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On the role of ppGpp and DksA mediated control of σ54‐dependent
transcription
Lisandro Bernardo
Department of Molecular Biology Umeå University Umeå , Sweden
2006
Copyright © 2006 by Lisandro Bernardo ISBN 91‐7264‐217‐3
Printed by Arkitektkopia AB, Umeå, Sweden
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Science is facts; just as houses are made of stone, so is science made of facts; but a pile of stones is not a house, and a collection of facts is not necessarily science.
Jules Henri Poincaré (1854‐1912) French mathematician.
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Table of Contents
ABSTRACT …………………………………….....................................…….....................6
PAPERS IN THIS THESIS ………………………………...…........................................7
INTRODUCTION ………………………………………………......................................8
1.1. Overview of specific and global regulatory signals of bacterial gene expression …..........8 1.2. The engine of the transcription cycle ………………………………………...……..................9 1.2.1. The promoter specificity subunit ……………………………………….…..…….........10 1.2.2. Families of σ‐factors …………………………………………………….……….............12 1.2.3. Structure to function relationships of σ‐factors ………………………...…….........…14 1.2.4. Transcription initiation step‐by‐step ……………………………………..……............16 1.2.5. σ54‐activators ‐ enhancer binding proteins (EBPs) ……………………………............18 1.3. Modulation of transcription initiation by nucleoid‐associated proteins …………............20 1.4. The alarmone ppGpp – the effector of the stringent response ………………...……..........21 1.4.1. ppGpp and transcription ………………………………..……………………................23 1.4.2. DksA ‐ an enhancer of ppGpp activity ……………………………………..…….........25 1.5. Global regulation of hydrocarbon catabolism ……………………………………...….........27 1.6. The experimental system derived from Pseudomonas sp. strain CF600 ……………...........28 1.6.1. Subservience of the DmpR/Po system to global regulatory input ………..…...........30
AIMS ……………………………………………………………….............…..................32
RESULTS AND DISCUSSION ……………………………………......…...................33
1. Do global regulatory systems found to impact a closely related system also impact DmpR/Po? ……………………...………………………………………......................33 1.1. Is FtsH required for σ54‐Po transcription? ……………………….........…………............33 1.2. Is the DmpR/Po system under the control of PtsN‐mediated glucose repression? ...............................................................................................................34 1.3. Influence of ppGpp on DmpR/Po and XylR/Pu circuits in E. coli and P. putida ...........................................................................................................................35
2. Do ppGpp and DksA have direct and/or indirect effects on any of the processes of specific regulation? ……………………………………..…………..................36 2.1. Are both ppGpp and DksA required for efficient σ54‐dependent transcription in vivo? .............................................................................................................38 2.2. What’s true of E. coli is (nearly true) of P. putida – an adaptive twist ……...................39 2.3. Do ppGpp and DksA have any direct effects on σ54‐dependent transcription in vitro?.............................................................................................................41 2.4. Can the requirement for ppGpp and DksA be explained by their effects on the levels of IHF?..................................................................................................43 2.5. Can the requirement for ppGpp and DksA be explained by their effects on the levels of DmpR or σ54?...…………………………………………................44
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3. Do ppGpp and DksA regulatory effects through σ70‐transcription explain the impact of these regulatory molecules on σ54‐transcription? …………………....45 3.1. Supressor mutations reveal a link between ppGpp and σ54‐dependent transcription .................................................................................................45 3.2. σ70 mutants are defective in competition with σ54 for core RNAP ……………............46 3.3. σ‐factor competition limits σ54‐dependent transcription in vivo ……………................48 3.4. Mutations within β and β´ reveal a dual bypass mechanism that directly links σ54‐transcription to properties associated with down‐regulation of stringent σ70‐promoters …………......................………...................50 3.5. ppGpp and DksA mediated regulation of σ54‐transcription – the current model .................................................................................53
CONCLUDING REMARKS ………………………...........................…………............55
ACKNOWLEDGEMENTS ……………………………………………………….........56
REFERENCES ……………………………………………………………………............58
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ABSTRACT The σ54‐dependent Po promoter drives transcription of an operon that encodes a suite of enzymes for (methyl)phenols catabolism. Transcription from Po is controlled by the sensor‐activator DmpR that binds (methyl)phenol effectors to take up its active form. The σ54‐factor imposes kinetic constraints on transcriptional initiation by the σ54‐RNA polymerase holoenzyme which cannot undergo transition from the closed complex without the aid of the activator. DmpR acts from a distance on promoter‐bound σ54‐holoenzyme, and physical contact between the two players is facilitated by the DNA‐bending protein IHF. The bacterial alarmone ppGpp and DksA directly bind RNA polymerase to have far reaching consequences on global transcriptional capacity in the cell. The work presented in this thesis uses the DmpR‐regulated Po promoter as a framework to dissect how these two regulatory molecules act in vivo to control the functioning of σ54‐dependent transcription. The strategies employed involved development of i) a series of hybrid σ54‐promoters that could be directly compared and in which key DNA elements could be manipulated ii) mutants incapable of synthesizing ppGpp and/or DksA, iii) reconstituted in vitro transcription systems, and iv) genetic selection and purification of mutant RNA polymerases that bypass the need for ppGpp and DksA in vivo. The collective results presented show that the effects of ppGpp and DksA on σ54‐dependent transcription are major, with simultaneous loss of these regulatory molecules essentially abolishing σ54‐transcription in intact cells. However, neither of these regulatory molecules have discernable effects on in vitro reconstituted σ54‐transcription, suggesting an indirect mechanism of control. The major effects of ppGpp and DksA in vivo cannot be accounted for by consequent changes in the levels of DmpR or other specific proteins needed for σ54‐transcription. The data presented here shows i) that the effects of loss of ppGpp and DksA are related to promoter affinity for σ54‐holoenzyme, ii) that σ54 is under significant competition with other σ‐factors in the cell, and iii) that mutants of σ70, and the β‐ and β’‐subunits of RNA polymerase that can bypass the need for ppGpp and DksA in vivo have defects that would favour the formation of σ54‐RNA holoenzyme over that with σ70, and that mimic the effects of ppGpp and DksA for negative regulation of stringent σ70‐promoters. A purely passive model for ppGpp/DksA regulation of σ54‐dependent transcription that functions through their potent negative effects on transcription from powerful σ70‐stringent promoters is presented.
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PAPERS IN THIS THESIS This thesis is based on the following articles and manuscripts that are referred to in the text by their roman numerals I. Sze C.C., Bernardo L.M.D. and Shingler V. (2002). Integration of global regulation of two aromatic‐responsive σ54‐dependent systems: a common phenotype by different mechanisms. J Bacteriol 184, (3) 760‐70. II. Laurie A.D., Bernardo L.M., Sze C.C., Skärfstad E., Szalewska‐Palasz A., Nyström T., and Shingler V. (2003). The role of the alarmone (p)ppGpp in σN competition for core RNA polymerase. J Biol Chem 278, (3) 1494‐503. III. Bernardo L.M.D., Johansson L.U.M., Solera D., Skärfstad E. and Shingler V. (2006). The guanosine tetraphosphate (ppGpp) alarmone, DksA and promoter affinity for RNA polymerase in regulation of σ54‐dependent transcription. Mol. Microbiol. 60, (3) 749‐764. IV. Szalewska‐Palasz A., Bernardo L.M.D., Johansson L.U.M., Skärfstad E., Stec E., and Shingler V. (2006). Dual by‐pass mechanisms of RNA polymerase mutants: implications for ppGpp‐ and DksA‐mediated control of σ54‐dependent transcription. Submitted manuscript. V. *Bernardo L.M.D., *Johansson L.U.M., Solera D., Skärfstad E., Szalewska‐Palasz., A and Shingler V. (2006). The role of the alarmone ppGpp and DksA in regulation of σ54‐dependent transcription in Pseudomonas putida. Manuscript. * The first two authors contributed equally to this work.
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INTRODUCTION 1.1. Overview of specific and global regulatory signals of bacterial gene expression.
In their natural habitats, bacteria are exposed to a wide range of physical, chemical, and nutritional stresses. In order to adapt and counteract these usually adverse conditions, bacteria alter their gene expression. The regulation of gene expression in response to environmental signals involves sensing the stimuli that, either directly or via signal transduction pathways, leads to up‐ or down‐regulation of the levels of different gene products to make appropriate changes in cellular physiology.
One group of environmental signals is involved in reporting on the availability of essential resources such as the presence or absence of trace minerals, carbon‐, nitrogen‐ and phosphate‐sources, electron acceptors, and other metabolites necessary for growth. Other environmental stimuli such as temperature, osmolarity, pH and other factors that evoke a general protective response can have both specific and global regulatory consequences (94). A third class of signals constitute molecules used for detection of the same or other organisms, which is an important process for certain bacteria in sensing population densities as in bioluminescence, or sensing the presence of a suitable niche such as a eukaryotic host.
Many levels of the control of gene expression can be the target of environmental signals. These include 1) regulation at the level of transcription ‐ the process by which DNA information is transcribed into RNA, 2) at the post‐transcriptional level e.g. by RNase‐mediated regulation of RNA stability, or through translational control brought about by the action of small regulatory RNAs, or 3) at the post‐translational level through a number of mechanisms such as covalent modifications of proteins, protein co‐factor requirements, correct folding, processing, or localization to an appropriate compartment of the cell.
A distinct feature of the prokaryotic cell is the multicistronic operon, such as those for metabolic pathways, in which genes that encode proteins involved in a related process are transcribed from a single promoter. This genetic organisation provides a ready means to coordinate expression in response to a signal that controls the activity of a dedicated regulator to provide specific regulatory circuits. The stimuli that the regulator(s) sense(s) are often directly relevant to the function of the operon(s), e.g. for catabolic operons, the signal is often the substrate or intermediates of the
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catabolic pathway. Therefore, these genes are under specific regulation. Specific regulation also occurs in regulons in which cellular functions that involve gene products of multiple operons are differentially but coordinately controlled by one or more regulators. A stimulon, on the other hand, refers to a group of genes or operons that respond to a defined stimulus, without regard to their regulatory organization. All specific gene regulatory circuits can be considered “guests” that are subject to a higher order of regulation involving global regulatory factors. The presence and/or net activity of these global factors that act through promoters are ultimately dictated by metabolic signals that result from adaptation of the bacterium to its current environment (reviewed in 29, reviewed in 121).
Once adapted to the physical conditions of a new environment, bacteria often have to compete with other organisms. Competing effectively might mean being able to competitively scavenge scarce nutrients, e.g. iron, or taking advantage of plentiful ones to achieve higher growth rates. Within these choices, different compounds may be available for use as carbon and energy sources. To optimise energy usage, a bacterium needs to choose the one(s) it can use most cost‐effectively. This thesis focuses on dissection of how global regulation dominates a specific regulatory circuit to affect this choice. Since the mechanism uncovered primarily functions at the level of transcriptional initiation, I will first introduce the transcriptional machinery and the processes it mediates, before introducing other factors involved in global regulation that are pertinent to the experimental system used in this study.
1.2. The engine of the transcription cycle
Bacteria use DNA‐dependent RNA polymerase (RNAP) to transcribe the genetic information from DNA into RNA. As illustrated in Fig. 1, the transcription cycle consists of distinct steps: promoter‐localization, initiation, elongation and termination. The core enzyme, which is the catalytic machine that synthesizes RNA, is a thermodynamically stable complex of five subunits (α2ββʹω, with a molecular mass of ~400 kDa). The core complex exhibits a “crab‐claw” like structure (made up by the β and βʹ subunits) that encompasses the active site of the enzyme that is located at the back‐end of the claw where essential Mg2+ ions are bound (91). The ω‐subunit is involved in assembly of the core enzyme, which starts with the α2 dimer that serves as the scaffold on which β and βʹ are assembled with the aid of ω (50).
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σσσcycle
1. Promoter localization
2. Initiation 3. Elongation 4. Termination
5. Release
6. Reconstitution
mRNAmRNA
aaaα ‐CTD
β/β’α ‐NTDaaa
α ‐CTDβ/β’
α ‐NTD
aaaα ‐CTD
β/β’α ‐NTD
σUP ‐35 ‐10
aaaα ‐CTD
β/β’α ‐NTD
σaaa
α ‐CTDβ/β’
α ‐NTD
σσUP ‐35 ‐10UP ‐35 ‐10
aaaα ‐CTD
β/β’α ‐NTD
UP ‐35 ‐10
aaaα ‐CTD
β/β’α ‐NTD
UP ‐35 ‐10 UP ‐35 ‐10
aaaα ‐CTD
β/β’α ‐NTD
UP ‐35 ‐10UP ‐35 ‐10
aaaα ‐CTD
β/β’α ‐NTD
aaaα ‐CTD β/β’
α ‐NTD
σaaa
α ‐CTD β/β’α ‐NTD
σσ
Fig. 1. The transcription cycle mediated by DNA‐bound RNA polymerase consists of distinct steps: 1. promoter localization, 2. initiation, 3. elongation and 4. termination.
Although core alone can initiate transcription from ends, nicks and
open regions of DNA, it is incapable of selective transcriptional initiation due to its inability to specifically recognize promoter sequences. To initiate transcription from promoters, the core RNAP requires binding to an additional subunit, the sigma‐factor (σ‐factor), that reversibly associates with core to form the σ‐RNAP holoenzyme. At the end of the elongation phase, the formation of a nascent RNA hairpin, or the helicase activity of the termination factor Rho, mark transcription termination, allowing the released core to associate with a σ‐factor and then initiate a new round of transcription from a promoter (114). 1.2.1. The promoter specificity subunit
Transcriptional initiation is a multi‐step process that involves initial binding of holoenzyme to promoter sequences and melting (isomerisation) of the double stranded promoter DNA prior to initiation of RNA catalysis. Within the context of the holoenzyme , the σ‐factor facilitates promoter recognition by binding to specific DNA signatures of the promoter, and participates in transcriptional initiation by aiding opening of the double‐stranded DNA (53, 152). Different σ‐factors exhibit various sequence specificities for DNA signatures of the promoters they control (78). These different classes of promoters often control groups of genes whose products are involved in related and specialized processes (see Table 1). Most bacteria contain multiple σ‐factors, and genome sequencing has shown that the number of σs vary greatly between different bacteria. For
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example, dedicated intracellular pathogens that live within a relatively constant environment frequently encode only one σ‐factor, e.g. Mycoplasma genetalium, while free‐living soil bacteria such as Pseudomonas putida and Streptomyces coelicolor encode, respectively, 24 (83) and 63 σ‐factors (14). In general, organisms with more diverse lifestyles contain more σ‐factors to rapidly adjust their metabolism and developmental programs in response to various environmental changes.
