Pseudomonas Putida

Multiple regulatory inputs for hierarchical control of phenol catabolism by Pseudomonas putida

W.K. Anjana Madhushani

Department of Molecular Biology Umeå University, Sweden Umeå 2015

To my Family…….

Contents

Table of Content…………………………………………………………………………………i Abstract………………………………………………………………………………………….iii Abbreviations……………………………………………………………………………...... iv Papers included in the Thesis…………………………………………………………………...v 1. Introduction………………………………………………………………………………..1 1.1 Environmentally useful bacteria………………………………………………………...1 1.2 Control of bacterial gene expression – from DNA to protein…………………………...3 1.2.1 The bacterial transcription cycle…………………………………………..4 1.2.2 Players in transcription……………………………………………………5 1.2.2.1 The catalytic transcription machinery……………………… …….5 1.2.2.2 Specificity (σ) factors……………………………………………..6 1.2.3 Regulation of transcription………………………………………………..7 1.2.3.1 Holoenzyme pools………………………………………………...7 1.2.3.2 Classical DNA-binding transcriptional regulators………………...7 1.2.3.3 bEBPs – specialized mechano-transcriptional activators of σ54……………………………………………………9 1.2.3.4 Global co-regulation by architectural proteins…………………..11 1.2.3.5 Transcriptional control through RNAP binding of (p)ppGpp……………………………………………..12 1.2.3.6 Control of transcriptional elongation through RNA switches – attenuators and riboswitches…………………..13 1.2.4 Translation……………………………………………………………….14 1.2.4.1 RNA structure and translational regulation……………………...14 1.2.4.2 Anti-sense sRNA mediated translational control.………………..15 1.2.4.3 RNA-binding proteins and translational control.………………...17 1.2.5 The metabolically versatile pseudomonads.……………………………..18 1.2.5.1 The major CRC mechanism of P. putida...………………………18 1.2.5.2 P. putida CF600 and the dmp-system……………………………19 2. Aims………………………………………………………………………………………..23 3. Results and Discussion……………………………………………………………………24 3.1 A promoter-distal ATAAATA DNA motif reduces the levels of mRNA transcripts ……………………………………………………………………………...24 3.1.1 Potential mechanisms underlying the action of the ATAAATA DNA motif ………………………………………………………………25 3.2 The CA-motif within the 5’-LR results in translational inhibition of dmpR…………..27 3.2.1 Crc-exerts translational regulation on dmpR in vivo……………………..29 3.2.2 Crc and Hfq act as co-partners in formation of riboprotein complexes in vitro………………………………………………………..29 3.3 Crc and Hfq both participate in catabolic repression………………………………….32 3.3.1 Crc-Hfq input on DmpR levels and (methyl)phenol catabolism………...32 3.3.2 The dmp-system has multiple additional CA-motifs for targeting by Crc-Hfq………………………………………………………………33

3.4 Current model for hierarchical regulation of (methyl)phenol catabolism by the dmp- System…………………………………………………………………………...34 Acknowledgements ………………………………………………………………………38 References ………………………………………………………………………………..40 Papers I-III

Abstract

Metabolically versatile bacteria have evolved diverse strategies to adapt to different environmental niches and respond to fluctuating physico-chemical parameters. In order to survive in soil and water habitats, they employ specific and global regulatory circuits to integrate external and internal signals to counteract stress and optimise their energy status. One strategic endurance mechanism is the ability to choose the most energetically favourable carbon source amongst a number on offer.

Pseudomonas putida strains possess large genomes that underlie much of their ability to use diverse carbon sources as growth substrates. Their metabolic potential is frequently expanded by possession of catabolic plasmids to include the ability to grow at the expense of seemingly obnoxious carbon sources such as phenols. However, this ability comes with a metabolic price tag. Carbon source repression is one of the main regulatory networks employed to subvert use of these expensive pathways in favour of alternative sources that provide a higher metabolic gain. This thesis identifies some of the key regulatory elements and factors used by P. putida to supress expression of plasmid-encoded enzymes for degradation of phenols until they are beneficial.

I first present evidence for a newly identified DNA and RNA motif within the regulatory region of the gene encoding the master regulator of phenol catabolism – DmpR. The former of these motifs functions to decrease the number of transcripts originating from the dmpR promoter, while the latter mediates a regulatory checkpoint for translational repression by Crc – the carbon repression control protein of P. putida. The ability of Crc to form repressive riboprotein complexes with RNA is shown to be dependent on the RNA chaperone protein Hfq – a co- partnership demonstrated to be required for many previously identified Crc-targets implicated in hierarchical assimilation of different carbon sources in P. putida. Finally, I present evidence for a model in which Crc and Hfq co-target multiple RNA motifs to bring about a two-tiered regulation to subvert catabolism of phenols in the face of preferred substrates – one at the level of the regulator DmpR and another at the level of translation of the catabolic enzymes.

