB PI (Paul et al., 2004a). One would predict that Soj would be able to drive RNAP from the D N A in the presence of Spo0A~P on templates where the open complex was unstable (wild type spoIIG), but not from templates in which the -10 element is missing (TIIG), or artificially and stably denatured (NT 14/3). Soj would also not be expected to influence the interaction of RNAP alone with any of these templates. The effect of Soj on the dissociation rate could be measured in a transcription-based assay following relative transcription as a function of time following heparin challenge. However, the t\/2 for open complexes formed on linear spoIIG in the absence of Soj is less than 113 30 s, which may require the development of a stopped-flow kinetic assay system. The transcription-based assay may be effective on a supercoiled template, as the t|/2 for open complexes at rrnB P1 increased from 6 min on linear templates to 600 min on supercoiled templates (Paul et al, 2004a). Alternately it is possible that 2-aminopurine, an adenine analog whose fluorescence is quenched in double-stranded but not single-stranded D N A (Guest et al, 1991), could be incorporated into linear templates between -8 and -3, and the rate of loss of signal followed by fluorescence spectroscopy. SinR must inhibit transcription by a more subtle mechanism. One possibility is that SinR and Spo0A~P form mutually antagonistic nucleoprotein structures with RNAP at the spoIIG promoter. Unpublished data (G. Spiegelman) suggests that reversal of SinR-mediated inhibition of transcription from supercoiled templates may account for the requirement for SpoOA during in vivo activation of spoIIG. In this scenario, Spo0A~P, through its interaction with RNAP, would reverse the repressive influence of SinR by altering the topology of the promoter; it is noteworthy that Spo0A~P and SinR bind to partially overlapping sites (Cervin et al, 1998a). It may also be informative to consider that the binding of Spo0A~P to the spoIIG promoter would be predicted to limit binding of the aCTDs to 2 and 3 helical turns upstream of the -35 element. The binding of SinR may prevent or alter this helical phasing, either of which would be expected to alter the transmission of torsional energy to the -10 element. 4 Transcription Initiation. Current understanding of the mechanisms driving transcription initiation in bacteria draws overwhelmingly from studies performed using E. coli R N A P a 7 0 . On the basis of genetic and biochemical studies, B. subtilis R N A P o A appears to recognize identical promoter sequences and 70 catalyze D N A strand separation in an identical fashion to E. coli RNAPa . However, open complexes formed by R N A P a A are noticeably less stable (Rojo et al, 1993; Voskuil and Chambliss, 2002; Whipple and Sonenshein, 1992) and presumptive UP elements and extended -10 motifs occur with greater frequency in B. subtilis aA-dependent promoters than in E. coli o 7 0 -dependent promoters (Helmann, 1995). In contrast to promoters in E. coli where extended -10 motifs appear to compensate for absent -35 elements (Bown et al, 1997), oA-dependent 114 promoters with extended -10 motifs in B. subtilis are likely to have highly conserved -35 elements (Voskuil and Chambliss, 1998). Lastly, a non-conserved region in the N-terminus of E. coli o 7 0 is absent B. subtilis o A; it is not clear how the absence of this region affects initiation in B. subtilis. 4.1 Closed Complex (RPc) Formation. In the early stages of transcription initiation, RNAP forms closed complexes (Browning and Busby, 2004). This involves the docking of RNAP domains to some combination of modular promoter sequences, including UP elements, extended -10 motifs and the -35 and -10 elements. At well-characterized promoters to which RNAP can bind independently of activators, two closed complexes have been observed, RPci and RPc2 (Record et al., 1996), differing in the length of promoter D N A that RNAP contacts. RNAP associates with one face of the D N A and does not bind the start site in RPci, while the D N A around the start site is completely enveloped by RNAP in the final closed complex, RPc2- At promoters that are recognized poorly by RNAP, DNA bound transcriptional activators such as CAP serve as docking sites, permitting RNAP to