Table 1. Sigma factors and their functions (adapted from 152).
Group Examples Functions Primary σ‐factors σ70 / σD (RpoD)
(Gram‐negative bacteria) σA (Gram‐positive bacteria) MysA (Mycobacteria) HrdB (Streptomyces)
Major σ‐factor in exponential phase, regulates house keeping genes
Non‐essential stationary phase / stress σ‐factors
σS / σ38 (RpoS) (Enterobacteria, Pseudomonas) SigB‐E (Cyanobacteria)
Major σ‐factor during entry into stationary phase and/or exposure to hyper‐osmotic or acid stress. Controls gene expression during circadian responses, C and N limitation and post‐exponential phase.
Flagella σ‐factors σF / σ28 (Enterobacteria) WhiG (Streptomyces) σD (Bacillus subtilis)
Expression of late flagellar components, chemotaxis and/or early sporulation genes
Extracytoplasmic function (ECF) σ‐factors
σE / σ24 (RpoE) (E. coli, Mycobacteria) CarQ (M. xanthus) AlgU (P. aeruginosa) SigV‐Z (B. subtilis)
Expression of genes involved in alginate biosynthesis, iron uptake, antibiotic production, induction of virulence factors, OM proteins or survival of high temperature
Heat shock and related σ‐factors
σH / σ32 (RpoH) (Gram‐negative bacteria) SigB (M. xanthus) SigC (S. aurantiaca) σB (B. subtilis)
Expression of genes during stress, fruiting body formation, myxospores maturation, or late sporulation
Sporulation σ‐factors
σH, σF, σE, σG, σK
(Bacillus, Clostridium) Transcription of genes involved in various stages of sporulation
σ54‐factors σ54 / σN (RpoN / NtrA) (Enterobacteria, Rhizobium, Pseudomonas, Bacillus, Caulobacter, M. xanthus)
Expression of genes involved in N fixation, nitrate utilization, glutamate synthesis, dicarboxylate transport, aromatic degradation, fimbriae and flagellar synthesis, and others
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The innate affinities of the promoter sequences for their cognate holoenzymes create hierarchies between promoters of the same class, while different levels of holoenzymes create hierarchies between different promoter classes. σ‐factors have different affinities for core RNA polymerase (estimated to be in the range of 1 to 10 nM) (52 and references therein). In addition, the levels of many of them are controlled by sophisticated regulatory systems involving transcription, translation and proteolysis. For example, in Escherichia coli, the nucleotide ppGpp acts as a positive regulator of transcription of rpoS that encodes σS, Hfq and HU stimulate translation by affecting the secondary structure of the rpoS mRNA, while phosphorylated RssB modulates σS availability by delivering it to the ClpXP protease (reviewed in 57).
The levels of other σ‐factors are controlled by anti‐σ‐factors, which sequester the cognate σ‐factor so that it is unavailable to associate with core RNA polymerase until condition trigger degradation of the anti‐σ‐factor, e.g. the anti‐σE protein RseA (3). In the case of σ28, the FlgM anti‐σ28 protein inhibits transcription of σ28‐dependent genes whose products are needed late in assembly of flagella. Once the hook‐basal body of the flagellar motor structure is in place, FlgM chaperoned by σ28 exits through this partially completed flagella structure, thus freeing σ28 to form the cognate holoenzyme to transcribe genes needed to complete the structure (4). In E. coli, the levels of the housekeeping σ70 are relatively constant (67); however, the stationary‐phase Rsd protein (65, 66) and the related phage‐encoded regulator, AsiA (120), associate with σ70 to reduce σ70‐transcription in vitro.
These limited examples illustrate some of the elegant strategies that allow bacteria to modulate the composition of σ pools, and consequently transcription, to efficiently respond to changing environmental conditions and sequentially control gene expression to give temporal order to complex events. In contrast to other alternative σ‐factors, the levels of σ54 remain constant under different growth conditions and growth phases in E. coli and P. putida (30, 67), hence alternative explanations are needed for growth‐phase and conditional regulation of promoters recognised by σ54‐RNAP. The mechanisms underlying these responses of σ54‐promoters are discussed in detail in the Results and Discussion section.
1.2.2. Families of σ‐factors
Based on sequence and functional similarity, σ‐factors can be grouped into two families, which have little if any sequence identity between them (Fig. 2). The largest of these is the σ70‐family, named after the
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70 kDa primary σ‐factor encoded by rpoD that is also sometimes referred to as σD (78). The second family is the σ54‐family, named after the 54 kDa nitrogen regulation σ‐factor, which is encoded by rpoN (ntrA) and is thus sometimes referred to as σN (88). In addition to nitrogen metabolism, σ54 is involved in controlling many processes that are responsive to environmental signals, including the utilization and transport of alternative carbon sources, chemotaxis, and alginate biosynthesis (reviewed in 141).
Activation ofPromoter melting
Region IIAcidic
Region IIIconserved
Region ILeu-gln rich
Core binding DNACross-link
RpoNbox
HTHand -12
recognition
σ54σ54
NCR
σ1.1
σNCR
σ3 σ4σ2σ 3.2 loop
Inhibits DNA bindingCore binding
DNA melting
-10 recognition
Extended -10 recognition
HTH box and
-35 recognition
σ70σ70
1.1 1.2 2.1 2.2 2.3 3.0 3.12.4 3.2 4.1 4.21.1 1.2 2.1 2.2 2.3 3.0 3.12.4 3.2 4.1 4.2
Fig. 2. Schematic illustration of the σ70‐ and σ54‐subunits of RNA polymerase. NCR: non‐conserved region (adapted from 90, 147, respectively).
Both σ54 and σ70‐like proteins have regions involved in DNA‐
binding and interaction with core RNA polymerase (Fig. 2). Unlike σ70, σ54 is able to bind to certain promoters in the absence of core polymerase, although σ54‐holoenzyme binds tighter than σ54 alone (reviewed in 88). The σ54‐factor programs RNAP to recognise the unusual ‐24, ‐12 class of promoters, consensus TGGCAC‐N5‐TTGCa/t with the almost invariant GG and GC residues underlined (12). This sequence is very different from those
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of σ70‐promoters, consensus TTGACA‐N17‐TATAAT, which are located at ‐35 and ‐10 with respect to the transcription start site.
The σ70 family has been broadly divided into four phylogenetic groups on the basis of gene structure and function (reviewed in 54). Group 1 consists of the essential primary σ70/D (78) that controls transcription of housekeeping genes. This group has two distinctive features: a) region 1.1 which auto‐inhibits promoter recognition by free σ‐factor (90 and references therein) and b) a non‐conserved region (NCR, Fig. 2) that is present only in some housekeeping σ‐factors (reviewed in 54). Group 2 proteins are closely related to the primary σ‐factors and direct the RNA polymerase to a large number of genes upon conditions of growth arrest, starvation and stress (e.g. the stationary phase and stress factor σ38/S), but are dispensable for growth (57). Members of group 3 lack region 1.1 and NCR, and have roles in sporulation (e.g. σ32/H, σ28/F and σK), in flagella biosynthesis (e.g. σ28/F) and in the heat‐shock response (σ32/H) (reviewed in 54). Finally, group 4 accommodates the extracytoplasmic function (ECF) σ‐factors that lack region 1.1, NCR and regions 3.0 to 3.1. These σ‐factors respond to signals from the extracytoplasmic environment, such as the presence of misfolded proteins in the periplasmic space and cell envelope stress (e.g. σ24/E) (reviewed in 54).
1.2.3. Structure to function relationships of σ‐factors
Based on the crystal structure of Thermus aquaticus σA (equivalent to σ70 of E. coli), the regions of homology of σ70‐like proteins encode functional domains that form extensive contacts with core RNAP (21). As illustrated in Fig. 3B, these subunits are spread across the upstream face of the core RNA polymerase crab‐claw, and consist of globular regions (σ2, σ3, and σ4) that are connected by flexible linkers. Region 1.1 functions as an auto‐inhibitory domain that prevents free σ70 from binding promoter DNA, but upon interaction with the core enzyme, this inhibitory effect is relieved (87). Regions of the globular domains that are involved in DNA‐binding are all solvent‐exposed, while the σ‐3.2 loop protrudes into the active site cleft (see Fig. 3B). The σ2 domain forms one of the primary interfaces with RNAP, and is involved in both recognition of the –10 hexamer DNA sequence of promoters and in DNA melting (21 and references therein). The σ3 domain binds extended ‐10 DNA sequences that are important for the functioning of promoters lacking –35 sequences (54), while the σ4
domain recognizes ‐35 DNA hexamers of σ70‐promoters. The σ4 domain contacts the β‐flap region that lies right over the RNA exit channel (Fig. 3B).
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The σ4 domain is also the target for interaction with transcriptional regulators at some promoters, and in some cases is simultaneously contacted along with one of the α‐carboxy‐terminal domains (α‐CTDs) of RNAP (51).
αI
αII
β β’
Mg2+
Active site
ω
σ1.1
β
β’
σ4
αII
RNA exit channel
template strand non-template strand
-5
-15
-25
-35
σ3σ3
σ2
downstreamDNA
900
A) B)
αI
Fig. 3. A) Schematic illustration of core RNA polymerase and B) the σ70‐RNA polymerase holoenzyme at the first step of open complex formation (see text). Active Mg2+ ions (circles). The path of the RNA to be synthesized is indicated by a dashed line to emphasise the local conflict between the σ‐3.2 loop and the nascent RNA (adapted from 62).
Until now, (to the best of my knowledge), no high‐resolution
structure of σ54‐proteins has been solved. Nevertheless, alignment of the highly conserved amino acid sequence of σ54 from different bacteria has identified three distinct regions (Fig. 2). The amino‐terminal Region I is a regulatory bifunctional domain that inhibits DNA isomerisation by the holoenzyme in the absence of activation (23, 73, 130, 144); but aids DNA isomerisation and stable open complex formation upon activation (27, 116, 130). Region II is an acidic‐residue‐trimer repeat linker connecting Region I and Region III. Deletion studies of σ54 suggested a role for region II in DNA‐binding and in stabilising the holoenzyme (22), as well as effects on open complex formation in the presence of an activator protein (126). The carboxy‐terminal Region III contains sequences required for core RNAP binding (49, 135) and determinants for DNA interactions, including the helix‐turn‐helix motif and a patch that UV‐crosslinks to promoter DNA (33, 49, 88).
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1.2.4. Transcription initiation step‐by‐step The multiple steps of transcription initiation involve major
sequential structural changes within both the DNA and the RNA polymerase. Extensive genetic, biochemical and structural studies have greatly helped understanding of these multiple steps in both σ70‐ and σ54‐dependent transcription initiation events, even though not all is fully known. A schematic overview of some of the intermediates in this process is shown in Fig. 4.
BinaryClosed
Complex
RNAPolymerase
+Promoter DNA
BinaryOpen
Complex
NTP
TernaryInitiationComplex
AbortiveInitiation
NTP TernaryElongationComplex
+Promoter
DNAσ
R+P RPC1/RPC2 RPinit RPERPO1/RPO2
BinaryClosed
Complex
RNAPolymerase
+Promoter DNA
BinaryOpen
Complex
NTP
TernaryInitiationComplex
AbortiveInitiation
NTP TernaryElongationComplex
+Promoter
DNAσ
R+P RPC1/RPC2 RPinit RPERPO1/RPO2
Fig. 4. Transcription initiation is a multi‐step process which involves promoter recognition, formation of a closed DNA complex, isomerization of the DNA to form the open complex, nascent RNA initiation in the ternary initiation complex, and finally promoter escape/clearance to form the ternary elongation complex.
Transcription is initiated when a σ‐bound RNA polymerase (R)
recognizes and binds to the double‐stranded promoter DNA sequence to form the so‐called binary closed complex (RPC). Within the RPC1 σ70‐holoenzyme‐promoter complex, σ2 engages the promoter –10 element, σ4 the –35 elements, while the σ‐3.2 loop and σ‐1.1 region protrude into the RNAP active site channel (21). The binding of downstream double‐stranded DNA in the active site displaces the σ‐1.1 region, bringing both the DNA upstream and downstream of the transcriptional start site into contact with the RNA polymerase, forming the binary closed complex RPC2 (40).
Unwinding and separation of the double‐stranded DNA from approximately position –10 to position +2 by the σ2 sub‐region‐2.3 form the transcription bubble, and marks the start of open complex formation, resulting in the binary open complex RPO (see Fig. 3B), that has at least two intermediates (RPO1 and RPO2) (91). Once the transcription bubble extends past the +1 transcription start, the single‐stranded template DNA is directed to the active site, where NTP (nucleotide triphosphate) substrates
16
are provided through the secondary channel and the synthesis of RNA is initiated, resulting in the ternary initiation complex RPinit.