iii

Abbreviations

AAA+ ATPases associated with various cellular activities αNTD The amino-terminal domain of the α-subunit αCTD The carboxyl-terminal domain of the α-subunit ATP Adenosine triphosphate bEBPs bacterial Enhancer Binding Proteins bp base pairs CA Catabolite Activity cAMP Cyclic adenosine monophosphate CAP Catabolite Activator Protein c-di-GMP Bis-(3’-5’)-cyclic dimeric guanosine monophosphate CRC Catabolite Repression Control DNA Deoxyribonucleic acid Dmp Dimethylphenol Fis Factor for inversion stimulation HNS Histone-like nucleoid-structuring IHF Integration Host Factor LB Luria-Bertani/Lennox broth LR Leading region mRNA messenger RNA nt Nucleotides ORF Open-reading frame ppGpp Guanosine tetraphosphate pppGpp Guanosine pentaphosphate (p)ppGpp ppGpp and pppGpp Po Promoter for the dmp operon Pr Promoter for dmpR PTS Phosphoenolpyruvate carbohydrate phosphotransferase system RBS Ribosome binding site rRNA ribosomal RNA RNA Ribonucleic acid RNAP RNA polymerase Rsd Regulator of sigma D SD Shine Dalgano sequence SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis sRNA small regulatory RNA tRNA transfer RNA UAS Upstream Activating Sites UTRs Untranslated regions

Papers included in the thesis

The thesis is based on the following published papers and a manuscript, which are referred to in roman numerals (I-III). Published papers are reproduced with permission from the journal publisher.

I. Madhushani, A., Del Peso-Santos, T., Moreno, R., Rojo, F. and Shingler, V. (2015), Transcriptional and translational control through the 5′-leader region of the dmpR master regulatory gene of phenol metabolism. Environmental Microbiology, 17: 119– 133. On line 2014-doi: 10.1111/1462-2920.12511

II. Moreno, R., Hernández-Arranz, S., La Rosa, R., Yuste, L., Madhushani, A., Shingler, V. and Rojo, F. (2015), The Crc and Hfq proteins of Pseudomonas putida cooperate in catabolite repression and formation of ribonucleic acid complexes with specific target motifs. Environmental Microbiology, 17: 105–118. On line 2014-doi: 10.1111/1462-2920.12499

III. Madhushani, A., Wirebrand, L. and Shingler, V. Multiple Hfq-Crc target sites are required to impose catabolite repression on (methyl)phenol metabolism in Pseudomonas putida CF600. Manuscript.

1. Introduction

1.1 Environmentally useful bacteria Many organic chemicals that are deliberately

“Microbes can and will do anything: introduced into the environment are specifically microbes are smarter, wiser and more designed to be beneficial in terms of pest and energetic than microbiologists, chemists, engineers and others” disease control. For example, in their use in enhancing crop production and a wide variety of David Perlmon (1980) Developments in Industrial Microbiology. Vol31 domestic and industrial applications that counteract

biofouling of different types – such as additives to

paints, lubricants and wood preservatives.

While beneficial at the site of introduction and/or in manufacturing, these same chemicals become environmentally damaging when they accumulate in soil and water ecosystems through runoff or are accidentally released in industrial effluents. Because of the stability of the basic aromatic (benzene) ring against microbial enzymatic attack and the low water solubility of most aromatic compounds, this class of chemicals has a particular propensity to accumulate (reviewed in 1, 2) and become sources of environmental pollution that adversely affects the well-being of wildlife as well as humans.

Both monetary costs and environmental considerations have prompted much research into microbial-based systems to safeguard the environment. These include, as just a few examples, alternative microbial solutions for crop protection and growth stimulation (3) as well as natural and engineered bacteria for pollution detection, clean-up purposes and green chemistry (e.g. 4, 5, 6). The remarkable biochemical diversity of soil and water dwelling microbes (bacteria and fungi) make them critical for all biogeochemical cycles. Their varied metabolic capacity includes the innate ability to degrade and recycle carbon from naturally generated aromatic compounds. Thus, through their voluntary efforts of consuming this class of compounds as food sources, native microbes can usually eventually renew most, but not all, sites polluted by aromatic compounds.

In addition to naturally occurring aromatics, microbes can also degrade the vast majority of man- made aromatics (so called xenobiotic); however, in these cases, degradation rates are generally slower or can be completely absent, leading to the use of expensive physicochemical treatment processes. Many factors contribute to inefficient biodegradation and hence environmental

1 problems caused by this class of chemicals (4, 5). Among these are inefficiency of pre-existing metabolic capacity to accommodate man-made compounds with unusual side groups on the aromatic ring, competition for limited available essential growth resources by other microbes, and abiotic factors that affect the ability of bacteria to access the growth substrates and/or perform their metabolic tasks (e.g. stress caused by toxicity of a polluting compound).

In their natural habitats, bacteria are exposed to constantly changing conditions. To accommodate such fluctuations, sophisticated specific and global regulatory networks have evolved to adapt gene expression patterns to optimally fulfil cellular demands. For example, bacteria with the ability to metabolize toxic aromatic pollutants possess specific regulatory circuits that respond to signals that report on the presence of the compound as an environmental growth substrate before producing the multiple enzymes required for their catabolism (5). These natural sensory systems have been co-opted for use in whole-cell biosensors as well as in genetic traps to isolate new metabolic enzymes, and in construction of synthetic pathways for bio- transformations (reviewed in 6, 7). However, not all carbon sources are metabolically equivalent. Natural microbial metabolic expertise and competitive fitness depends on global regulatory networks that ensure that, when presented with alternatives, growth substrates which provide the highest energetic gain are preferentially used (8).