make contacts with the -10 element and transcriptional start site (Lawson et al., 2004). By comparison, a striking feature of this work was the binding of RNAP to the TIIG promoter fragment, and the subsequent assembly of a ternary complex composed of RNAP, Spo0A~P and upstream promoter DNA. This observation suggested, at least at this promoter, that the -10 element and all D N A downstream were not essential for the earliest stages in transcription initiation. Based on structural models of RNAP-promoter complexes (Borukhov and Nudler, 2003; Murakami and Darst, 2003; Murakami et a l , 2002a; Naryshkin et al., 2000) and the 22 base pair separation of conserved promoter elements, one might predict that specific contacts between oA2.4 and the -10 element at the native spoIIG promoter would be unimportant, though it seems surprising that non-specific interactions with D N A downstream of the -35 element are also unnecessary. This situation is reminiscent of early models for transcription initiation in which promoter binding and promoter melting were proposed to be mediated independently by upstream and downstream promoter elements, respectively. Recent reports of promoter melting by a minimal RNAP assembly incapable of making upstream contacts supports the idea that 115 promoter denaturation can occur in the absence of contact to upstream D N A (Young et al., 2004). Similarly, the work presented in this thesis supports the idea that complex assembly can occur in the absence of downstream DNA. However, it is important to remember that such findings artificially isolate a limited number of events from what are probably an interdependent series of transitions on the native promoters. For example, on linear templates upstream sequences control the trajectory of D N A around RNAP and therefore influence the presentation of downstream elements for promoter melting and start site selection (Davis et al., 2005). It may be that on D N A with different topology, RNAP contacts upstream and downstream promoter elements simultaneously. It is not clear what relation RP and R P 0 A c at spoIIG have to RPci and RPc2 defined at activator-independent promoters using E. coli RNAP (Record et al., 1996). The definitive feature of RPc2 is the double-stranded protection of the promoter around the start site that was observed in low temperature footprints of both D N A strands. Such experiments with the spoIIG promoter were 70 not attempted. Envelopment of the start site by E. coli RNAPo is correlated with the acquisition of heparin resistance at least at A P R (Craig et al, 1998; Saecker et al, 2002) and /acUV5 (Buc and McClure, 1985; Spassky et al, 1985) and appears to be coupled to D N A strand separation. Heparin resistance at spoIIG is not acquired until synthesis of an R N A trimer (a stage necessarily following D N A strand separation), leading to the prediction that RNAP may not enclose the start site until this stage. In that case, a closed intermediate complex formed by RNAP in the presence of Spo0A~P at spoIIG would be structurally distinct from the E. coli RPc2, and as a result the effect of Spo0A~P would not simply be to convert an RPci-like complex to an RPc2-Hke complex. This delayed start site enclosure by B. subtilis RNAP may account for the observed instability of RP complexes formed at some promoters and could be further investigated by DNase I protection analysis of both strands at 7 and 37 °C in the presence of iNTPs and heparin at spoIIG and G2, a strong (p29 promoter at which B. subtilis RNAP makes stable complexes. It may be informative to perform the same series of footprints on the A2c promoter, a SpoOA-independent q>29 promoter that also requires initiation to form heparin-resistant complexes (unpublished observation, G.B.S). 