As illustrated in Fig. 3B, the path of the σ‐3.2 loop in the active site is in conflict with the path that the synthesised RNA will take through the RNA exit channel under the β‐flap. Within the RPinit complex, the elongating RNA chain must either displace the σ‐3.2 loop out of its way or dissociate from the complex and be released (probably through the secondary channel), a process that is called abortive initiation. Once the RNA chain elongates to ~12 nucleotides, it is of sufficient size to fill the RNA‐DNA hybrid and upstream RNA exit channel under the β flap (90). The displacement of the σ‐3.2 loop thus ends the potentially abortive stage of initiation. The displacement of the σ‐3.2 loop and exiting of the RNA is also believed to be linked to the initial steps of promoter escape, resulting from destabilization of the σ4‐β flap interaction as RNA exits through the underlying channel. As a consequence of the release of σ4 from the β flap, the interactions between σ4 and the ‐35 element would be destabilized, allowing the RNAP to be released from the promoter and to translocate downstream as it elongates the RNA. Eventually the σ70 is completely released, through sequential breaking of its multiple interactions with core, regenerating core RNAP in its processive elongation mode as ternary elongation complex; RPE (90).
The levels of formation of the non reversible ternary initiation complex are primarily determined by the ease of melting of the DNA strands, the stabilities of the intermediates which are all reversible and the ability of the nascent RNA to out‐compete the σ‐3.2 loop (90 and references therein); and at some promoters can also be greatly influenced by the concentration of the initiating NTPs (117).
In contrast to σ70‐RNAP, σ54‐RNAP forms an unusually stable closed complex around positions −24 (GG‐region) and −12 (GC‐region) from the transcription start site, and is incapable of self‐isomerization to an open complex. Thus, transcription by this holoenzyme always requires activation. The interactions between σ54‐regions I and III with the ‐12 DNA sequence are responsible for preventing spontaneous isomerization of the closed complex to the open promoter complex in the absence of activation (55, 147). Initiation of open complex formation by σ54‐RNAP requires interactions between σ54‐region I with a specialized AAA+‐family activator protein (ATPases associated with diverse cellular activities), which utilizes ATP hydrolysis to drive conformational changes in the σ54‐RNAP closed promoter complex, resulting in the formation of the open promoter
17
complex (20, 27, 48). The initial open complex (RPO1) formed is heparin‐unstable and the activator interaction with σ54‐RNAP further assists in its conversion to a heparin‐stable, NTP binding proficient state (RPO2) (143). A functional interplay between σ54‐region I and the β and the β’ subunits stabilizes the σ54‐RNAP open complex (148 and references therein). Once the open‐promoter complex has formed, the pathway for σ54‐RNAP is much as described for σ70‐RNAP except that abortive initiation appears more efficient, since the abortive products are 2 nucleotides long as compared to 3‐12 nucleotides for σ70‐RNAP (63, 136).
1.2.5. σ54‐activators ‐ enhancer binding proteins (EBPs)
Classical regulators of transcription initiation act by binding to DNA regions in or proximal to a promoter to affect RNAP recruitment to the promoter and/or to enhance rate‐limiting steps of the transcriptional initiation pathway (reviewed in 19). However, as outlined above, σ54‐RNAP forms stable closed complexes, and the cognate regulators of this polymerase are thought to act solely by acting on pre‐bound σ54‐RNAP locked in the closed complex form.
V4R
Major effectorspecificity sub-region
GAFTGA HTH
Sensory/ Regulatorydomain
DNA-binding domain
AAA+ σ54-activation domain
1 211 234 472 558 563
Linker
V4R
Major effectorspecificity sub-region
GAFTGA HTH
Sensory/ Regulatorydomain
DNA-binding domain
AAA+ σ54-activation domain
1 211 234 472 558 563
Linker
Fig. 5. Domain structure of a σ54‐activator, e.g. DmpR, illustrating the locations of the hydrocarbon‐binding V4R signature motif, the σ54‐RNAP interacting GAFTGA motif, and the helix‐turn‐helix DNA binding motif (adapted from 128).
The activity of σ54‐activators is controlled in response to a wide
variety of physiological and environmental signals, affecting cellular responses ranging from nitrogen assimilation and utilization (e.g. NifA and NtrC), utilization of alternative carbon sources (e.g. DmpR and XylR), expression of specific transport systems (e.g. DctD), to production of virulence determinants and extracellular structures (reviewed in 128). This family of proteins usually consists of three domains involved in signal reception, transcriptional activation, and DNA binding (Fig. 5). The amino‐
18
terminal signal reception domain is a regulatory domain that represses the activity of these proteins until they receive an activating signal in the form of a small ligand, phosphorylation, or by an interacting protein (reviewed in 122). The central catalytic AAA+ domain is required, and is often sufficient, for activating transcription from σ54‐promoters in vivo and in vitro (15, 71, 108, 149). The carboxy‐terminal HTH domain mediates DNA‐binding to sites located unusually far upstream from the promoters they control that are referred to as UASs (upstream activating sequences). Since σ54‐activators can frequently still mediate some level of transcription when their binding sites are moved even further from the promoter, these proteins are also referred to as enhancer‐binding proteins (EBPs) by analogy to transcriptional regulators of eukaryotes (106, 154).
UAS
(EBP)2
(EBP)6
-24 -12 +1
σ54
A
C
B
ATP
ADP+Pi
IHF+1
σ54
+1
σ54
Core
Core
Core
Fig. 6. Illustration of activation of transcription from σ54‐dependent promoters by EBPs. (A) σ54, within the context of the holoenzyme, binds to specific promoter sequences at positions ‐24/‐12 relative to the transcriptional start, forming a closed σ54‐RNAP‐promoter complex. EBPs bind to UASs upstream of the promoter sequence. (B) The EBP hexamer contacts the σ54‐RNAP‐promoter complex via DNA looping, often assisted by IHF. (C) ATP hydrolysis by the AAA+ domain of the EBP results in a σ54‐RNAP‐promoter open complex with melted DNA (adapted from 119).
19
EBPs are usually dimeric in their inactive state and the formation of active higher‐order oligomers (usually hexamers) is required to stimulate ATPase activity and to allow interaction with σ54‐RNAP (e.g. 112, 149). EBPs bound to UASs that are located at ‐80 to ‐150 bp from the transcriptional start site contact the promoter bound σ54‐RNAP via DNA looping (see Fig. 6) (129, 145). In many cases, this event is assisted by the DNA bending protein IHF (Integration Host Factor) (61). Upon contact, conformational changes induced by the EBP as it hydrolyses ATP lead to the disruption of the repressive interaction of σ54‐region I on the core polymerase. This step involves the GAFTGA motif of the central domain, which is the hallmark of this family of proteins. In particular, the conserved threonine of GAFTGA that resides on a mobile loop is suggested to play a crucial role in the energy transfer process derived from ATP hydrolysis to restructure the σ54‐RNAP promoter DNA complex into a transcriptionally competent open complex (Fig. 6C) (17, 112).
1.3. Modulation of transcription initiation by nucleoid‐associated proteins
Bacterial chromosomes far exceed the length of the cell and are compressed thousands of fold in the nucleoid structure that serves as the scaffold within which transcription, replication and recombination take place. The highly compacted bacterial chromosome is ordered by supercoiling, RNA, and nucleoid‐associated proteins that represent the prokaryotic equivalent to histones. Negative supercoiling, due to the negative charges on the phosphates of DNA, can facilitate both DNA folding and compaction and also the unwinding of DNA, which is required for the initiation of transcription as well as DNA replication and recombination. In E. coli, major nucleoid‐associated proteins involved in DNA compaction include FIS (Factor for Inversion Stimulation), IHF, H‐NS (heat‐stable [or histone‐like] nucleoid‐structuring protein) and HU (histone‐like nucleoid‐structuring proteins) (reviewed in 139).
The levels of nucleoid‐associated proteins vary according to growth phase of the cell, e.g. IHF increases up to ~7‐fold upon transition from exponential to stationary phase in E. coli and P. putida (93, 140), while the level of FIS is high during exponential phase but drops to almost nil at stationary phase (10). In particular, the levels of HU, H‐NS and IHF all increase before cell size reduction, probably to better compact the nucleoid to fit the smaller‐sized cells of stationary phase (10).
20
In addition to their roles in compacting chromosomal DNA, nucleoid‐associated proteins can also function as transcription factors through their influence on DNA topology in and round regulatory DNA sequences (118, 151). For example, upon binding, FIS bends DNA between 50° and 90° (102), whilst IHF induces a 160° bend (113). The consequences of changes in DNA topology caused by nucleoid proteins at promoters are varied and include i) enhanced or reduced binding of RNA polymerase and/or regulator(s), ii) establishment or disruption of interactions between RNA polymerase and regulator(s), iii) enhanced or reduced unwinding of DNA duplex (affecting formation of open complexes), and/or iv) nucleation and formation of higher order structure leading to activation or repression of transcription (reviewed in 38).
IHF plays a role at promoters of different types, but is particularly important for σ54‐dependent promoters in facilitating contact between the distally bound EBP and the σ54‐RNAP (see Fig. 6). IHF also has some additional roles at σ54‐dependent promoters such as i) restricting the transcriptional activation specificity to a single regulator bound to specific UASs (35, 45, 109), ii) helping to recruit σ54‐RNAP to the promoter (16), iii) helping to engage regulators to UASs (70), and iv) promoting and/or stabilizing the σ54‐RNAP open complex (131).
One of the most extensively characterized regulatory roles of H‐NS and FIS is their involvement in transcriptional repression (H‐NS) and activation (FIS) of the seven rRNA operons of Escherichia coli that are driven by promoters that are amongst the most powerful known in prokaryotes (2, 118). Ribosomal genes are highly transcribed during the exponential growth phase when nutrients are abundant and large numbers of ribosomes are needed for protein synthesis. When cells reach stationary phase, and the demand for protein synthesis lessens, the decreased levels of the activator protein FIS and increased levels of the repressor protein H‐NS are involved in the down‐regulation of transcription of ribosomal genes (reviewed in 105). However, as detailed in the following sections, the nucleotide ppGpp and its co‐factor DksA, which directly bind and modulate RNAP function, are the most potent factors involved in down‐regulation of these powerful promoters.
1.4. The alarmone ppGpp – the effector of the stringent response
The stringent response occurs when the supply of charged tRNA cannot keep up with the demands for protein synthesis and is elicited under conditions of amino acid starvation, carbon‐, nitrogen‐ and
21
phosphate‐limitation and common physical‐chemical stresses that cause decreased growth rates and/or growth arrest (32). The effector molecule of the stringent response is the nucleotide guanosine tetraphosphate, ppGpp. Since ppGpp production heralds metabolic stress, it serves to raise the alarm for adaptation for difficult times that lie ahead.
In E. coli, this simple‐structured signalling molecule has far reaching global regulatory affects on gene expression (Fig. 7) through its influence on the levels of global regulators such as IHF, σS and σH (77). Regulatory effects of ppGpp on cellular functions have also been reported in many other bacteria for example in sporulation of Bacillus subtilis (46), antibiotic production by Streptomyces spp. (142) and alginate and polyphosphate production by Pseudomonas aeruginosa (74). In addition, the existence of a bacteria‐type stringent response has been recently identified in plants, where increased levels of ppGpp occur in chloroplasts in response to stresses such as wounding, heat shock, high salinity and jasmonic acid (134).
GTP+ATP
Amino Acid Starvation
RelAStress and Starvation
SpoTAmino Acid Starvation
RelAAmino Acid Starvation
RelAStress and Starvation
SpoTStress and Starvation
SpoT
pppGpp
GDP+PPippGpp
pppGpp
GDP+PPippGpp
Glycolysisgenes
Stasis survivalgenes
Oxidative stresssurvival genes
Osmotic stress survival genes
StableRNA
Ribosomalproteins &Elongation
factor
DNA replication
rpoSFatty acids,Lipids &Cell wall
UniversalStress
proteins
Amino acidbiosynthesisProteolysis
StableRNA
Ribosomalproteins &Elongation
factor
DNA replication
rpoSFatty acids,Lipids &Cell wall
UniversalStress
proteins
Amino acidbiosynthesisProteolysis
σS
aaaα -CTD β/β’
α - NTD
σaaa
α -CTD β/β’α - NTD
σσ
Fig. 7. ppGpp is produced from GTP and ATP by 2 routes in response to starvation and stress. It targets RNAP and redirects transcription from growth‐related genes to genes involved in stress resistance and starvation survival. SpoT also hydrolyzes ppGpp (adapted from 80). ‐ indicates down‐regulation, + indicates up‐regulation.
22
In E. coli and P. putida, the accumulation of ppGpp is mediated by the RelA‐ppGpp synthetase I and the SpoT‐ppGpp synthetase II dependent pathways. A strain lacking both RelA and SpoT is completely unable to produce ppGpp and is referred to as a ppGpp0 strain, while the response to starvation of such a strain is called a relaxed response. The specific feature of the relaxed response is a continued accumulation of rRNA during starvation (32). E. coli cells lacking ppGpp are also characterized by being polyauxotrophs (153). The production of ppGpp in response to nutrient limitation and other stresses leads to coordinated changes in gene expression. In general, genes involved in cell proliferation and growth are negatively regulated by ppGpp, whereas genes implicated in maintenance and stress survival are positively regulated by the alarmone; see Fig. 7 (reviewed by 98).
The RelA protein is activated to catalyze the synthesis of ppGpp from ATP and GTP during amino acid starvation or when cells enter stationary phase. Initially pppGpp is produced, which is then rapidly converted by guanosine pentaphosphate hydrolase to ppGpp (75). RelA detects uncharged tRNA stalled on the ribosomes and as a result activates its ppGpp synthesis while hopping to other stalled ribosomes (146). Less is known about how SpoT senses starvation conditions. SpoT is a dual‐functional protein and is also responsible for hydrolyzing ppGpp (32). Most bacteria only contain a single protein that, like SpoT, has two catalytic sites in its amino terminal region that are involved in its dual activity (60, 89). Allosteric changes through its carboxy‐terminal end are proposed to control switching between its two opposing activities. SpoT is primarily responsible for the accumulation of ppGpp in response to most stresses and nutrient limitations other than amino acid starvation (92). Very recently it has been suggested that interaction with a central co‐factor in fatty acid synthesis may be the mediator for controlling the opposing activities of SpoT (13).