Molecular level understanding of the metabolic strategies of bacteria in terms of potential exploitation has come from detailed studies of individual bacterial species. With respect to catabolism of aromatic compounds, toluene and xylene degradation by Pseudomonas putida mt- 2 (9) and phenol and methylated phenol degradation by P. putida CF600 (5) have been extensively used as prototypes to probe both specific and global regulatory systems that impact catabolism of this class of compound. While many advances have been made using these and many other bacterial species, much remains to be learnt about how bacteria integrate multiple environmental signals to optimise growth. Since these global regulatory inputs can lead to poor predictability and/or performance of regulatory circuits employed under the harsh conditions of polluted environments and bioreactors, efficient and sustainable exploitation of microbial-based technologies demands a clearer understanding of the mechanisms involved.

Bacteria can grow and proliferate almost everywhere. Irrespective of where they live, optimal energy generation is a key aspect for their adaption and survival. In this thesis, I describe recent advances in the understanding of how multiple regulatory inputs converge to control and modulate adaptive expression of the genes required for (methyl)phenol catabolism in 2

Pseudomonas putida. As a prelude to discussion of my own work, in the following sections I give a general overview of the strategies bacteria can use to control gene expression, with particular emphasis on those implicated in hierarchical control of phenol catabolism in P. putida strains.

1.2 Control of bacterial gene expression – from DNA to protein Bacterial genomes usually contain several thousand different genes. In addition to chromosomal DNA, self-replicating plasmids provide additional genetic information. Expression of protein- encoding genes occurs in two basic steps – transcription and translation – which provide major entry points for both specific and global regulatory control for adaptive gene expression (see Fig. 1).

Many bacterial genes are arranged in operons, in which a single promoter is responsible for the transcription of all the genes, providing one system for co-ordinate expression of multiple gene products involved in a given physiological process. In contrast to eukaryotes, bacterial transcription and translation occur simultaneously, since bacterial chromosomal DNA resides in a nucleoid that is not enclosed by a membrane and, thus, has access to cytoplasmic ribosomes. The transcribed messenger RNA (mRNA) is simultaneously used by ribosomes as the template for translation and synthesis of the cognate protein. This close coupling of transcription and translation events, alongside alterations in the structure of RNAs, leads to an impressive diversity of bacterial regulatory processes that can modulate the final levels of a gene product at both the transcriptional and/or the translational level. Some examples of these are detailed in later sections.

Transcriptional initiation, elongation and termination RNA stability and translation

Classical Global Regulators DNA-binding e.g. (p)ppGpp & DksA Riboswitches & Attenuators regulators sRNAs, RNA-binding & processing proteins DNA

Promoter engagement Anti--factor

 ribosome RNA Proteins

-factor competition & holoenzyme formation Figure 1. Regulatory factors that can control transcriptional output and translational efficiency in bacteria. Transcriptional regulation (left) can occur at any of the different steps of the transcription cycle – from promoter engagement, through initiation, elongation and termination, to association of -factors to generate the holoenzyme. As detailed in the text, regulation at this level can involve many different factors, including alterations in the structure of the RNA product through riboswitches and attenuation mechanisms. Similarly, RNA structure and its manipulation by interaction with sRNA or proteins underlies many regulatory processes that function through the stability of RNA products and/or translational efficiency (right) in the production of protein gene products. Adapted from (10).

Irrespective of whether the final gene product is a protein or a non-coding but functional RNA, such as a regulatory small RNA (sRNA), ribosomal RNA (rRNA), or transfer RNA (tRNA), all involve production of RNA complementary to the DNA template. This process is catalysed by DNA dependent RNA polymerase (RNAP), which drives transcription from promoters.

1.2.1 The bacterial transcription cycle RNAP is a complex molecular machine that undergoes numerous conformational changes as it binds and transcribes the DNA. RNA production involves three major steps: transcriptional initiation, elongation and termination. For transcriptional initiation, core RNAP requires association with an additional subunit – a σ-factor – to form the -RNAP holoenzyme before it can recognize and bind to the distinct DNA motifs of a promoter (11). In the initial stage of promoter engagement, RNAP forms a complex with closed, double stranded, DNA that is hence referred to as a closed complex. Both core RNAP and the -factor contribute to DNA melting to its open, single stranded form, in the vicinity of the +1 transcriptional start point just downstream of the promoter. Within this open complex, core RNAP initiates catalysis of the RNA from the template strand. When the transcript exceeds ~12 to 14 nucleotides in length, it starts exiting RNAP, causing conformational changes and concomitant breaking of some of the interactions

4 between the σ-factor and core RNAP, as well as σ-factor/promoter interactions (12). This event allows RNAP to leave the promoter (promoter escape) and proceed into the elongation phase. During elongation of the first ~100 to 200 nucleotides, the σ-factor is stochastically released and recycled to the pool of σ-factors available to associate with core RNAP (13). Transcription elongation continues until the core RNAP reaches a terminator comprised of a stem-loop hairpin structure within the RNA, which leads to dissociation of RNAP and the RNA from the DNA template, an event that sometimes requires the aid of the helicase activity of the Rho termination protein (14). At this point, core RNAP again becomes available for association with a σ-factor to restart the transcription cycle, see Fig. 1.