116 4.2 Open Complex (RPo) Formation. In the later stages of transcription initiation, RNAP separates the D N A strands to form open complexes (Browning and Busby, 2004). Seminal work in the catalysis of strand separation was performed using B. subtilis RNAP, demonstrating the involvement of aromatic and basic residues in a 2 j in the nucleation of D N A melting (Aiyar et al., 1994b; Jones and Moran, 1992; Juang and Helmann, 1994b; Rong and Helmann, 1994). Contacts with the P and P' subunits maintain D N A strand separation (Brodolin et al., 2000; Naryshkin et al, 2000; Severinov and Darst, 1997; Studitsky et al, 2001). At most E. coli promoters, the D N A may be melted from the upstream edge of the -10 element to +2 (-14 to +2 at spoIIG), although intermediate open complexes have been observed at A P R and T7 A l in the absence of Mg (Suh et al, 1993; Zaychikov et al, 1997). Activators that increase the rate of open complex formation are postulated to stabilize the formation of a rate-limiting intermediate (Dove et al, 2000). The data presented in this thesis indicate that the thermodynamic properties of initiation are driven by transitions involving only RNAP and the promoter, and that Spo0A~P is not involved in the energetics of D N A melting. The thermal profiles of transcription from the nested series of artificially denatured templates provided insight into the process of D N A melting during open complex formation. The differences in transition temperatures, Tm, of transcription allowed an estimate of the energetic requirements of denaturing short segments of the spoIIG promoter by calculating the van't Hoff enthalpy (AHapparent) required to separate the strands. This determination of enthalpy required that RNAP-DNA binding equilibrated in the transcription reaction (Bird et al, 1996), and assumed that the differences between the Tm reflected the thermodynamic properties involved in D N A melting. Using this approximation, the differences in Ai/appai-em between initiation at spoIIG and NT14/13 or NT14/11 indicated that denaturation of the bases between -14 and -11 required 9 kcal mol"' (Table 7 and Fig. 34). The differences in A//apparent between initiation at NT14/11 and NT14/9 or NT14/7 suggested that denaturation of the 4 base pairs between -10 and -7 required an additional 9 kcal mol"1. Separating the D N A strands 4 base pairs further to denature -6 to -3 imposed an energetic cost of 11 kcal mol"' because of the difference in L\Happarent between NT14/7 and NT14/3. In total, formation of an open complex in which the D N A strands were denatured between -14 and -3 required 29 of the Figure 34. Thermodynamically separable steps in DNA strand separation at the spoIIG promoter. In the model, P, P' (light blue) and o (dark blue) interact with the promoter to drive DNA strand separation beginning at the upstream edge of the -10 element (white box) at base pair -14 and proceeding towards the transcription start site (bent arrow; +1). The portions of the promoter DNA that are buried within RNAP are grey. The A / 7 a p p values are calculated from the Tm of transcription from the heteroduplex templates as described in Table 6. A/7, represents the enthalpic cost of a single thermodynamic step that results in the melting of the promoter from -14 to -11. A/7 2 represents the enthalpic cost of a single thermodynamic step that results in the melting of the promoter from -10 to -7, and A/7 3 represents the enthalpic cost of a single thermodynamic step resulting in the denaturation of the promoter from -6 to -3 . A/7 4 represents the enthalpic cost of denaturing -14 to -3 in a single step. See text for details. 118 42 kcal mol"1 required for the overall transcription initiation reaction at the spoIIG promoter. The latter value is similar to the 41 kcal mol"1 estimated for transcription from lacUVS (Nakanishi et al, 1975) and 45 kcal mol"1 for open complex formation at T7 A l (Johnson and Chester, 1998). The apparent energy input for spoIIG promoter melting was strikingly discontinuous. Despite the requirement to denature an additional 2 base pairs, transcription from NT 14/13 required the same enthalpic input as did transcription from NT14/11. Transcription from NT14/9 and NT 14/7 also required the identical enthalpic inputs even though 2 additional base pairs need to be denatured at NT14/9. There also did not appear to be a simple relationship between the type (A:T versus G:C) of base pairs melted and the energetic cost. Denaturing the base pairs between -7 and -3(11 kcal mol"') was more energetically costly than denaturing the base pairs between -10 and -7 (9 kcal mol"1) even though both stretches contained 3 A:T and 1 G:C base pair. Similarly, denaturing the 4 A:T base pairs between -14 and -11 might have been expected to be less energetically costly