1.4.1. ppGpp and transcription
In contrast to classical DNA‐binding transcriptional regulators, ppGpp binds directly to RNAP to modulate its activity. Mutations affecting ppGpp‐mediated control of transcription have been identified in the β (encoded by rpoB), β’ (encoded by rpoC) and σ70 (encoded by rpoD) subunits (58), and crosslinking of ppGpp‐analogues to RNAP has shown that ppGpp interacts with β and β’ subunits (34). These studies have been complemented by the recent co‐crystallisation of RNAP in complex with
23
ppGpp, in which ppGpp was found to bind in either of two orientations adjacent to the active site (8). The authors suggest that ppGpp binding might affect interactions of NTPs with RNAP by altering the configuration of the active site, and that ppGpp might interact with the non‐template DNA strand of the transcription bubble. However, supporting biochemical data for the involvement of residues proposed to bind ppGpp, and hence the significance of the structure found, is still awaited.
A)
Fig. 8. Thermodynamic representation of promoter activation versus inhibition by ppGpp/DksA (adapted from 104). Free energy diagrams for an amino acid promoter (A) and an rRNA promoter (B) showing the progression of transcription initiation. The diagram illustrates two steps, RP1 and RP2, on the pathway to open complex formation with a single transition state ([RP]‡). A two‐step mechanism is depicted for simplicity, but RP1 and RP2 could refer to any two intermediates on the pathway to open complex formation, and ppGpp/DksA conceivably could affect any transition state between the two intermediates. ΔG1 and ΔG2 represent the changes in free energy required to transition from RP1 to [RP]‡ and [RP]‡ to RP2, respectively. The dashed line represents the free energy of the transition state in the presence of ppGpp/DksA. ΔG1D and ΔG2D represent the changes in free energy in the presence of ppGpp/DksA. ppGpp/DksA are proposed to decrease the free energy of [RP]‡, allowing for a more rapid conversion from RP1 to RP2 and from RP2 to RP1. The ground state of both RP1 and RP2 at a particular promoter would dictate whether ppGpp/DksA would facilitate activation as in the case of A) or inhibition of the promoter as in B).
Despite the availability of the co‐crystal structure and the proposed
interaction of ppGpp with DksA, which is described in the next section, it is still unclear how ppGpp mechanistically alters the function of RNA polymerase. Nevertheless, ppGpp has been found to directly act on many steps of transcriptional initiation from different σ70‐promoters in vitro, including: 1) The rate of ternary initiation complex formation through competition between ppGpp and NTPs at the active centre of RNAP (8, 69),
ΔG
RP1
RP2
[RP]‡
ΔG1
ΔG2
ΔG1DΔG2D
RP1
RP2
[RP]‡
ΔG1
ΔG2
ΔG1DΔG2D
ΔG
RP1
RP2
[RP]‡
ΔG1ΔG2
ΔG1DΔG2D
Progression of Transcription Initiation
B)
RP1
RP2
[RP]‡
ΔG1ΔG1ΔG2
ΔG1DΔG2D
Progression of Transcription Initiation
24
which would particularly effect promoters that are sensitive to initiating NTP levels; 2) destabilisation and reduced life‐time of open RNAP‐promoter complexes. This is of particular importance for negative regulation of promoters which have this as a rate limiting step, for example those of rRNA promoters which are intrinsically unstable and short‐lived (11, 103, 156); 3) accelerated transition to the open complex, which is proposed to underlie ppGpp‐stimulation of transcription from promoters controlling certain amino acid biosynthetic and transport pathways (104 and references therein); and 4) negative effects on promoter clearance, which has been proposed to underlie down‐regulation of transcription from the λPR promoter by ppGpp (110). In addition, ppGpp has also been observed to increase pausing during transcriptional elongation (69). The outcome of the effects of ppGpp and DksA at different promoters can be explained by changes on the kinetics of the pathway to open complex formation, depending on the relative stabilities of transition state intermediate(s) (Fig. 8) (104 and references therein).
In addition to acting directly on transcription from σ70‐promoters, ppGpp also acts as a positive regulator of gene expression from some promoters recognised by alternative σ‐factors, e.g. σS (68, 76), σH (68), σ54 (26, 133), and σE (37). In these cases, ppGpp is proposed to primarily mediate its effect through modulating the outcome of competition of the σ‐factors for core RNAP to co‐ordinate responses to nutritional and other stresses (68 and papers II‐IV). This model in relation to σ54‐transcription is the primary focus of this thesis and is developed and discussed further in the Results and Discussion section.
1.4.2. DksA ‐ an enhancer of ppGpp activity
The 151‐amino‐acid long DksA protein was initially isolated in Escherichia coli as a multicopy suppressor of temperature‐sensitive growth and filamentation phenotypes of dnaKJ mutants (72). Deletion of dksA results in pleiotropic effects in the cell such as defects of σS expression (59), cell division, amino acid biosynthesis, quorum sensing, phage sensitivity and virulence in a number of organisms (104 and references therein). Many, but not all, of these effects are probably explained by the identification of DksA as a co‐factor that binds RNAP and amplifies both the positive and negative effects of ppGpp on σ70‐transcription (103, 104 and references therein, 107). However, DksA also mediates ppGpp‐independent effects on transcription (103). The phenotypes of E. coli DksA null and ppGpp0 strains do not completely overlap. For example, both strains exhibit somewhat
25
different amino acid requirements and the promoter sequence requirements for the two regulatory components have been reported to differ (59). Thus, it is possible that DksA and ppGpp in some instances act alone and in other cases act in concert.
Structural studies have shown that DksA is a structural, but not sequence, homologue of Gre factors that are involved in reactivating stalled RNAP by stimulating the intrinsic endonucleolytic transcript cleavage activity of RNAP (107 and references therein). The acidic residues at the tip of the coiled‐coil of E. coli GreA and GreB are positioned close to the active site by protrusion of the coiled‐coil through the secondary channel, see Fig. 9, where they are proposed to help coordinate one of the two catalytic Mg2+ ions at the active centre of RNA polymerase (107 and references therein). Based on the structural similarity of DksA to Gre factors and computer modelling, Perederina and co‐workers propose that the coiled‐coil of DksA also protrudes through the secondary channel of RNAP and that the two aspartate residues at the tip of the DksA coiled‐coil interact with a Mg2+ ion coordinated by ppGpp to directly facilitate and/or stabilize the ppGpp‐RNAP complex, thus providing a structural basis for the synergistic effects of ppGpp and DksA (107).
Mg2+
DownstreamDNA
UpstreamDNA
DksA/GreA/GreB
SecondaryChannel
TemplateDNA
NTPs and ppGppβ
flap
RNA
Fig. 9. Schematic illustration of transcribing RNA polymerase. The secondary channel (represented in white) is the site of entrance for the NTPs and ppGpp and also has to accommodate regulatory proteins such as DksA, GreA and GreB that access the active site to directly modulate the activity of RNAP. β subunit: dark grey and β’ subunit: light grey (adapted from 97).
26
The existence of multiple secondary channel binding proteins such as GreA, GreB and DksA of E. coli, may suggest competition for this site. This competition would most likely be governed not only by the relative concentrations of the regulators, but also by their relative binding affinities for the secondary channel. All these levels of regulation add complexity that can contribute to the diversity of responses to nutritional and environmental signals that have evolved for regulating different promoters.
1.5. Global regulation of hydrocarbon catabolism
Soil micro‐organisms such as Pseudomonas, Alcaligenes, Burkholderia and Acinetobacter are capable of degrading aliphatic (e.g. alkanes, styrene) and aromatic hydrocarbons (e.g. phenols and toluenes). Pseudomonas genomes (> 6 million bp) contain the highest proportion of regulatory genes observed, and a high proportion of genes dedicated to the catabolism, transport and efflux of organic compounds (95). The process of hydrocarbon biodegradation requires sets of specialized enzymes that catalyze sequential biochemical reactions to convert these compounds to central metabolites. Operons encoding aromatic catabolic enzymes are often found on plasmids, or, if located on chromosomes, are flanked by transposons, thus ensuring a degree of horizontal transferability (150). The aerobic catabolism of various classes of hydrocarbons by pseudomonads has been extensively studied due to their environmental bioremediation potential.
The ability to catabolise aromatics is nonessential, serving only to confer an advantage to the host bacterium under conditions where aromatic compounds are readily available. However, aromatic catabolic pathways, like other catabolic processes, have to function efficiently within the context of the host and the signals presented by complex environments. Thus, tight regulation is required in order to avoid detrimental energy fluxes, with these auxiliary metabolic pathways only becoming truly favourable if their usefulness does not compromise the host’s fitness and survival. Therefore, the functioning of aromatic catabolic pathways within the network of host cellular processes requires both specific and global regulation (reviewed in 121).
27
Table 2. Hydrocarbon catabolic systems subject to global regulation.
Pathway/operon and phenotype Reference
Down‐regulation of pentachlorophenol metabolism by glutamate, glucose and cellobiose.
(137) (138)
Inhibition of benzene consumption of P. putida ML2 by succinate. (84) Down‐regulation of activity of PnahG promoter involved in salicylate/naphthalene degradation of P. fluorescens H K44 by rich media, glucose and toluene.
(56)
Inhibition of ethyl benzene catabolism of P. fluorescens CA4 by glutamate but not glucose or citrate.
(36)
Down‐regulation of TOL pathway of pWW0 of P. putida mt2 by rich media, rapid growth, and some carbohydrates e.g. glucose.
(82)(43)(44) (42)(30)
Inhibition of degradation of styrene in P. putida CA3 by glutamate and citrate, but not glucose.
(99)
Down‐regulation of (methyl)phenol degradation dmp pathway of Pseudomonas sp. CF600 by media that support rapid growth.
(132)
Down‐regulation of chlorobenzoate degradation clc pathway of P. putida AC27 by fumarate.
(85) (86)
Inhibition of benzoate consumption of Ralstonia eutrophus by succinate. (6)(5) Down‐regulation of n‐alkane degradation alk pathway of P. oleovorans by rich media and rapid growth.
(155) (127)
Down‐regulation of styrene catabolism in P. fluorescens ST by glucose, acetate and glutamate, and to a lesser extent by succinate and lactate.
(115)
The phenomena of global regulation overriding specific regulation
of hydrocarbon catabolism have been reported in many systems, and some examples are listed in Table 2. The mechanisms of how specific systems are integrated and made subservient to global regulatory input are many and varied, and can involve competition between pathway substrates and central metabolites for binding sites of a regulator (e.g. ClcR), through the action of components of an alternative PTS system (e.g. at the Pu promoter for toluene catabolism of pWW0), to switching of the use of different σ‐factors (e.g. Pm promoter of pWW0) (reviewed in 121). The experimental system used in this study, that of the dmp‐operon of pVI150 from Pseudomonas sp. CF600 is also subservient to global regulation (132).
1.6. The experimental system derived from Pseudomonas sp. strain CF600
Interest in the regulatory and metabolic processes of the soil bacterium Pseudomonas sp. strain CF600 mainly derives from its ability to degrade phenols and substituted phenols. This organism was first isolated by selection in media containing 3, 4‐dimethylphenol as the sole carbon
28
source, and subsequently found to similarly grow on phenol, 2‐, 3‐ and 4‐methylphenols. These compounds are more usually known for their toxicity than roles as growth substrates. The genetic setup that allows such unusual growth is conferred by the dmp‐system, which includes the dmp‐operon and the divergently transcribed regulatory gene dmpR that are carried on pVI150, an Inc‐P2 megaplasmid (124). The dmp‐operon is composed of fifteen genes transcribed from the Po promoter. These genes encode the nine enzymes required for sequential conversion of (methyl)phenols to pyruvate and acetyl CoA (Fig. 10), which then feed into central metabolism (125).
Transcription from the Po promoter of the dmp‐operon is dependent on σ54‐RNAP (123), and shows all the typical hallmarks of a σ54‐dependent promoter, including a functional IHF binding site located between the ‐24, ‐12 promoter and distally located UAS sequences that comprise the binding sites for DmpR (131). Likewise, the DmpR regulator that controls the output from Po is a mechano‐transcriptional regulator of the AAA+ family (101, 149). DmpR is a sensory transcriptional regulator that directly binds phenolic pathway substrates (and structurally related compounds) via its amino‐terminal domain to unlock its transcriptional promoting activity and take up its active multimeric form (100, 149). As such, the specific regulatory circuit provided by DmpR and Po serves to ensure that the metabolically expensive Dmp‐enzymes are only produced when pathway substrates are present.
Fig. 10. Genetic organization of the dmp‐system and the (methyl)phenol catabolic pathway catalyzed by the enzymes it encodes. Phenolic substrates are converted to catechols by the multi‐component phenol hydroxylase Dmp(K)LMNOP. Subsequently, catechol 2,3‐dioxygenase, (DmpB), mediates ring‐cleavage to initiate the sequential transformation to acetyl CoA and pyruvate by downstream meta‐pathway enzymes (DmpC to I).
29
1.6.1. Subservience of the DmpR/Po system to global regulatory input When the dmp‐system of pVI150 is transferred from Pseudomonas sp.
strain CF600 to the now genome sequenced strain of P. putida KT2440 it enables the new host to degrade phenolic substrates via the DmpR/Po‐controlled dmp catabolic pathway (124). However, even in the presence of phenol, the enzymes of the pathway are not produced during rapid growth on rich media (high‐energy conditions), becoming detectable only at the exponential to stationary phase transition (Fig. 11).