1.2.2 Players in transcription 1.2.2.1 The transcription machinery The catalytic core RNAP, with a molecular mass of ~400 kDa, is itself a multi-subunit enzyme consisting of five subunits: α2, β, β’, and ω (see Fig. 2). In contrast to the -RNAP holoenzyme, core RNAP can only bind DNA non-specifically, since it is the -factor that provides most of the promoter DNA recognition determinants (reviewed in 10, and 15).

The active site of RNAP is formed by the large β- and β′-subunits (16). The amino-terminal domains of the two α-subunits (αNTD) are involved in the assembly of the β- and β′-subunits. The carboxyl-terminal domains (αCTD) are separated from the αNTD by a flexible linker and can bind to A+T rich DNA UP-motifs, which lie upstream of the major promoter recognition elements, to increase both basal and activated transcription from certain promoters (17). The small ω-subunit was initially identified as an assembly factor for RNAP and, subsequently, as an important component for modulating the activity of RNAP in response to some regulatory molecules (18, 19, 20, see section 1.2.3.5).

aNTD aNTD b/b’ aCTD ω 

UP -35 -10 +1 TTGACA TATAAT Figure 2. Subunits of RNAP and promoter DNA binding.

Schematic illustration of the different subunits of the RNAP holoenzyme (α2, β, β’, ω and σ) and its interaction with DNA elements of a promoter. The consensus sequence of the major recognition determinants of the σ70-factor are also shown. The box adjacent to the -10 element marks the location of an additional recognition sequence present in some promoters, which is alternatively termed an extended - 10 or a -15 element. Adapted from (16).

1.2.2.2 Specificity (σ) factors As alluded to in the preceding sections, the dissociable σ-factor both facilitates unwinding of the DNA duplex near the transcript start site and builds promoter binding specificity into the holoenzyme. Most free-living bacteria contain more than one σ-factor, with the multiplicity of alternative σ-factors generally reflecting the ability to thrive in more diverse environments (16). For example, enteric Escherichia coli has seven different σ-factors whilst the soil, water and root dwelling P. putida has twenty-four (21). Hence, a single species of core RNAP is used to generate many alternative forms of the holoenzyme, see Fig. 1.

Based on sequence and functional similarity, alternative σ-factors are categorized into either the σ70-family or the σ54-family, named after their counterparts in E. coli (reviewed in 10). Each σ- factor programs RNAP with a distinct promoter binding specificity, and thereby directs RNA polymerase to the different classes of promoters in the bacterial genome. For example, σ70 programs RNAP to recognise promoters with the consensus elements TTGACA‐N17‐TATAAT, which are usually located ~ ‐35 and ‐10 with respect to the transcription start site (Fig. 2, 22), while the promoter recognition elements for σ54 are located at ‐24 and ‐12 relative to the transcription start site and have a very different consensus recognition sequence: TGGCAC‐N5‐ TTGCa/t (23).

The broad array of alternative σ-factors allows for co-ordinate regulation of transcription from different classes of promoters involved in responding to different needs (10, 15). The housekeeping or household σ-factor (σ70 in E. coli and P. putida; also known as RpoD), is the most abundant σ-factor and is responsible for the bulk of transcription of essential genes during rapid growth. Other σ70-family members are less abundant and control adaptive transcriptional responses. For example, σ38 (RpoS), a general stress response σ-factor used in counteracting nutritional and other stresses (24), σ24 (RpoE) and σ32 (RpoH) for counteracting heat and other physico-chemical stress imposed by the environments that bacteria inhabit. Yet other σ-factors, e.g. the σ70-family member σ28 (FliA) and σ54 (RpoN), are required for expression of flagella and motility responses (15). In addition to motility, σ54 is involved in the expression of many other genes, including those for uptake and utilization of alternative carbon sources and those involved in nitrogen metabolism (25, 26).

1.2.3 Regulation of transcription Under this section I provide a brief overview of the regulation of the first steps of transcription – initiation and elongation. 1.2.3.1 Holoenzyme composition A primary checkpoint in transcription from any bacterial promoter is the level of the cognate σ- RNAP holoenzyme required for promoter engagement. Hence, changes in the composition of the holoenzyme pool determine the extent to which different promoter classes can be occupied. Different σ-factors compete for limiting numbers of core RNAP molecules (see schematically represented in Fig. 1). The levels, activities and availability of different σ-factors is constantly adjusted in response to environmental cues (reviewed in 10). This is achieved by multiple tiers of regulatory mechanisms involving transcriptional and translational control, as well as post- translational processing via proteases, and sequestering by anti-sigma factors.