than denaturing the 3 A:T and 1 G:C base pair between -10 and -7, but denaturing both stretches required 9 kcal mol"1. This would seem to imply stepwise melting, as found for the B. subtilis flagellin promoter (Chen and Helmann, 1997), and may reflect a series of conformational rearrangements in RNAP domains that enable precise control over D N A strand separation. In support of such a model, a mutant RNAP with a p subunit containing a deletion forms open complexes that separate only the upstream 4 to 6 base pairs (Severinov and Darst, 1997). No evidence for such intermediates was found by potassium permanganate footprinting using the spoIIG promoter, suggesting either that these proposed rearrangements are allosterically coupled (though thermodynamically separable), or perhaps that they interconvert more rapidly than the technique can resolve. 5 Transcription Activation. 5.1 SpoOA. The stimulatory properties of SpoOA reside in its C-terminal domain (Grimsley et al, 1994; Rowe-Magnus and Spiegelman, 1998a) and crystal structures of the C-terminal activation domain alone and complexed to D N A are available (Lewis et al, 2000a; Zhao et al, 2002). The 119 activation domain contains a helix-turn-helix D N A binding motif, a central bundle of three a-helices, and a helix protruding from one side (aE), connected by flexible linkers to adjacent helices. When a dimer of SpoOA~P sits on the DNA, the central axis of the aE helix is nearly parallel to the axis of the D N A helix. SpoOA~P binds D N A as a dimer (Asayama et al, 1995; Lewis et al, 2002), typically to direct repeats. The consensus binding sequence is 5'-ttTGTCGAAaaa-3' where the lower case bases are much less conserved (Liu et al, 2003; Molle et al., 2003). At the aA-dependent spoIIG and spoIIE promoters, Spo0A~P binds to sites overlapping the -35 element to stimulate transcription (Satola et al, 1991; Satola et al, 1992; York et al, 1992). The -35 and -10 elements at the spoIIG and spoIIE promoters are separated by 22 and 21 base pairs, respectively. This conserved promoter architecture suggests that Spo0A~P stimulates transcription from both promoters by an identical mechanism. Analysis of spoOA mutations that specifically eliminate transcription activation from aA-dependent promoters found that the mutants were found either in helix aE or in the flexible linkers connecting aE to the remainder of the domain (Buckner et al, 1998; Hatt and Youngman, 1998; Kumar et al, 2004a). Removal of the entire aE helix also abolishes transcription from spoIIG while not affecting repression of the abrB promoter, a clear indication that the protein still binds D N A (Kumar et al, 2004a). Table 1 lists all characterized mutations in spoOA affecting sporulation. Corresponding mutations in a A that specifically affect transcription from SpoOA-dependent promoters (Baldus et al, 1995; Schyns et al, 1997) and a mutation in SpoOA that suppresses SpoOA-dependent mutations in a A have been identified (Buckner et al, 1998). This mutation maps to aA4. 2. Structural modeling led to the prediction that an interaction between a negatively charged residue in SpoOA, Glu221, and a positively charged interaction in o A , Arg355, might also be involved. Simultaneous substitution of both amino acids enhanced transcription relative to single substitutions in either SpoOA or a A , but the "restoration" of activity in the double mutant was less than 0.4 of the wild type (Kumar et al, 2004b), suggesting that other effects of the substitutions may be in play. Regardless, the evidence is compelling that SpoOA activates transcription by interacting with o \ 2 through a flexible helix jutting from the central helical bundle responsible for D N A binding. 