0 2 4 6 8 10 12
0,1
1
Gro
wth
(OD
650)
Time (hr)
0,0
0,2
0,4
0,6
0,8
1,0
C230 S
pecific activity
+ phenol - phenol
Fig. 11: The expression of the dmp‐operon can be monitored by an enzyme assay of one of the operon gene products, DmpB, which has catechol‐2,3‐deoxygenase (C23O) activity. P. putida KT2440 carrying a plasmid encoded dmp‐system does not express DmpB even in the presence of phenol in a rich media until the transition from exponential to stationary phase.
The finding that the genetic components of the dmpR gene and the
Po regulatory region are sufficient to mimic the growth phase response in both P. putida and E. coli suggested that host‐encoded factors involved in this response are functionally common to both strains (132). This allowed the use of E. coli mutants to help decipher the mechanisms underlying this phenomenon. Roles for some potential global regulators in mediating the growth phase response were excluded, such as σS, FIS, and quorum signalling molecules (132), and these early studies identified DmpR levels and IHF as important for the net transcriptional output from the σ54‐Po promoter. Subsequently, the alarm signalling molecule ppGpp was shown to be both required for transcriptional output from the σ54‐dependent Po promoter and to be a major link between the specific DmpR‐Po regulatory circuit and the physiological status of the cell. This level of global regulation dominates the DmpR‐mediated regulatory circuit so that Po is
30
essentially silent under nutritionally rich growth conditions (low ppGpp levels) and is only active when levels of ppGpp increase, for example i) under nutritionally poor conditions such as media lacking amino acids, ii) when nutrients are depleted as during the transition into stationary phase, or iii) when ppGpp levels are artificially manipulated (133).
transcription- + - + 2-mp- - + + ATP
ATP
NTPsDmpR
IHF
OH
σ54Core
-24 -12 +1UASs
terminator
transcription- + - + 2-mp- - + + ATP
transcription- + - + 2-mp- - + + ATP
ATP
NTPsDmpR
IHF
OHOH
σ54Core
σ54Core
-24 -12 +1UASs
terminator
Fig. 12. Aromatic effector‐ and ATP‐dependent transcription from the σ54‐Po promoter reproduced in vitro. The required components include the σ54‐RNAP holoenzyme, the Po regulatory region, DmpR, an aromatic effector and ATP required for DmpR activity, IHF required for maximal output, and NTPs.
Aromatic effector and ATP dependent transcription from the σ54‐Po
promoter can be fully reproduced in vitro with just a few purified components. These include the σ54‐RNAP holoenzyme, Po regulatory region, DmpR, an aromatic effector and ATP required for DmpR activity, IHF required for maximal output and NTPs (Fig. 12) (101, 149). Hence, these components represent targets for integration within the global regulatory network of the cell.
In the next section, I will present studies conducted on global regulation to deduce the molecular mechanisms of how the ppGpp signal is integrated to superimpose its message at the level of transcription from σ54‐dependent promoters.
31
AIMS The general aim of my research was to determine the molecular
mechanism(s) of how ppGpp, and later on DksA, exert their effects at the level of transcription from σ54‐dependent promoters. This general aim can be subdivided into different mechanistic questions as illustrated below:
-24 -12 +1
σ54
Core
-24 -12 +1
σ54
Core
ATP
OH
DmpR
ATP
OHOH
DmpR
IHF
Core andσ-factors
ppGppDksA
ppGppDksA
?
?
?
FtsHPtsNFtsHPtsN
1.Do global regulators
found to impact a closely relatedspecific regulatory circuit
also impact DmpR/Po?2.
Do ppGpp and DksAhave direct and/or indirect effects
on any of the processesof specific regulation?
3.Do ppGpp and DksA regulatory effects
through σ70-transcription explain the impact of these regulatory molecules
on σ54-dependent transcription ?
32
RESULTS AND DISCUSSION In this section, rather then going through the papers
chronologically, I present the key data under the mechanistic questions posed above.
1. Do global regulatory systems found to impact a closely related system also impact DmpR/Po? (Paper I)
The major regulators of the dmp‐ and xyl‐systems, DmpR and XylR, are two mechanistically related σ54‐dependent regulators that respond to distinct sets of aromatic effector compounds representing cognate pathway substrates and structural analogues (reviewed in 121). The Po promoter is the sole target for DmpR, while XylR controls the Pu (upper operon) promoter and the Ps promoter of second regulatory gene involved in controlling toluene catabolism. The Po and Pu promoters have three similar regions: a) the ‐24, ‐12 promoter motif recognized by σ54‐RNAP, b) an integration host factor (IHF) binding site required for optimal promoter output, and c) related upstream activating sequences (UAS) for regulator binding (1, 39, 125, 131). These common properties allow cross‐regulation of the Pu and Po promoters by DmpR and XylR in response to appropriate aromatic effectors (47).
Both systems are subject to dominant global regulation, which results in repression of transcription during exponential growth in rich media. In this work we were interested to determine (i) if two global regulatory factors (FtsH and PtsN) that impact the XylR/Pu system also influence the DmpR/Po system and (ii) if the alarmone ppGpp, previously analyzed predominantly in E. coli, affects the two systems to the same extent in the native P. putida host.
1.1. Is FtsH required for σ54‐Po transcription?
The levels of E. coli ATP‐dependent FtsH protease are controlled in response to physiological cues. This protease is an essential factor for aerobic survival of E. coli, with ftsH mutations requiring an additional suppressor mutation in order for the bacteria to grow (111). In an FtsH null E. coli strain, XylR‐mediated transcription from Pu is drastically reduced (24). As shown by the comparison in Fig. 13 (Paper I, Fig. 3A and B), the DmpR/Po system is fully functional while the XylR/Pu system is clearly defective in the absence of FtsH. Hence we concluded that FtsH is not required for DmpR/Po transcription, and is clearly not an essential factor
33
for σ54‐dependent transcription per se. As shown in the lower panels of Fig. 13, regulator swapping experiments clearly indicate that the regulator, rather than the promoter, determines sensitivity to the absence of FtsH (Paper I). Although the mechanism behind FtsH dependence of the XylR/Pu system is unknown, one speculation, based on work from others (96), is that the chaperone activity of FtsH may be required for proper assembly and/or disassembly of active multimers of XylR but not DmpR.
0,1
1
10
Gro
wth
OD
600
0
5
10
15
20
25A) XylR/Pu (pVI677)
B) DmpR/Po (pVI466)
E. coli: A8514 ( FtsH+ ) A8926 ( FtsH- )
0
1
2
3
4
5
RLU
/OD
600 x 10,000 (open symbols)
0 1 2 3 4 5 6
0,1
1
10
Gro
wth
OD
600
C) XylR/Po (pVI678)
Time (hr)0
5
10
15
20
25
0 1 2 3 4 5 6
D) DmpR/Pu (pVI679)
Time (hr)0
1
2
3
4
5
Fig. 13. Luciferase transcriptional profiles of Po and Pu promoters activated by their native regulators (panel A and B) or non‐native regulators (panel C and D) in E. coli FtsH+ (squares) and FtsH‐ (circles). Culture growth is shown with closed symbols and the luciferase response with open symbols (adapted from Paper I).
1.2. Is the DmpR/Po system under the control of PtsN‐mediated glucose repression?
The XylR/Pu system has been reported to be under the control of glucose repression by the ptsN gene, which encodes the IIANtr enzyme of an alternative phosphoenolpyruvate‐sugar phospho‐transferase system (PTS) (28, 31). As seen in Fig. 14 (Paper I, Fig. 4), the presence of glucose has no effect on the DmpR/Po system in P. putida while a previously observed 2‐fold decrease was found in the XylR/Pu system. Regulator‐swapping experiments again indicate that the susceptibility to glucose repression
34
depends on the identity of the regulators (Fig. 14). Hence, like FtsH, PtsN represents a regulatory factor that impacts the XylR/Pu system but not the DmpR/Po system and mediates its effect via the activity of the regulator rather than the properties of the promoter.
0
100
200
300
400
P. putida KT2440 derivatives in M9+CAA M9+CAA+glucose
XylR/Pu XylR/PoDmpR/PuDmpR/Po
Peak
act
ivity
(RLU
/OD
600 x
100
0)
Fig. 14. Peak luciferase activities of P. putida KT2440::dmpR‐Km and KT2440::xylR‐Km, carrying plasmids Po‐luxAB or Pu‐luxAB. Cells were grown in M9 media with 0.2% casein amino acids (CAA) and the appropriate aromatic effector, and further supplemented with 10 mM glucose where indicated (adapted from Paper I). 1.3. Influence of ppGpp on DmpR/Po and XylR/Pu circuits in E. coli and P. putida.
Previous work in E. coli identified ppGpp as the major signal that couples the DmpR/Po system to cellular physiology. However, while transcription from Po in ppGpp0 E. coli is dramatically reduced, transcription from Pu is minimally affected (26, 133). To assess the dependence of the two systems on ppGpp in their native host, a full ppGpp0 strain of P. putida was generated, as confirmed by chromatographic analysis (Paper I, Fig. 5). The transcriptional profiles of both DmpR/Po and XylR/Pu in P. putida confirmed previous observations in E. coli, i.e. that the DmpR/Po system is very dependent on ppGpp, although to a some what lesser extent than in E. coli (Paper I, Fig. 6A to D). In comparison, the expression profiles of the XylR/Pu system in both ppGpp+ and ppGpp0 P. putida still exhibited an abrupt shift in expression at the transition from exponential to stationary phase. Thus while the basic trends and main conclusions drawn from the work in E. coli were verified in P. putida, the magnitude of dependence of the two systems in the different hosts varies.
35
To see if the difference in the influence of ppGpp on the DmpR/Po and XylR/Pu systems could be ascribed to regulator or promoter properties, regulator‐swapping experiments were performed with both hosts. The data showed that in the E. coli ppGpp0 background transcription from both DmpR‐driven Pu and XylR‐driven Po were reduced (Paper I, Fig. 6E and F). Thus, having either component of the DmpR/Po pair renders the system more dependent on ppGpp, although not to the same extent (10‐fold) as for the original pair. In P. putida, the nature of the regulator appeared as the predominant component that determines the transcriptional outcome, since profiles with the cognate and non‐cognate promoters are very similar. Hence, both the nature of the regulator (most likely expression levels) and the promoter play a role in controlling the level of expression seen along the growth curve.
These studies led to the conclusions that: i) neither of the two factors identified to impact the XylR/Pu system (FtsH and PtsN) influences the DmpR/Po system, and ii) ppGpp impacts the DmpR/Po system to a much greater extent than the XylR/Pu system. Thus, even two very similar specific regulatory circuits can be locked into the global regulatory networks of bacteria by very different means. The predominant role of ppGpp in controlling transcription mediated from the DmpR/Po system lead to a focus on the role of this bacterial alarmone and how it dominates the specific DmpR/Po regulatory circuit in particular, and σ54‐dependent transcription in general. In addition, the observation that responses differed when monitored in E. coli and P. putida, prompted continual comparison between these two organisms.
2. Do ppGpp and DksA have direct and/or indirect effects on any of the processes of specific regulation?
To address this question we needed to develop a series of tools that included i) mutants incapable of synthesizing ppGpp and/or DksA, ii) a series of σ54‐dependent promoters that could be directly compared and in which key DNA elements could be manipulated with minimal disruptions, and iii) reconstituted in vitro transcription systems from both E. coli and P. putida. E. coli mutants were already available and corresponding P. putida mutants were constructed (Paper I and V). Likewise, the necessary purified E. coli derived proteins were already available, and P. putida counterparts were either purified or obtained from generous colleagues (Papers III and V).
36
Promoter upstream regions
XhoI NdeI
-170 -127 -52
UAS1 UAS2
-24,-12
hybrid σ54-promoters-24 -12 +1
CATATCAGACCTTGGCACAGCCGTTGCTTGATGTCCTGCGC Po/PoCATATCAAGCAATGGCATGGCGGTTGCTAGCTATACGAGAC Po/PuCATATCCACGGCTGGTATGTTCCCTGCACTTCTCTGCTGGC Po/nifHCATATCCACGGCTGGTATGTTTTTTGCACTTCTCTGCTGGC Po/nifH049CATATCAAAAATTGGCACGCAAATTGTATTAACAGTTCAGC Po/pspACATATCAAAAGTTGGCACAGATTTCGCTTTATCTTTTTTAC Po/glnA
IHFluxABdmpRdmpRdmpR
-388BglII
Hin
dIII
Po/Px
×xh-Pu/Px hybrids
xh-Po(-IHF)/Px hybrids
UP
Fig. 15. Hybrid σ54‐promoter regions of luxAB transcriptional reporters. Schematic illustration (not to scale) of the Po promoter region is shown together with the locations of the coding regions of dmpR and the luciferase luxAB reporter genes as open boxes. Arrow heads indicate the direction of transcription. The location of the DNA binding sites for DmpR (inverted shaded arrows; UAS1, UAS2) and the IHF recognition sequence (shaded box) are also indicated. The upstream regions that vary in xh‐Px/Px hybrid promoters, which contain a non‐native XhoI site (xh), have a disrupted IHF binding site or the upstream region from the Pu promoter (adapted from Paper V).
To obtain a series of directly comparable σ54‐dependent promoters,
we used the frame‐work of the Po promoter to ensure that appropriate promoter geometry was maintained and to place other well studied σ54‐promoters under control of DmpR. These series of hybrid promoters are schematically illustrated in Fig. 15 (Papers III and V). In the first series, designated Po/Px, the ‐33 to +2 region spanning the conserved −24, −12 promoter region from Po was replaced by the corresponding regions of other σ54‐promoters. These included the P. putida‐derived Pu promoter, the Klebsiella pneumoniae‐derived nifH promoter and its mutant derivative nifH049, in which substitution of the −17 to −15 CCC by TTT increases both affinity and transcriptional output, and the strong E. coli‐derived pspA and glnA promoters. In a second series, designated xh‐Po(‐IHF)/Px hybrids, the different promoters were combined with a disruption of the IHF‐binding site consensus that maintained the base composition of Po. Abolished IHF‐binding capacity as a result of these manipulations was confirmed by in vitro transcription assays (Paper III, Fig. 8B). In the final series, designated xh‐Pu/Px hybrids, the entire ‐116 to ‐40 region was replaced with that of the
37
Pu promoter. These hybrids still have the UAS binding sites for DmpR in the same location relative to the ‐24, ‐12 sequence, but have the appropriately positioned Pu UP‐element and IHF binding site that is located one helical turn from that in Po (see Fig. 15). At Pu, this region provides enhanced affinity for σ54‐RNAP by means of IHF mediated DNA architecture changes that allow simultaneous binding of both the α‐subunits of σ54‐RNAP to the distally located UP element (16, 25, 79). 2.1. Are both ppGpp and DksA required for efficient σ54‐dependent transcription in vivo?