Sequestering of a σ-factor by a cognate anti-sigma factors until its activity is needed provides a means of rapidly adjusting the composition of the holoenzyme pool, and thus promoter output, without de novo protein synthesis. The Rsd protein (regulator of sigma D) has also been implicated in controlling the levels of alternative holoenzymes via reducing the availability of 70 (10, 27). By forming 1:1 complexes with 70, the naturally elevated levels of Rsd in stationary phase E. coli are sufficient to sequester ~25% of the relatively constant levels of 70 (28), which would predictably facilitate access of alternative σ-factors to core RNAP. However, the precise mechanism(s) underlying the action of Rsd and when it is most effective are still unknown (27).

Changes in the composition of the σ-RNAP holoenzyme pool provide the background in which promoter activity can be further adjusted by DNA-binding transcriptional regulators (repressors and activators) and global regulatory molecules that directly targeting the RNAP to modulate its activity.

1.2.3.2 Classical DNA-binding transcriptional regulators Classic bacterial transcriptional repressors and activators bind to control regions (operators), adjacent to or overlapping promoter DNA motifs, to manipulate – either negatively or positively – one or more of the steps of transcriptional initiation pathway of RNAP. While the activity of some promoters can change dramatically in the apparent absence of any DNA-binding transcriptional regulator, most bacterial promoters respond to different environmental and internal signals with the aid of at least two (reviewed in 5, and 16). 7

Signal-responsiveness can be built into promoter control via many different mechanisms, which ultimately determine either the production of the regulators or their activities (29). Classic examples of direct signal-responsive mechanisms come from trp and lac operons of E. coli in which the DNA-binding affinity of the cognate regulator is altered in response to ligands that report on nutrient availability. In the case of the TrpR repressor for tryptophan biosynthesis, binding of tryptophan unlocks its DNA binding capacity, resulting in repression of transcription when the operon gene products are not needed. Conversely, for the catabolic lac operon, LacI binding of allolactose (that signals the presence of lactose in the growth medium), results in blocking of its DNA-binding ability and de-repression of the system.

The lac operon also provides a ready example of where two transcription regulators are involved in integrating a specific signal and a global metabolic signal to subvert expression of gene products until they are needed. In addition to de-repression caused by LacI repressor binding of allolactose, transcription from the lac promoter also requires activation to compensate for its sub-optimal σ70 promoter sequence. This activation is mediated by the global regulatory protein CAP (catabolite activator protein) that helps recruitment of σ70-RNAP to the promoter. CAP is the major mediator of global catabolic control that determines the hierarchical utilization of sugars as growth substrates in E. coli.

This global level of control is imposed by the requirement of monomeric CAP to bind cyclic- AMP (c-AMP) to take up its active DNA-binding dimeric form. In E. coli, cellular levels of c- AMP are ultimately controlled through the uptake of glucose – the preferred carbon source – which is actively transported into the cell. The phosphoenol pyruvate carbohydrate phosphotransferase system (PTS) couples glucose transport to its phosphorylation. When glucose levels are high, the PTS system inhibits active transport of other less preferred sugars through interaction between EIIAGlc component and their specific transporters (inducer exclusion). Under these metabolic conditions the levels of cAMP are low. However, when glucose levels are low or exhausted, EIIAGlc becomes phosphorylated, preventing its inhibitory interaction with transporters. Instead, the phosphorylated form of EIIAGlc, along with other factors stimulates the activity of the adenylate cyclase [which generates cAMP from ATP], to result in high c-AMP levels and consequent high cAMP-CAP dependent transcriptional activation (reviewed in 8).

As illustrated by the example above, not all signal sensing by transcriptional regulators is direct. Bacterial surveillance of their environment often involves conversion of external signals into 8 either changes in the concentration of a nucleotide secondary-messenger (e.g. c-AMP, c-di-GMP and ppGpp) that functions as global regulatory molecule with multiple targets, or a phosphorylation event to couple a signal to promoter output (5, 29). In these regulatory circuits, initial sensing by one or more proteins is converted to control of the activity of the transcriptional regulator. This is the case in the major class of response regulators of bacterial two-component systems. Two components systems employ sensory histidine kinases that autophosphorylate at a histidine residue in response to a signal. The activity of the cognate response regulator is determined by its phosphorylation status, which is controlled by phospotransfer of the high energy phosphoryl group from the histidine kinase to an aspartate residue of the response regulator. The basic scheme of two-component systems is extremely adaptable and is used in signal integration in a wide variety of physiological processes, including metabolic control (reviewed in 30).

1.2.3.3 bEBPs – specialized mechano-transcriptional activators of σ54 Although some promoters need an activator to recruit or stimulate the activity of RNAP, σ70- RNAP and other holoenzymes using σ70-like factors are intrinsically competent to initiate transcription on their own. This is not the case for σ54-RNAP, which forms thermodynamically stable closed complexes at the promoters it controls. Hence, σ54-promoters always strictly require activation by a member of the specialized family of bacterial enhancer binding proteins (bEBPs), which utilize ATP hydrolysis to activate σ54-dependent transcription.