120 A fairly detailed picture of how Spo0A~P stimulates transcription at the spoIIG promoter can be constructed. Binding of Spo0A~P to the upstream OA boxes (site 1) appears to increase the local concentration of SpoOA~P available to bind the promoter-proximal OA boxes. SpoOA~P is recruited to the downstream OA boxes through a combination of SpoOA~P-DNA interactions and SpoOA~P-RNAP interactions. Once bound to the DNA, SpoOA~P positions RNAP by preventing it from sliding back upstream off the -10 element. Practically, this has the effect of facilitating D N A strand separation by stabilizing RNAP during the nucleation of D N A melting and increasing the rate at which RNAP establishes interactions with the non-template strand of the -10 element. The effect of SpoOA~P during open complex formation is limited; contacts between SpoOA~P and a A 4 2 are unnecessary when RNAP can exchange them for contacts between a A 2 and the single-stranded -10 element. It appears that SpoOA~P helps RNAP stay in the right place once it gets there. SpoOA controls transcription at no less than 54 promoters (Molle et al, 2003), of which its effects are now well-understood at only the spoIIG promoter. Although it is reasonable to presume that SpoOA might stimulate transcription from the aA-dependent spoIIE promoter by a similar mechanism since its consensus promoter elements are separated by 21 base pairs, SpoOA also stimulates transcription from aA-dependent promoters that lack this unusual promoter spacing. SpoOA also stimulates transcription from many oH-dependent promoters. In vitro analyses of the role of SpoOA at these promoters have barely been attempted, but the diversity in the positions and orientation of the OA boxes suggests the involvement of different subunits of RNAP and different surfaces in SpoOA (Spiegelman et al., 1995). How SpoOA represses transcription is also largely unknown. In fact, only one detailed study (Greene and Spiegelman, 1996) has investigated the mechanism by which SpoOA prevents transcription, though SpoOA directly represses twice as many genes as it activates (Molle et al., 2003). 5.2 Other Response Regulators. On the basis of their C-terminal effector domains, most response regulators that act as transcription factors can be classified into three major subfamilies, represented by NtrC, NarL, 121 and OmpR (Stock et al, 2000). SpoOA has a unique effector domain structure that defies categorization into any of these subfamilies (Lewis et al, 2000a). 5.2.1 The NtrC Subfamily. The mechanisms by which members of the NtrC subfamily of activators stimulate transcription likely have little relevance to how SpoOA stimulates transcription. These proteins activate transcription from RNAP holoenzymes containing a 5 4 (or its homologs) that are evolutionarily unrelated to a 7 0-like sigma factors. R N A P a 5 4 form closed complexes but cannot spontaneously isomerize to form open complexes (Buck et al, 2000). NtrC-like proteins use ATP hydrolysis, mediated by an A A A domain not found in other response regulators, to drive conformational rearrangements in the p' jaw domain of RNAP (Burrows et al, 2004; Popham et al, 1989; Wigneshweraraj et al, 2004; Zhang et al, 2002). These activators can stimulate transcription from solution without binding DNA (North and Kustu, 1997; Wyman et al, 1997) though they typically bind hundreds of base pairs upstream of the promoter and interact with RNAP through D N A looping (Wedel et al, 1990). 5.2.2 The NarL Subfamily. The NarL subfamily contains a number of well-characterized response regulators. The effector domain structure is also shared with GerE, the ultimate transcriptional regulator of spore formation, and auto-inducer homoserine lactone-dependent activators typified by LuxR (Ducros et al, 2001). Crystal structures are available for full-length NarL (Baikalov et al, 1996) as well as the isolated C-tenninal domain bound to D N A (Maris et al, 2002); the C-terminal domain of NarL forms a four helical fold including a classic helix-turn-helix motif. A bewildering number of these proteins and some of the genes they regulate have been identified. In many cases, D N A binding sites for these proteins have been found at particular genes. In relatively few cases is any information available regarding how these response regulators influence R N A polymerase and transcription initiation. RcsB stimulates RNAP binding at osmC PI through binding sites located just upstream of a poor -35 element (Davalos-Garcia et al, 2001; Sturny et al, 2003). ComA binds to sr/'between -79 and -61 (Roggiani and Dubnau, 1993), and is essential for transcription (Nakano et al, 1991). Disruption of the 122 interaction between ComA and RNAP by blocking the aCTDs results in the dissociation of ComA, but not RNAP, from the promoter. This is consistent with the possibility that RNAP might recruit