The absence of ppGpp or DksA in both E. coli and P. putida results in a severe decrease in the transcriptional output from the σ54‐Po promoter as shown in Fig. 16 (Paper III and V). The simultaneous lack of both ppGpp and DksA essentially abolishes transcription from Po in E. coli (Paper III, Fig. 1A). The additive negative effect upon loss of both of these regulatory molecules is consistent with the proposed role of DksA in stabilizing the binding of ppGpp to RNAP. However, as DksA can also modulate transcriptional properties in the absence of ppGpp (103, 104), the ppGpp‐independent effects of DksA may also contribute to the net in vivo effect on σ54‐Po and other σ54‐promoters.
1
1
0 2 4 6
0.
Fig. 16. Both ppGpp and DksA are required for efficient transcription from the σ54‐Po promoter in P. putida and E. coli. Isogenic strains were used in both cases, and cultured in LB containing 2‐methylphenol. For P. putida KT2440 derivatives (left‐hand panel), dmpR was expressed in monocopy from the host chromosome, and transcription monitored using a Po‐luxAB reporter plasmid. For E. coli MG1655 derivatives (right‐hand panel), transcription was monitored using a dmpR‐Po‐luxAB plasmid (adapted from Paper III and Paper V).
Gro
wth
(OD
600)
(clo
sed
sym
bols
)
Time (hr)0 2 4 6
0.1
1
E. coli
Time (hr)
0.0
0.5
1.0
1.5P. putida
0.0
0.5
1.0
1.5
Wild-type DksA null
ppGpp0 ppGpp0 / DksA null
Transcription (open symbols)
(Wild-type set as 1)
38
The requirement for ppGpp for transcription from σ54‐Po in vivo appeared to be associated with the intrinsically low affinity of the Po promoter for σ54‐RNAP (131). This observation encouraged the direct testing of the requirement of ppGpp and DksA for transcriptional output from the Po/Px series of hybrid promoters described above. The relative affinities of the hybrid Po derivatives in the presence of increasing concentrations of σ54‐RNAP in electro mobility shift assays were shown to lie in the order Po/nifH < Po/Pu < Po/Po < Po/nifH049 < Po/glnA < Po/pspA (Paper III, Fig. 5B), with the first three hybrids (Po/nifH, Po/Pu, and Po/Po) constituting a group of relatively low‐affinity promoters and the second set (Po/nifH049, Po/glnA, Po/pspA) constituting a group of relatively high‐affinity promoters. Additionally, in vitro transcription data (Paper III, Fig. 6) shows that promoter occupancy kinetics differs such that the higher‐affinity promoters rapidly approach plateau values, while the lower‐affinity promoters show slower kinetics. These results defined the relative affinities and hierarchy of promoter occupancies of the hybrid promoters under in vitro conditions, which would presumably result in higher affinity promoters being occupied more easily by the available pool of σ54‐RNAP in vivo than lower affinity counterparts.
In E. coli, the temporal in vivo expression of all the hybrid promoters showed growth‐phase dependent profiles similar to Po, with absolute levels that differed by 18‐fold (Paper III, Fig. 7C). These differences in promoter output were essentially reiterated with an E. coli‐based in vitro transcription system (Paper III, Fig. 7D). The powerful high‐affinity hybrid promoters, such as the Po/pspA and Po/glnA, also allowed some level of transcription even in the exponential phase of growth. Most importantly, all the Po/Px hybrids were found to require ppGpp and DksA for full transcriptional output to an extent that reflects their affinity for σ54‐RNAP in vitro (Paper III, Figs. 5B and 7C), i.e. transcription from high‐affinity promoters was reduced ~2‐ to 3‐fold in the absence of ppGpp or DksA while transcription from low‐affinity promoters was reduced ~6‐ to 10‐fold.
2.2. What’s true for E. coli is (nearly true) for P. putida – an adaptive twist?
The nutritionally and environmentally adaptable Pseudomonads can thrive in and occupy a wide variety of niches including soils, aquatic environments, and in association with plants. Transcription from σ54‐promoters, as with any type of promoter, has to function within the context of the evolutionary selected regulatory network of the host bacteria. The
39
different lifestyles of P. putida and enteric E. coli are likely to have lead to adaptations to integrate responses to very different external stimuli. These considerations prompted a similar series of experiments in P. putida as those described for E. coli (Paper V).
Consistent with their nutritional adaptability, P. putida ppGpp0 and DksA null strains are prototrophic which contrasts the polyauxotrophic amino acid requirements of E. coli counterparts (Paper V). In P. putida, rather than the 18‐fold difference observed in transcriptional output from the lowest‐ to the highest‐affinity hybrid promoter of the Po/Px series in E. coli, only an ~3‐fold difference in the transcriptional output was found, but again dependency on ppGpp and DksA was found to correlate with promoter affinity for σ54‐RNAP (Paper V, Fig. 8A). This difference found with the Po/Px hybrid promoters as assessed with the two species was initially puzzling. However, we found that while P. putida derived σ54‐RNAP recognised the Po/Px hybrid promoters with the same hierarchy as that of E. coli, in vitro transcription with fully P. putida‐derived components reproduced the same ~3‐fold difference in the transcriptional output as seen in vivo (Paper V, Fig 8B). This suggests that the P. putida derived σ54‐RNAP is less discriminative of promoter differences that influence affinity for E. coli σ54‐RNAP. We infer from this data that this less‐discriminative property of P. putida σ54‐RNAP probably accounts for the robust transcription seen from even comparative low affinity promoters in P. putida.
In contrast to the high transcription observed in the post‐exponential phase of growth in P. putida, exponential phase transcription is very tightly controlled (Paper V, Fig. 4A to D). A combination of high‐affinity ‐24, ‐12 promoter features with the IHF‐dependent UP element of Pu to increase σ54‐RNAP binding, and the use of the XylR regulator that is naturally expressed at higher levels than DmpR, were simultaneously required to see any transcription in this phase of growth (Paper V, Fig. 4 F to I).
It had previously been established that ppGpp levels, as in E. coli, are vastly elevated at the exponential‐to‐stationary phase in P. putida (133). We used monocopy reporter gene fusions to the PdksA promoters in the chromosomes of these two organisms, and immuno‐detection of DksA levels to determine that transcriptional regulation and changes of protein levels across the growth curve were similar in each case, Fig. 17 (Paper V, Fig. 6). DksA levels peak in mid exponential phase but only vary ~2‐fold across the growth curve in both species. Thus, while σ54‐transcription
40
responses to ppGpp and DksA are similar in P. putida and E. coli, P. putida appears to have evolved and integrated these responses to provide both tighter control of σ54‐transcription under high energy conditions of the exponential phase of growth, with vigorous transcription upon nutrient limitation at the exponential‐to‐stationary phase transition.
0 1 2 3 4 5 6 7 8 9
0.1
1
Gro
wth
(OD
600)
(clo
sed
sym
bols
)
Time (hr)
0
5
10
15
20
P. putida E. coli
Transcription (open symbols)
(RLU
/OD
600 x 10,000)
DksA
t1 t2 t3 t4
E. coli
P. putida
PdksA-luxAB
Fig. 17. DksA expression is similar in E. coli and P. putida. Growth and luciferase activity of monocopy transcriptional reporter gene fusions on the chromosomes of P. putida (KT2440::PdksA‐luxAB) and E. coli (MG1655::PdksA‐luxAB). Both P. putida (squares) and E. coli (circles) strains were cultured at 30oC in LB and luciferase activity monitored over time. Triangles indicate the time points that cells were harvested for immuno‐blot analysis of DksA levels, which are shown in the lower part of the Figure. Immuno‐detection of DksA used 2 and 4 μg of SDS‐PAGE separated soluble protein prepared from the indicated strains (adapted from Paper V). 2.3. Do ppGpp and DksA have any direct effects on σ54‐dependent transcription in vitro?
In addition to the series of hybrid Po/Px σ54‐promoters, the potential direct effects of ppGpp and DksA on σ54‐transcription in vitro was compared with that of two σ70‐dependent promoters that had previously been shown to be either positively stimulated by ppGpp and DksA (σ70‐PthrABC) or negatively regulated by ppGpp and DksA (the stringent σ70‐rrnB P1) (103, 104). As illustrated in Fig. 18 (Paper III, Fig. 2B), the presence of ppGpp and DksA had no major stimulatory effect on transcription from σ54‐Po under conditions that recapitulated the known negative and positive effects on σ70‐dependent transcription. As is the case for Po, addition of
41
ppGpp and DksA had no significant stimulatory effects on in vitro transcription from the hybrid Po/Px derivatives with either E. coli or P. putida purified components (Paper III, Fig. 7D; Paper V, Fig. 8B).
0 1 2 3 4 50.1
1
0 75 150 225 3000.1
1
PthrABC Po rrnB P1
PthrABC
Po
rrnB P1
2 μM DksA + ppGpp 200 μM ppGpp +DksA
Fig. 18. Multiple‐round in vitro transcription in the absence or presence of ppGpp and DksA. Autoradiographs of titrations of ppGpp in the presence of constant concentrations of DksA (2 μM; left‐hand panel), and titrations of DksA in the presence of constant concentrations of ppGpp (200 μM; right‐hand panel). Templates: σ54‐Po, σ70‐PthrABC, and the σ70‐rrnB P1 promoters. Graphs show the normalized data for each promoter with the zero ppGpp or DksA value set as one (adapted from Paper III). Despite having no direct influence on σ54‐dependent transcription in vitro, both ppGpp and DksA do target σ54‐RNAP just as they do σ70‐RNAP. This was demonstrated by their negative effects on the lifetimes of competitor‐resistant open complexes (Paper III, Fig. 3; Paper IV, Fig. 6A to C). Decrease in competitor‐resistant open complex lifetime will only have impact on promoters that have this as a rate limiting step, e.g. σ70‐rRNA promoters. However, this reduction in lifetime does not result in reduced transcription from the σ54‐Po promoter under in vitro conditions, suggesting that open complex stability is not the rate‐limiting step for σ54‐dependent transcription from this promoter. The possibility that σ54‐dependent transcription from this promoter was influenced by the nature and levels of the initiating nucleotide was also investigated, but was not found to play any role (Paper III, Fig. 2).
Tran
scrip
ts
+ 2 μM DksA + 200 μM ppGpp
ppGpp [μM] DksA [μM]
42
The lack of any detectable direct effect of ppGpp and DksA on in vitro σ54‐transcription implies that these two regulatory molecules exert their effects in vivo by an indirect mechanism. The studies described in the following sections focus on the possible effects of ppGpp and DksA on the levels of key players involved in σ54‐Po transcription that could couple to host physiology.
2.4. Can the requirement for ppGpp and DksA be explained by their effects on the levels of IHF?
Transcription from the σ54‐Po promoter requires IHF‐mediated DNA‐bending for optimal DmpR and σ54‐RNAP interaction. The levels of IHF are partially under the control of ppGpp and increase at the transition from exponential to stationary phase of growth (9, 41, 140). Therefore, the influence of ppGpp on IHF levels might contribute to the ppGpp/DksA dependency of σ54‐Po transcription in vivo. To test this possibility, the hybrid Po/Px series and cognate derivatives containing disrupted IHF binding site consensus were used. This approach, as opposed to the use of an IHF‐deficient strain, avoids pleiotropic effects of the loss of IHF, which has global regulatory consequences in vivo (7).
In vitro transcription assays confirmed that all hybrid σ54‐promoters tested were strongly dependent on capacity to bind IHF (Paper III, Fig. 8B). Likewise, all hybrid σ54‐promoters were dependent on the IHF binding capacity for maximal in vivo transcriptional output in both wild‐type E. coli and P. putida (Paper III, Fig. 8C; Paper V, Fig. 3B). However, using the xh‐Po(‐IHF)/Px hybrids that lack the capacity to bind IHF, ppGpp and DksA were still both required for maximal promoter output from all derivatives tested, Fig. 19 (Paper III, Fig. 8D; Paper V, Fig. 5C). As with IHF binding proficient derivatives, simultaneous loss of both molecules essentially abolished detectable in vivo transcription in E. coli. Additionally, over‐expression of IHF had no significant effect on the transcriptional profile from σ54‐Po across the growth curve in wild‐type E. coli (133) or on any of the hybrid σ54‐promoters in P. putida (Paper V, Fig. 3C). These studies led to the conclusion that while IHF levels are important for optimal output from these σ54‐promoters, the mechanism by which ppGpp and DksA influence transcription from these σ54‐promoters is not through their effects on IHF levels.
43
0 2 4 6 8
0.1
1
G
row
th (O
D60
0) (c
lose
d sy
mbo
ls)
Time (hr)
0.0
0.5
1.0
xh-Po(-IHF)Po
Wild-type
ppGpp0
DksA null
ppGpp0 / DksA null
Transcription (open symbols)
(RLU
/OD
600 x 10,000)
Fig. 19. Transcription of an IHF‐binding incompetent σ54‐Po derived promoter still requires ppGpp and DksA. Growth (closed symbols) and luciferase activity (open symbols) of isogenic E. coli strains harbouring a transcriptional reporter plasmid (dmpR‐xh‐Po(‐IHF)/Po‐luxAB), cultured in LB containing 2‐methylphenol (adapted from Paper III).