The bEBP-family of transcriptional activators derives its name from the location of its DNA binding sites and mode of action. In contrast to classical activators, bEBPs bind to specific DNA sites far (more than 100 bp) from the promoter they control, and in some cases these sites can be moved up several kb away and still function, albeit inefficiently, in correct promoter control (25, 31). The DNA binding sites for inactive bEBPs dimers are comprised of a long imperfect inverted repeat termed a UAS (upstream activating sequences). As the name implies, these sites are usually located upstream of the cognate promoter; however, there are exceptions where the bEBP binds to downstream UASs to control σ54-dependent transcription, e.g. P. aeruginosa FleQ at the flhA, fliE, and fliL promoters, and Bacillus subtilis RocR at the rocG promoter (32, 33). From these distal sites, the active multimeric forms of bEBP acts on pre-bound σ54-RNAP, as illustrated in Fig. 3, to activate transcription through by a common mechanism.

54-RNAP 54-RNAP Closed complex Open complex

IHF

bEBP dimer bEBP activation ADP + Pi ATP

Figure 3. Transcriptional initiation by 54-RNAP. Schematic illustration of the sequential steps of activation of σ54-RNAP by bEBPs. The bEBP activators act on pre-bound σ54-holoenzyme that is locked in a closed-complex. Multimerization of the bEBP to the transcriptional-promoting active form requires a bEBP-specific signal. Subsequent ATP hydrolysis by the bEBP promotes open complex formation with the aid of re-oriented GAFTGA-loop. Alternative configurations of the GAFTGA-loop, masked (black) or exposed (red), at different stages is indicated. Figure adapted from (26). bEBPs usually exhibit a domain structure with individual domain performing distinct functions: a signal reception domain at the N-terminal region, a helix-turn-helix DNA binding domain in the C-terminal region, and central catalytic domain for interaction and activation of σ54-RNAP. The signal reception domains usually determines the transcriptional promoting property of these proteins by controlling ATP-triggered multimerisation in response to an environmental signal that cues the need for its activity (26, 34). These can be ligands, as is the case for P. putida derived DmpR and XylR that control catabolism of aromatics, phosphorylation events [e.g. of NtrC from enteric bacteria involved in nitrogen assimilation and DctD from Sinorhizobium meliloti controlling C4-dicarboxylic acid transport], or removal of inhibitory protein:protein interactions, as exemplified by Azotobacter vinelandii NifA that requires release from a complex with the redox-responsive NifL protein before it can play its role in nitrogen assimilation.

Although natural variants lacking a sensory domain and/or DNA binding domain exist, all bEBPs possess a central catalytic AAA+ domain [ATPases associated with various cellular activities]. This domain also contains the signature GAFTGA motif of bEBP on a flexible loop – a key feature involved in converting the energy derived from ATP hydrolysis into a mechanical force exerted on σ54-RNAP to drive conformational changes essential for the transition from a closed complex to an open complex (reviewed in 26). The N-terminal sub-region I of σ54 is primarily responsible for maintaining σ54-RNAP in a close complex through its initial interactions with -12 promoter element. These interactions off-set σ54-RNAP such that loading of promoter DNA into the active site cleft is blocked. Upon ATP hydrolysis by the bEBP, conformational changes expose and stabilize the GAFTGA-loop to engage region I of σ54,

10 leading to relief of its inhibitory conformation and subsequent correct promoter DNA-σ54 alignment for open-complex formation (Fig. 3).

As schematically illustrated in Fig. 3, correct interfacing of bEBPs and σ54-RNAP requires DNA looping between the distally bound bEBP and promoter bound σ54-RNAP. This process can be assisted by either intrinsic curvature in the intervening DNA [e.g. B. subtilis RocR at the rocABC promoter (33)] or by a site specific DNA bending protein such as integration host factor (IHF) that acts as a co-regulator at many σ54-promoters (26).

1.2.3.4 Global co-regulation by architectural proteins Dedicated transcription regulators are generally present in low abundance and their action can be modified by co-regulatory nucleoid-associated proteins that are present in comparatively high abundance. These nucleoid-associated proteins, such as integration host factor (IHF), factor for inversion stimulation (Fis), and histone-like nucleoid-structuring proteins (HNS and HU), all modify the architecture of the regulatory regions of many different promoters (31, 35) and are, therefore, considered as global regulators. Although generally present in large numbers, the levels of nucleoid-associated proteins are also controlled in response to environmental cues (35), adding another level at which signals can be interpreted to control promoter output.

One of the major mechanisms of co-regulation by this class of proteins involves facilitating or block interactions of other regulatory proteins through modifying the location of the DNA in three-dimensional space (31). For example, in addition to its role in compacting the nucleoid, HNS and HNS-like proteins act as repressors that can completely silence gene expression by forming bridging structures between distal DNA sites and/or extended nucleoprotein structures that occlude binding of other regulatory molecules. IHF, on the other hand, induces sharp DNA bends of >1600 that, depending on the location of its binding site, can simulate or inhibit regulatory protein:protein interactions at a wide variety of promoters.