ComA to the srf promoter in a similar fashion to the recruitment of SpoOA~P to the spoIIG promoter, though the subunits of RNAP mediating this recruitment would be different (Nakano et al, 2003). Probably the two best understood transcription factors in this subfamily are UhpA, which controls the expression of genes encoding sugar phosphate transporters in E. coli, and BvgA, which controls the expression of virulence determinants in Bordetella pertussis. At the uhpT promoter that lacks a -35 element, three dimers of UhpA bind cooperatively between -80 and -32 to recruit RNA polymerase through contacts with a 7°4(Chen and Kadner, 2000; Olekhnovich et al, 1999; Olekhnovich and Kadner, 1999). A precise analysis of the effect of UhpA on RNAP is complicated somewhat by the lack of RNAP binding in the absence of UhpA, and a secondary stimulation of transcription by CAP (Olekhnovich et al, 1999). Three dimers of BvgA also bind cooperatively to the fha promoter between -100 and -35 to recruit RNAP (Boucher et al, 1997; Boucher et al, 2001). It appears that BvgA recruits RNAP through the aCTDs, although it may do so indirectly since the aCTDs and BvgA proteins bind to opposite helical faces. Consequently, it has been suggested that bending of the D N A bound by BvgA would narrow the minor groove and enhance association of aCTDs with the D N A (Boucher et al, 2003). No evidence exists for contacts between BvgA and a, but it has been noted that the two proteins are appropriately juxtaposed to interact at the fha promoter. In fact, mutational analysis of members of this sub-family to define contact surfaces with RNAP subunits has yet to be reported. 5.2.3 The OmpR Subfamily. Several members of the OmpR subfamily have been intensively studied, but relatively little information is available regarding the mechanisms by which they stimulate transcription. Crystal structures are available for the full-length DrrD (Buckler et al, 2002) and DrrB (Robinson et al, 2003), the effector domains of OmpR (Kondo et al, 1994; Martinez-Hackert and Stock, 1997) and PhoB (Okamura et al, 2000), and of a dimer of PhoB bound to D N A 123 (Blanco et al, 2002). The structure of these activators includes an N-terminal four-stranded P sheet, a central three-helical bundle and a C-terminal P hairpin. A winged helix-turn-helix motif is responsible for D N A binding. Trans-activation mutants in OmpR (Pratt and Silhavy, 1994; Russo et al, 1993), PhoB (Makino et al, 1996) and PhoP (Chen et al, 2004), map to the turn of the D N A binding motif. Despite this similarity, OmpR and PhoB contact different subunits of RNAP. PhoB binds to the pstS promoter from -65 to -38 and recruits RNAP through interaction with a (Kim et al, 1995; Makino et al, 1993), while OmpR requires the aCTDs at ompC and ompF to stimulate transcription (Igarashi et al, 1991; Sharif and Igo, 1993; Slauch et al, 1991). Whether OmpR enhances RNAP binding or stimulates a step following RNAP recruitment has not been addressed. ResD, a member of this subfamily in B. subtilis, binds to the hmp promoter between -83 and -41 and likely recruits RNAP through interactions with the a subunit CTD (Geng et al, 2004; Nakano et al, 2003). Significantly, the turn of the D N A binding motif to which the trans-activation mutants map protrudes laterally from the D N A bound protein reminiscent of the aE helix in SpoOA (Fig. 35). One would predict this arrangement to be a common property of response regulators (and bacterial transcription factors in general). A laterally presented interaction surface is available to a helically-displaced DNA-bound binding partner. 5.3 C A P . E. coli CAP is the best-studied transcriptional activator. At several promoters, CAP stimulates transcription by RNAP in the absence of any accessory factors. As a consequence of sustained investigation at such promoters, a structural and mechanistic description of how CAP works is more nearly complete than for any other transcriptional activator (Busby and Ebright, 1999; Lawson et al, 2004). CAP was the first transcriptional activator to have its structure determined (McKay and Steitz, 1981) and structures of a binary CAP-DNA (Parkinson et al, 1996) and a ternary CAP-aCTD-promoter (Benoff et al, 2002) complex are available. Simple CAP-dependent promoters can be classified into two groups on the basis of the position of the D N A site for CAP, and the corresponding mechanism of activation. 