2.5. Can the requirement for ppGpp and DksA be explained by their effects on the levels of DmpR or σ54?
Western blot analysis revealed that the temporal expression profiles of DmpR and σ54 in wild‐type and mutant E. coli ppGpp0 and DksA null strains are similar (Paper III, Fig. 1B). In both E. coli and P. putida, σ54 levels remained constant across the growth curve, while DmpR levels vary (Paper III, Fig. 1B, and Paper V, Fig. 2B). In E. coli DmpR levels increase ~2‐fold over the growth while in P. putida the elevation in DmpR levels is greater. Only lack of ppGpp was found to influence the levels of either of these two key proteins: σ54 levels were unaffected while DmpR levels were reduced ~2‐fold in both organisms (Paper III, Fig. 1B; Paper V, Fig. 2B). Compensatory experiments were performed in E. coli to provide DmpR levels in the ppGpp0 strain that slightly exceeded those in the wild‐type. However, this compensation did not alter the degree of ppGpp‐dependency of transcription from σ54‐Po. This lead to the conclusion that while DmpR levels are obviously critical for the output from the σ54‐Po promoter, ppGpp and DksA exert their action in vivo through a mechanism that is independent of changes of either DmpR or σ54 levels.
44
3. Do ppGpp and DksA regulatory effects through σ70‐transcription explain the impact of these regulatory molecules on σ54‐transcription?
The work outlined in the proceeding sections established i) that ppGpp and DksA have little if any direct influence on the performance of σ54‐RNAP on a range of σ54‐ promoters tested in in vitro transcription assays, ii) that the major effects on transcription in vivo can not be accounted for by ppGpp‐associated changes in DmpR, IHF or σ54‐levels, and iii) that the influence of ppGpp and DksA on σ54‐dependent transcription is related to the innate affinity of the promoters for σ54‐RNAP. However, these data do not provide any obvious link between ppGpp and σ54‐dependent transcription. So how is ppGpp linked to σ54‐dependent transcription?
3.1 Suppressor mutations reveal a link between ppGpp and σ54‐dependent transcription
Some RNAP mutants isolated in the laboratory of M. Cashel on the basis of restoration of prototrophy had previously been shown to restore efficient transcription to the σ54‐Po promoter in ppGpp‐deficient E. coli (133). This suggested that ppGpp and DksA are likely to have their effects through RNAP, but given the data already described, it was not obvious how this could be so. This prompted a genetic approach to try to identify the players or mechanisms that could provide the link (Paper III). The strategy employed to isolate suppressor mutants was designed to specifically target DmpR‐regulated Po promoter activity. This involved the use of a selection plasmid carrying the minimal dmp‐system controlling the expression of a promoter‐less tetracycline resistance gene in an E. coli ppGpp0 strain. This approach resulted in the isolation of many spontaneous mutants. The vast majority mapped to the components of the transcriptional machinery. DNA sequencing identified two independent mutations within the rpoD gene that encodes σ70, nineteen independent mutations within rpoB that encodes the β‐subunit of RNAP, and twelve independent mutations within rpoC that encodes the β’‐subunit of RNAP (Paper II, III and IV).
The degree of restoration of transcription from σ54‐Po by these different mutants ranged from a barely detectable difference from that observed in the ppGpp0 parent to >3‐fold higher levels than that observed in the ppGpp proficient wild‐type strain in vivo (Papers II, Fig. 1B and IV, Fig. 1A), which translate to >30‐fold enhanced transcription in the absence of ppGpp. This data clearly suggested that ppGpp operates through the
45
transcriptional apparatus, but raised the obvious question of how do mutations in σ70 restore transcription to the σ54‐dependent Po promoter? Given the distinct classes of promoters recognized by the two σ‐factors, it is unlikely that the σ70 mutants directly affect Po output. The most plausible interpretation of this data was that the σ70 mutants mediate their effects through defects in their association with the common substrate for the two σ‐factors, namely core RNAP.
3.2. σ70 mutants are defective in competition with σ54 for core RNAP
This series of experiments used the two σ70 mutants identified in the genetic screen described above (designated σ70‐40Y and σ70‐35D), and two previously isolated σ70 mutants identified on the basis of restoration of prototrophy (designated σ70‐P504L and σ70‐S506P) (58). The σP
70 mutants all have substitutions that are likely to directly or indirectly alter the path of the σ70‐3.2 loop and/or interaction of σ with the β‐flap (Paper II). 4
To test for possible defects in competition for core RNAP, we established an in vitro competition assay that allowed simultaneous monitoring of transcription from the σ54‐Po promoter and the σ70‐dependent promoter PRNA1. As a first step in the in vitro analysis, we established the parameters of wild‐type proteins. Consistent with previous data from others, we found that the apparent affinities of σ54 and σ70 for core RNAP are similarly high when assessed in isolation (Paper II, Fig. 2B and C). However, under a situation of competition provided by a constant level of core and σ70 with increasing amounts of σ54, or vice versa, clear differences became apparent: σ54 is significantly poorer at out‐competing a fixed concentration of σ70 than the converse (Paper II, Fig. 3A and 4B).
In vitro competition between the σ70‐mutants and σ54 demonstrated that they indeed were defective in competitive association for core RNAP, with their deficiencies lying in the order σ70‐40Y > σ70‐P504L > σ70‐S506P > σP
70‐35D (Paper II, Fig. 3). This order correlated with the phenotypes of these mutants on transcription from σ54‐Po in ppGpp0 E. coli (Paper II, Fig. 1). The most severely defective mutant (σ70‐40Y) restored transcription to approximately 1.5‐fold the levels seen in the ppGpp+ parent, the intermediately affected mutants (σ70‐P504L and σ70‐S506P) restored transcription to approximately wild‐type levels, while the least affected mutant (σ70‐35D) had barely detectable effects in vivo. The σ70‐40Y and σ70‐35D mutants were also tested for their ability to restore transcription from σ54‐Po in the absence of DksA. Again the magnitude of their defects in competitive association for core RNAP correlated with their in vivo phenotypes; σ70‐40Y resulting in
46
restoration of transcription from Po in the DksA null mutant to ~1.5‐fold of the levels observed in wild‐type, while the minimally affected σ70‐35D resulted in ~0.5‐fold the levels in wild‐type E. coli (Paper III, Fig. 1D)
These data established that the requirements for ppGpp and DksA for σ54‐transcription from Po could be bypassed in vivo by mutants of σ70 that are defective in competitive association for limiting core RNAP. This data strongly suggested that ppGpp and DksA acted together, either directly or indirectly, to control σ54‐transcription through competition for core RNAP. However, no detectable effects of ppGpp and DksA could be detected in in vitro competition assays between σ70 and σ54, Fig. 20 (Paper III, Fig. 4). Hence, we favour a purely passive model for ppGpp and DksA effects on σ54‐transcription in vivo.
σ54-Po transcripts
σ54 [nM]0 50 100 150 200
0
20
40
60
80
100
No additions + ppGpp/DksA
0
20
40
60
80
100
No additions + ppGpp/DksA
σ70-R
NA
1tra
nscr
ipts
σ70-R
NA
1tra
nscr
ipts
σ70-R
NA
1tra
nscr
ipts
PoRNA1
Fig. 20. Multiple‐round in vitro competition between σ70 and σ54 for core RNAP in the absence or presence of ppGpp and DksA. Transcript levels are given as a percentage of those achieved for σ70‐RNA1 in the presence of 20 nM σ70 and the absence of σ54 (‐‐‐‐‐‐‐), or for σ54‐Po in the presence of 160 nM σ54 and in the absence of σ70 (_______), under each condition (adapted from Paper III).
47
The concept that alternative σ‐factors compete for their common substrate – core RNA polymerase – has a long history and has been proposed to account for a number of observations relating to the poor performance of promoters controlled by alternative σ‐factors. Up until recently, the numbers of σ‐factors in the cell was thought to only be slightly in excess of the number of core RNAP available for association to form the RNAP holoenzyme. However, recent reassessments estimate that in E. coli grown in rich defined media, ~13,000 molecules of core RNAP are available for association with ~17,000 molecules of σ70 (52). While being approximately constant throughout growth, the levels of σ70 always exceed those of σ54, and σ54 levels are maintained at about 16‐20% of the level of σ70 (67). Upon entry in stationary phase, the levels of σS also come into play, and are dramatically increased to about 30% of the level of σ70 (64). Hence, conditions in the cell are such that competition for core RNAP is likely to be fierce.
3.3. σ‐factor competition limits σ54‐dependent transcription in vivo
Manipulation of the availability of different σ‐factors was used to experimentally establish if competition between σ‐factors limited σ54‐dependent transcription in vivo. The first clues that competition for limiting core might be involved in controlling the output from the σ54‐Po promoter came from the early observation that transcription from this promoter is higher in the absence of σS (132). This observation suggested that σS and σ54
competed with each other and led to a direct comparison of transcription from the σ54‐Po promoter in cells either devoid of σS or expressing excess σ54. The data confirmed that lack of σS results in ~2‐fold higher transcriptional output, while elevated levels of σ54 resulted in ~2.5‐fold higher transcriptional output (Paper II, Figs. 5C and A).
σ70 levels are slightly elevated in ppGpp0 E. coli (58). Therefore, we employed a genetic system with the σ70 gene (rpoD) under the control of the Ptrp promoter to i) be able to determine the effect of ppGpp on σ54‐Po transcription independently of its effects through σ70 levels and ii) to be able to artificially under‐produce the essential σ70 factor. As summarised in Fig. 21 (paper II, Fig. 6), using this approach, simple balancing of the levels of σ70 in ppGpp+ and ppGpp0 counterparts to those of wild‐type ppGpp+ E. coli drastically reduced the effect of the absence of ppGpp on σ54‐Po transcription (compare open and shaded bars in left panel in Fig. 21). Furthermore, underproduction and sequestering of σ70 by the σ70‐binding protein Rsd resulted in >10‐fold increase in maximal σ54‐Po transcription,
48
and transcription during exponential growth where ppGpp levels are low and the Po promoter is normally silent (right panel in Fig. 21). Subsequent to these findings, work from the laboratory of T. Nyström demonstrated that underproduction of σ70 mimics a stringent response as assessed on the proteomic level (81).
Fig. 21. Effect of modulation of in vivo σ70 levels on transcription from the Po promoter. Expression of rpoD is from the IAA‐dependent Ptrp promoter (Ptrp‐rpoD). Left‐hand panel shows the transcriptional output from the Po promoter of a luciferase reporter plasmid (dmpR‐Po‐luxAB) in ppGpp0 CF1693Δlac Ptrp‐rpoD (open bars) or in ppGpp+ MG1655Δlac Ptrp‐rpoD (shaded bars) that harbour either the plasmid pBR‐Rsd (indicated as Rsd ↑) or the pBR322 vector control (indicated as Rsd ‐). Results are the average of data from two independent experiments with cultures inoculated into LB supplemented with 0.2 mM IAA (indicated as σ70 – ) or 0.002 mM IAA (indicated as σ70 ↓). Right‐hand panel shows the immediate Po‐transcriptional response (open symbols) upon down‐regulation of σ70 levels in CF1693Δlac Ptrp‐rpoD (circles) and its ppGpp proficient counterpart (squares) MG1655Δlac Ptrp‐rpoD. Growth as measured by OD600 is indicated with closed symbols (adapted from Paper II). The results from the in vivo assays establish that σ54 is under significant competition by both σ70 and σS which results in restricted transcription from σ54‐Po, and supports the idea that ppGpp effects on this process are indirect (passive) since the same effects were observed in ppGpp+ and ppGpp0 cells alike.
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49
3.4. Mutations within β and β´ reveal a dual bypass mechanism that directly links σ54‐transcription to properties associated with down‐regulation of stringent σ70‐promoters.
All the mutations in the β‐ and β´‐subunits of RNAP that were identified in the genetic screen described under section 3.1, map to the active site cleft of RNAP (Paper V, Table 2). Five of these mutants (β‐R451C, β‐H551Y, β‐H1244L, β‐Q1264P and β’‐L432R) were found to give a hyper‐suppressor phenotype on σ54‐Po transcription, resulting in 20‐ to 60‐fold higher transcription levels than wild‐type RNAP in ppGpp0 E. coli, Fig. 22 (Paper IV, Fig. 2B to F).
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Fig. 22. rpoBC alleles bypass the requirement for ppGpp and DksA. Growth profiles (closed symbols) and transcriptional profiles (open symbols) of strains harbouring a dmpR‐Po‐luxAB plasmid. Strains possess either wild‐type (wt) alleles or the indicated rpoBC alleles in ppGpp+ DksA+ (squares), or its otherwise isogenic ppGpp0 (circles) and DksA null (triangles) counterparts. The data for rpoBC‐wt are shown for comparison. Note the change of scale for the rpoBC mutations in panels B–F, in which the dashed line indicates the maximal wild‐type level (adapted from Paper IV).
These hyper‐suppressors were also able to restore transcription
from the σ54‐Po promoter in E. coli lacking DksA to 0.5‐ to 2‐fold levels of those found in the wild‐type strain (Fig. 22). However, transcription from σ54‐Po in the absence of DksA was found to be notably lower than that in the absence of ppGpp in all cases, suggesting a critical role for this protein
50
in the normal functioning of the cell that is not fully compensated for by the mutations in core RNAP. The essentially ppGpp‐independent behaviour of these mutants in vivo occurs despite the fact that core RNAP mutants remain co‐responsive to ppGpp and DksA in vitro when assessed on the σ70‐rrnB P1 promoter (Paper IV, Fig. 5B). This suggests that transcription from the σ54‐Po promoter has reached a ceiling in each of these strains that is the maximal level that can be driven by each of the mutant polymerases.