In addition, IHF can contribute to transcriptional regulation by other means. These include: i) IHF-mediated architectural changes that aid binding of regulators at specific promoters, ii) IHF- aided changes in DNA topology that allows the simultaneous binding of the two α-subunits to a distally located UP-element that would otherwise be out of reach, iii) restricting transcriptional activation specificity to a single bEBP by forming structures that block undesirable and nonspecific activation by bEBPs bound to nonspecific binding sites, or from solution, and iv) IHF- mediated effects on DNA supercoiling that can stimulate open-complex formation and/or 11 stability of σ54-RNAP (26). Intracellular levels of IHF in E.coli and P. putida become elevated as cells enter stationary phase and stimulate output from many σ54-promoters. This increase in IHF levels is primarily mediated by the second messenger (p)ppGpp (reviewed in 26).

1.2.3.5 Transcriptional control through RNAP binding to (p)ppGpp Under energy limiting and stress conditions, bacteria enter into the stationary phase with reduced growth rates. A major player in this process is the stringent response mediated by guanosine tetraphosphate (ppGpp) and guanosine pentaphosphate (pppGpp) – collectively known as (p)ppGpp. These nucleotide “alarmones” help the cells balance translational capacity to the reduced demand in protein synthesis, and generally allow the cell to re-orchestrate transcription towards functions required for survival rather than proliferation. While different bacteria accomplish this by different routes (20), in E. coli and P. putida, (p)ppGpp directly targets RNAP to alter its properties at kinetically sensitive promoters dependent on σ70 and σ70-like proteins e.g. σ24/E, σ28/FliA, σ32/H and σ38/S in vitro (reviewed in 10).

Recently, the elusive binding site for (p)ppGpp on E. coli RNAP has been identified as a cleft formed between the α, β and ω subunits, located far (~30 Å) away from the active site (reviewed in 20). DksA, a transcription factor that amplifies the effects of (p)ppGpp, binds to the opposite side of RNAP and inserts its coiled-coiled domain through the secondary channel to access the active site (36). Synergistic effects of these two regulatory factors are proposed to result from DksA-mediated widening of the secondary channel and long-range structural changes, although the precise repercussions of these changes still need to be elucidated (20).

Direct targeting of RNAP by (p)ppGpp and DksA can account for much of their action in down- regulating transcription from powerful σ70-dependent promoters such as tRNA and rRNA operon promoters and up-regulation of transcription from promoters that control factors that are needed for adaptation and survival (26). However, this cannot account for all the transcriptional consequences of these regulatory molecules in intact cells. For example, neither (p)ppGpp nor DksA have any effect on σ54-dependent transcription in vitro, while lack of these regulatory molecules essentially abolishes σ54-dependent transcription in E. coli and P. putida (37, 38). The in vivo amplification of (p)ppGpp and DksA effects have been attributed to passive and active modulation of -factor competition for core RNAP to result in elevated levels of holoenzymes using alternative -factors (reviewed in 10, and 39). Since there is no known anti-sigma factor for σ54, and the levels of σ54 are constant in E. coli and P. putida under different growth

12 conditions, such regulation appears to be the primarily mode of controlling the functional pool of the σ54-RNAP holoenzyme (37, 38, 40).

1.2.3.6 Control of transcriptional elongation through RNA switches – attenuators and riboswitches 5’-leader regions of mRNA can also serve as regulatory targets that control transcriptional elongation to fine tune signal responsive gene expression. Both attenuators and riboswitches provide fast and sensitive mechanisms to terminate elongation by RNAP until gene products are required. These regulatory mechanisms involve cis acting elements involved in sensing, which in turn, determine the formation of alternative stem-loop structures that either allow progression of RNAP or induce its detachment, leading to premature termination (41).

These attenuation systems are associated with a wide range of sensing mechanisms. One major class involves translation of a short upstream open reading frame (uORF), and includes the first attenuation system proposed by Charles Yanofsky and coworkers (42) to account for premature transcriptional termination of the polycistronic trp mRNA in the presence of high level of tryptophan. Variations on the basic theme were subsequently discovered in other amino acid biosynthetic operons. This class of attenuator – generally referred to as “peptide leader” attenuators – involve sensing through translation of uORFs enriched in codons for the amino acid synthesized by the enzymes encoded by downstream genes. High levels of the cognate charged tRNA allow swift ribosomal reading of the uORF, resulting in formation of the transcriptional termination structure. Conversely, pausing of ribosomes due to lack of the cognate charged tRNA (signalling the need for the gene products) allows formation of an alternative anti-termination structure and transcriptional progression.

Riboswitches comprise another major class of RNA regulatory elements that can attenuate transcription and similarly work through alternative secondary structures. In these cases, they are primarily located within 5’-untanslated regions (5’-UTRs) and commonly bind specific metabolites using complex three-dimensional aptamers to either promote or inhibit transcription. Such metabolites can include certain amino acids, nucleosides and sugars. Other analogous RNA-based switches operate in a similar way through binding of a second messenger (e.g. c-di- GMP), metal ions (Mg2+), an uncharged tRNA [e.g. as in so called T-Box switches], or a protein [e.g. some ribosomal proteins within their cognate leader regions] (41, 43).

Recently, riboswitches have also been found in non-coding sRNAs that bind and sequester transcriptional activators required for ethanolamine utilization. These sRNAs – EutX of Enterococcus faecalis and Rli55 of Listeria monocytogenes – become truncated when they bind factors required for enzyme activity, and thus provide systems that couple expression of a metabolic pathway to the presence of an essential co-factor (44, 45).