124 Figure 35. Response regulator-DNA interactions and RNAP contact surfaces. Dimer-DNA interactions are presented from the side (left column) and top (right column). The DNA is 5' to 3'. (A) SpoOAc-OA box interaction. The G^activating region around the aE helix is orange and overlaid with the transparent mauve ovals. Based on PDB file 1LQ1 deposited by Zhao and Varughese, 2002 in the RCSB protein data bank (http://pdbbeta.rcsb.org/pdbAVelcome.do). (B) PhoB c -DNA interactions. The activation loop defined by positive control mutants in PhoB and OmpR that contacts o 7 0 in E. coli is overlaid with the transparent mauve ovals. Note that these mutations map to the flexible loop, not the green-yellow helix. Based on PDB file 1GXP deposited by Blanco et al., 2002 in the RCSB protein data bank. (C) NarL c -DNA interactions. No contact surface has been defined for NarL family members. Based on PDB file 1JE8 deposited by Maris et al., 2002 in the RCSB protein data bank. Figure constructed using PyMOL (Delano Scientific). 125 At Class I CAP-dependent promoters, a dimer of CAP proteins binds to a site centered at -61.5, best characterized at the lac and synthetic CC(-61.5) promoters (Busby and Ebright, 1999). At such promoters, transcription activation involves a protein-protein interaction between activating region 1 of CAP and one of the two RNAP a subunit CTDs and proceeds via a simple recruitment mechanism to stimulate the formation of a closed complex. At Class II CAP-dependent promoters, dimeric CAP binds to a site centered at -41.5, best characterized at the gal PI and the artificial CQ-41.5) promoters. At these promoters, transcription activation involves three independent sets of protein-protein interactions between CAP and RNAP including contacts to the aCTD, the aNTD and c 7 0 4 .2 . CAP stimulates formation of the initial closed RNAP-promoter complex as well as the subsequent isomerization leading to promoter opening. Since Spo0A~P stimulates transcription by facilitating D N A melting after RNAP isomerization, a closer examination of the mechanism underlying Class II CAP-dependent transcription is merited. At Class II CAP-dependent promoters, RNAP is recruited by the interaction between activating region 1 in CAP and an aCTD. Isomerization is stimulated by the additional interactions 70 between CAP activating region 2 and aNTD and between CAP activating region 3 and a 4. A detailed structural model of the CAP-RNAP-promoter complex has been constructed at CC(-41.5) by superimposing the structures of the CAP-aCTD-promoter complex, the -35 element-a4 complex and the RNAP-fork junction D N A complex, refining local D N A helix parameters and modeling downstream D N A according to structural and biophysical data (Lawson et al, 2004). Importantly, a restructuring of the D N A from a sharp kink to a smooth bend within the downstream half on the D N A site for CAP was necessary to permit contacts between CAP and the aNTD and o 7 0 4 . It was suggested that CAP binding restricted promoter topology initially permitting only the interaction of activating region 1 and aCTD. A scenario can be envisioned whereby transient fluctuations in the initial nucleoprotein structure might redistribute the kink 70 over several base pairs, allowing the formation of contacts to aNTD and a 4 , that in turn accelerate promoter melting by more precisely aligning the -10 element with the appropriate regions in RNAP (Lawson et al, 2004). 126 5.4 tel. E. coli A C I is the paradigm for activators that stimulate open complex formation without affecting RNAP binding (Ptashne and Gann, 2002). The A C I NTD is necessary and sufficient for D N A binding and transcription activation and when bound to its operator centered at -42, 70 stimulates transcription by RNAP at A P r m through interactions between A C I and a 4 (Bushman et al, 1989; Hochschild et al, 1983; Kuldell and Hochschild, 1994; L i et al, 1994; Nickels et al, 2002). Structures of Ael-operator (Beamer and Pabo, 1992) and Acl-ovpromoter complexes have been determined (Jain et al, 2004) and provide a framework for understanding the acceleration of the D N A strand melting step. Based