In vitro comparison of σ54‐dependent and σ70‐dependent transcription with the core RNAP mutants showed that they mediate 1.1 to 1.6 fold higher relative σ54‐dependent transcription (Paper IV, Fig. 3E), suggesting that these mutants may be comparatively defective in their association with σ70. In vitro assessment of their potential defects in competitive association with σ70 and σ54 supported this suggestion, since the levels of the defects were recapitulated under in vitro competition conditions (Paper IV, Fig. 4). However, while all the core RNAP mutants exhibited defects, they were not as extensive as those found for the σ70‐40Y mutant, and were closer to those observed for σ70‐P504L and σ70‐S506P. Hence these differences, although significant, appeared insufficient to fully explain their hyper‐suppressor phenotypes.
P
To identify other potential defects of the core RNAP mutants that might contribute to their in vivo hyper‐suppressor phenotypes, we assessed the lifetimes of competitor‐resistant RNAP‐promoter complexes in relation to the lifetimes of the wild‐type RNAP holoenzymes in the presence of ppGpp and DksA, Fig. 23A to C (Paper IV, Fig. 6). This analysis used the σ54‐Po promoter, the σ70‐λPL promoter, and the rrnB P1 promoter at which ppGpp and DksA effects on this step are thought to be critical for transcriptional down‐regulation in response to these molecules. Although ppGpp and DksA decrease the lifetimes of RNAP‐promoter open complexes at both the σ54‐Po promoter and the σ70‐λPL promoter, this step is not rate limiting, and does not result in reduced transcription.
As summarised in Fig. 23D to F, all the mutants reduced the lifetime of RNAP‐open complexes at all the promoters. However, for σ54‐Po and σ70‐λPL, the pattern observed with the mutant RNAP holoenzymes was similar, and the reduced lifetimes that result from these mutations is above those evoked by ppGpp and DksA with the wild‐type RNAP holoenzyme (Fig. 23D and F). This contrasts the results with the σ70‐rrnB P1 promoter. At this promoter, with the exception of β‐Q1264P, the mutant polymerases form intrinsically unstable RNAP‐promoter complexes that are close to or below those found with the wild‐type in the presence of ppGpp and DksA (Fig.
51
23E). These findings prompted re‐examination of the σ70 mutants. The σ70 mutants were also found to mediate specific defects in the lifetimes of open RNAP‐promoter complexes at the σ70‐rrnB P1 promoter (Paper IV, Fig 7), that like their defects in σ‐factor competition assays, also correlated with their in vivo phenotypes.
The vast majority of the suppressor mutants have two defects which could combine to explain their in vivo phenotypes on σ54‐dependent transcription from Po: i) modulation of σ‐factor to core association to favour σ54 over σ70, and ii) reduced innate stability of competitor‐resistant RNAP‐promoter complexes, which mimics the effects of ppGpp and DksA for negative regulation of stringent σ70‐promoters. Both these properties support the purely passive model described below that is put forward to explain regulation of σ54‐transcription by ppGpp and DksA.
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Fig. 23. The effects of ppGpp and DksA on the lifetimes of open promoter complexes with wild‐type or mutant RNA polymerases. The lifetimes of competitor‐resistant complexes generated with 10 nM core RNA polymerase and 80 nM σ‐factor. A)–C) Time courses in the presence of no additions (squares), 200 μM ppGpp (triangles), 200 nM DksA (circles), or both (open diamonds), are shown with the values found at 20 sec in the absence of any additions set as 100%. Templates used: A. σ54‐Po (11 nM), B. σ70‐rrnB P1 (7 nM), C. σ70‐λPL, (11 nM). D)–F) Were performed as for panels A–C using wild‐type or mutant core enzymes in the absence of ppGpp and DksA, and monitored at the indicated times after competitor additions. Bars 1 to 6 in panels E and F are as indicated in panel D. Dashed lines indicate the corresponding level of the wild‐type holoenzyme in the presence of ppGpp and DksA (adapted from Paper IV).
52
3.5. ppGpp and DksA mediated regulation of σ54‐transcription – the current model
The effects of ppGpp and DksA on σ54‐dependent transcription are major, with simultaneous loss of these regulatory molecules essentially abolishing transcription from the σ54‐Po promoter. Based on the collective results presented in this work, any model for ppGpp and DksA regulation of σ54‐transcription has to take into consideration the following: 1) That the major effects of ppGpp and DksA do not function indirectly through the levels of DmpR or other proteins directly required for σ54‐transcription. 2) That σ54 is under significant competition with other σ‐factors in the cell. 3) That ppGpp and DksA have no apparent direct effects on σ54‐dependent transcription in vitro, either alone or when placed in competition with σ70. 4) That the effects of ppGpp and DksA in vivo are related to the affinity of promoters for σ54‐RNAP. 5) That mutants of σ70, and the β‐ and β’‐subunits of RNAP that can bypass the need for ppGpp and DksA in vivo have defects that would favour the formation of σ54‐RNAP, and mimic the effects of ppGpp and DksA for negative regulation of stringent σ70‐promoters.
The proposed passive model for control of σ54‐dependent transcription functions through predicted global regulatory consequences of the negative action of ppGpp and DksA at the seven powerful stringent σ70‐rRNA operon promoters. While the levels of ppGpp change dramatically, DksA levels only vary ~2–fold across the growth curve. Thus, this passive model primarily operates through changes in ppGpp levels. The powerful stringent σ70‐rRNA operon promoters sequester ~60‐70% of the transcriptional machinery during rapid‐to‐intermediate growth on rich media (18). Thus, under these conditions, in which ppGpp levels are low (Fig. 24A), sequestering of the majority of the transcriptional machinery by the rRNA operons would leave little free core RNAP available for association with the constant levels of σ54 and σ70, leading to low levels of σ54‐RNAP holoenzyme and consequent low promoter occupancy and output from cognate σ54‐promoters. However, under slow growth and stress conditions that elicit high levels of ppGpp (Fig. 24B), the dramatic down‐regulation of transcription from the σ70‐rRNA operon promoters would lead to increased levels of core RNAP available for association with σ‐factors. As a result, σ54‐RNAP holoenzyme levels would increase, leading to enhanced σ54‐promoter occupancy and output.
53
Nutrition & other stressA) Low ppGpp B) High ppGpp
σ70-rRNA operons
σ54-operons
σ70-rRNA operons
σ54-operons
core RNAPσ70
σ54
Fig. 24. Passive model for ppGpp/DksA control of σ54‐dependent transcription. A) Growth conditions that elicit low levels of ppGpp. Availability of key transcriptional components is illustrated schematically. As detailed in the text, ~60‐70% of the transcriptional machinery is sequestered in expressing the abundant transcripts from the multiple powerful rRNA operon promoters under these conditions. Thus, only low levels of core RNAP would be available for association with different σ‐factors, leading to low levels of σ54‐holoenzyme. B) Conditions that elicit high levels of ppGpp. Reduced transcription from the rRNA operon promoters would increase the levels of core RNAP available for holoenzyme formation, leading to higher levels of σ54‐RNAP holoenzyme (adapted from Paper III).
Within this model, both core RNAP‐ and σ70‐suppressor mutants
would mimic the effects of ppGpp and its co‐factor DksA in two ways: 1) by directly reducing transcription from the powerful stringent σ70‐rRNA operon promoters to elevate the levels of core RNAP for association with other σ‐factors, and 2) by altering RNAP core to σ‐factor association to favor the formation of σ54‐RNAP holoenzyme over that of σ70‐RNAP holoenzyme. In the case of the core RNAP mutants, these effects would further amplify the enhanced output from the σ54‐Po promoter that is found with these mutants RNAP holoenzymes.
The model as outlined above would predict that low‐affinity promoters that have σ54‐RNAP holoenzyme binding as a rate‐limiting step would be more susceptible to loss of these regulatory molecules than their high‐affinity counterparts. This indeed appears to be the case since among six σ54‐promoters tested, those with lower affinity for σ54‐RNAP were found to exhibit greater sensitivity to loss of ppGpp and DksA in vivo than high‐affinity counterparts.
54
CONCLUDING REMARKS The mechanism proposed above for ppGpp/DksA‐mediated passive
regulation of σ54‐dependent transcription is also pertinent to the functioning of other alternative σ‐factors. In addition to σ54, DksA and/or ppGpp have been shown to be important for transcription from promoters dependent on σS, σH and σE that control processes critical for stress survival (37, 68). In the case of σS, mutant RNAPs that mimic the effects of ppGpp have also been shown to bypass the requirement for ppGpp in vivo by a mechanism that is independent of ppGpp effects on σS levels (68, 76). Although the direct effects of ppGpp in synergy with DksA have not been tested at these σS‐, σH‐ and σE‐dependent promoters, it appears likely that passive regulation of the levels of the cognate holoenzyme would at least in part contribute to regulation of their transcription and to the pleiotropic roles of ppGpp and DksA in controlling the activity of promoters in response to nutritional and other stresses.
So is it all purely passive? The findings that ppGpp and DksA do not affect competition between σ54 and σ70 in vitro, contrasts with the previously observed direct effects of ppGpp on competition between σH and σ70 (68). Hence it is certainly possible that for some σ‐factors, ppGpp and DksA may directly enhance their ability to compete with σ70 and/or each other. In addition, the results presented do not exclude the possibility that a currently unknown regulator(s) could also contribute to the requirement for ppGpp and DksA for efficient σ54‐dependent transcription in vivo. Akin to the discovery of the role of DksA in ppGpp‐mediated regulation of σ70‐transcription, it is possible that some yet unknown factor could also act in conjunction with ppGpp and DksA to directly control σ54‐dependent transcription, or alternatively, could directly affect σ‐factor competition for limiting core RNAP. However, only in the latter case could the existence of such an unknown player account for the σ54‐promoter affinity‐dependent differences in the requirement for ppGpp and DksA.
55
ACKNOWLEDGEMENTS
When I first landed in Umeå, my first thought was: “Where the hell am I? This is going to be a short term experience!! Take a deep breath and make the most of it while you’re here!”. Seven years later, here I am happily thanking everybody that made my life here in Umeå unforgettable, in and out of the department of Molecular Biology. I want to start to thank the person that had the courage to take me in her lab and teach me a lot of what I know today, my supervisor Victoria Shingler. Thank you very much for all the guidance, the pushing forward, the enthusiasm and the problem solving and, last but not least, the help during the writing of this thesis. Many thanks to past and present members in the lab: Andrew, Chun Chau (pity I didn’t have a chance to work with you longer), Inga, “crazy‐red‐haired polish” Agnieszka, “always giggling” Petra, “adopted spanish proper lady” Teresa, “always giggling part II” and training addict Linda (wish you the best of luck!) and the mother of us all, my dear Eleonore. Mustafa Jason! Go Newcastle! Go Hull! Thanx for the music in the lab, thanx for the England trip and many more. Hope to see you soon, mate! Ciao, bella Dafne, you are a very good and special friend, in and out of the lab. I wish you and Bunny Boy the best of luck, you deserve it! A special obrigado to Fernando and Perpétua for helping me a lot during the very beginning, for always advising me and for keeping the portuguese alive. I big smile up to Sugar Baby Mário! I will always remember those great dinners and the picture we took at Nydala with the rest of the foreign gang!! Swiss António, Jojo, Boióla Vinicius and querida Tania (tenho saudades vossas; boa sorte com o junior!) lets do that Brasil trip again and finish killing our liver!! Thanx to “curly baixinha” Nadia, “crazy rhum‐drinking” Joan, Ingela, “kanguru” Kurt for the great dinners and parties. Wish you guys best of luck. Magnus, Maria, Hans and Gosinha, thanx for the great conversations, airsoft and dark parties ;) I want to thank Carlos “Hombre” and Anna (soon it’s your turn, hehe!) for being good personnel friends and ppGpp partners in crime. Big thanks to all the people that listened and/or slept during bacterial seminars, the Hans Wolf‐Watz clan: Sara (your going to be done in 6 days!), Margareta, Janne (thanx for reparing my bike so many times!), Elin, Ann‐Cathrine, Rolle I, Rolle II; Matt Francis (thanx for always being around for a good chat and a cold beer, mate!) & Co: Katrin, Petra; Cathrine Persson, Jenny and Miss 2M Maria; Debra Milton, Barbara, Medisa and the Åke Forsberg and his FOI crew. For those that are not listed, I apologize! Thanx to “Almighty” Johnny (don’t forget your friend when you retire early!) that has always delivered the goods on time; to the secretaries and the media personnel many thanx for making our lives easier at the department.
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Tack så jätte mycket för allt möjligt till mina svenska kompisar: Nasse, Jenny, Åsa, Andreas, Kiwi, Dr. Johanna. Um muito obrigado especial para ti Lotta, ajudaste‐me imenso e estiveste ao meu lado durante estes ultimos anos. Tiraste‐me do lab para viajarmos, sem ti nao tinha conhecido o que conheco da Suécia, sem ti nao tinha conhecido muitos dos meus amigos suecos, foste tu que me ajudaste a aprender sueco, a esquiar e muito mais. Sem tudo isto nao tinha conseguido fazer esta tese. Desejo‐te muita sorte para o teu doutoramento. Beijo e muito obrigado! Cumprimentos ao clã Sandberg.
Prá malta do Porto: Joao Juani, Nuno Cigano, Duarte Zum, Daniel Demo, Raquel e Joana Desalojada, espero que possamos manter contacto e amizade para sempre. Boa sorte prá vocês!! Para o meu irmao Hugo (contigo para sempre até ao fim!), Xana e sobrinho Diogo e pró resto da familia Tavares um forte abraco e um muito obrigado!
Por fim, um grande muito obrigado prós meus pais. Obrigado pela ajuda, por me terem sempre apoiado e aconselhado, e por nunca dizerem não (salvo raras excepcões, hehe!!). Sem vocês nao tinha chegado onde cheguei nem tinha conseguido o que consegui. Beijo e abraco!
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