on a clever series of genetic experiments in which A C I could stimulate transcription with RNAP bearing a 0 4 fused to the aNTD from a promoter bearing an ectopic -35 element (Dove et al, 2000), it was proposed that A C I could stimulate transcription on the basis of a cooperative interaction between itself and o. The crystal structure of the ternary complex of A C I , o 7 0 and the A . P R M promoter suggested that isomerization is the result of a cooperative binding mechanism and allowed a detailed structural model of the Ael-RNAP-promoter complex to be constructed (Jain et al, 2004). In the proposed model, as in the CAP-RNAP-promoter model, the interactions between A C I and a that had been defined genetically could only be accommodated by altering the D N A topology immediately upstream of the -35 element. In support of this idea, subtle changes in DNase I digestion patterns at a number of promoters have been observed between -40 and -35 during the closed to open complex transition (Craig et al, 1998; Kovacic, 1987; L i and McClure, 1998; Ross et al, 2003). Thus, the fact that that A C I stimulates isomerization rather than the initial binding of RNAP is a consequence of the stage at which the two proteins are appropriately positioned to interact. During closed complex formation, RNAP and A C I are closely apposed but do not touch. During, or as a consequence of open complex formation, the surfaces of a and A C I are brought together and may interact. The proposed mechanisms by which CAP and A C I stimulate open complex formation are virtually identical. In the case of CAP at CQ-41.5) and A C I at A P r m , a relatively subtle restructuring of D N A topology by the activators stimulates open complex formation. At the 127 spoIIG promoter where promoter elements are dramatically mis-aligned, the effect of SpoOA~P on D N A topology around RNAP must be similarly dramatic. Thus the proposed re-positioning of RNAP over the -10 element in the presence of SpoOA~P could represent an exaggerated version of the mechanism proposed for CAP and \c\. 6 Transcriptional Regulation: General Mechanisms. This thesis addressed the specific question of how SpoOA~P works at the spoIIG promoter, but one can generalize from the large body of data available to derive major principles underlying the stimulation of transcription. Sequence-specific D N A binding of transcription factors increases their local concentration and effectively limits activity to promoters in the immediate vicinity of the regulators' binding site. The precise position and orientation of these D N A binding sites relative to the consensus promoter elements that direct RNAP binding, arrange the surfaces on transcription factors and RNAP to permit interaction. Protein-protein interactions between regulators and RNAP exert an effect on RNAP function. This interaction may prompt the allosteric modification of RNAP domains. For example, PspF (an NtrC-like phage shock protein), and gp2 (a phage t7-encoded inhibitor) appear to stimulate and oppose conformational changes respectively in the P' subunit jaw domain required for open complex formation by R N A P a 5 4 (Wigneshweraraj et al., 2004). Other protein-protein interactions between regulators and RNA polymerase stabilize particular intermediates during transcription initiation. This appears to be how CAP, A C I and SpoOA~P work. The effect such stabilizing protein-protein contacts have on transcription is context-dependent. For example, identical contacts between the B. subtilis bacteriophage protein p4 and aCTD activate transcription from A3 by stimulating closed complex formation and repress transcription from A2c by over-stabilizing RNAP and preventing promoter clearance (Mencia et al, 1996; Monsalve et al, 1996a; Monsalve et al, 1996b). Alternatively, wild type A C I activates transcription at A P R M with R N A P a 7 0 bearing a single amino acid substitution, by enhancing closed complex formation rather than by stimulating the rate of open complex formation (Li et al, 1997). In essence, regulators act as catalysts and can be thought of from a thermodynamic perspective as altering the activation energy required to drive transitions 128 between intermediates (Roy et al, 1998). 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