1.2.4 Translation Translation, like transcription, consists of stages of initiation, elongation and termination, which can all be played upon to exert regulatory effects (e.g. inhibition of ribosome assembly on the mRNA, translational frame shifting and ribosomal pausing, 46 and references therein). The 70S bacterial ribosome is a riboprotein machine, consisting of a small 30S subunit (primarily involved in initiation and reading of the mRNA) and a large 50S subunit (that catalyses peptide bond formation). Initiation of translation involves two key steps that are aided by initiation factors. First, the mRNA and the 30S ribosomal subunit form a pre-initiation complex with tRNAfMet. Pairing between the Shine-Dalgarno (SD) sequence – a 4 to 6 nucleotide conserved region within the mRNA – and a complementary region of the 16S rRNA of the 30S subunit, stabilises the pre-initiation complex and allows correctly positioning of the initation codon (usually AUG) for incorporation of the first amino acid. The second step is a slow, essentially irreversible conformational change, which facilitates pairing between the initiation codon and the anti-codon region of tRNAfMet to form the stable initiation complex. After the dissociation of the initiation factors, the 50S subunit associates to form the assembled 70S ribosome ready to proceed into the elongation phase. Polypeptide chain elongation continues until the ribosome reaches a stop codon. At this point, translation terminates in the presence of release factors, which trigger dissociation of the newly synthesised polypeptide from the ribosome.

1.2.4.1 RNA structure and translational regulation In addition to secondary structures within their 5’-leader regions, mRNAs also contain secondary structures within their downstream coding sequences. The ribosome itself possesses a helicase activity that can often disrupts downstream structures in mRNAs during elongation; however, this property of ribosome is not functional in translational initiation, making this step of translation particularly vulnerable to regulation through RNA structure (47).

Regulation of translational initiation in bacteria is often exerted by secondary structures within the mRNA that undergo changes that alternately sequester or expose the SD and/or AUG start codon – the two essential elements of the ribosome binding site (RBS). Some of these do not 14 require any trans-acting factor for environmental signalling. For example, RNA thermometers, such as those of the mRNAs encoding the transcriptional activator PrfA in L. monocytogenes and the heat-shock -factor RpoH in E. coli, influence translational efficiency by controlling access of ribosomes to the SD in a temperature dependent manner (48). Translational riboswitches can also function in an analogous manner – e.g. vitamin B1, which through binding to a conserved RNA aptamers, can inhibit several mRNAs encoding enzymes for its own synthesis (49).

1.2.4.2 Antisense sRNA mediated translational control Antisense sRNAs play upon the mRNA through base pairing to provide very rapid adaptive responses. These interaction usually involve discreet motifs presented by one or more loops on the sRNA (50, 51). As simplistically illustrated in Fig. 4, pairing with the mRNA can modify translation in different ways, including direct translational repression, by blocking the initiation site (either the SD and/or the AUG initiation codon). Conversely, they can directly promote translational initiation by precluding inhibitory secondary structure of mRNA that would otherwise sequester the SD and/or the AUG in step loop structures (46 and references therein). A more indirect way they can act is by interacting within the coding regions of uORFs, to either i) impede the passage of the ribosome to a downstream RBS and thereby disrupt translational coupling, or ii) through disruption of inhibitory intra-molecular structure of the mRNA, as has been found in the case of the sRNA PhrS (52). Ribosome occupancy in these case can increase or decrease the stability of the target mRNA as a secondary effect of sRNA mediated regulation. Other antisense sRNAs target mRNAs for destruction without interfering with initiation of translation (53).

No sRNA sRNA present

5’

5’ ORF 3’ 3’ activation 5’ ORF

Translational 3’

ORF 3’ ORF 3’ repression

Translational 5’ 5’ 5’ 3’

5’ 3’ uORF 5’ 5’ uORF

3’ ORF coupling

translational 3’ Facilitation of Facilitation

5’ 3’

5’ uORF uORF

5’

coupling translational Disruption of Disruption 3’ 3’

Figure 4. Translational control by sRNA Schematic illustration of different pairing mechanisms between sRNA (blue) and the target mRNA (red) for control of translational efficiency. Upper: direct control, either by disrupting inhibitory secondary structures for translational activation or occluding ribosome access for translational repression. Lower: indirect control, by facilitating or disrupting translational coupling.

The rate-limiting step in sRNA–mRNA pairing is the initial base-pairing interaction. The presence of a U-turn loop structural motif is a general binding-rate-enhancer that promotes RNA–RNA pairing (reviewed in 51). However, because antisense sRNA–mRNA pairing involves short and discontinuous regions of complementarity, it frequently requires an additional

16 factor to ensure the specificity and magnitude of the response over an appropriate time scale. Thus far, the only factor described to fulfil this function is the RNA chaperone Hfq (54).

The signal-responsive regulatory functions of antisense and other sRNAs results from transcriptional control of their production in response to environmental cues, such as nutritional stress (SgrS), iron limitations (RyhB), membrane stress (MicA) (reviewed in 54). Antisense RNAs usually have a single complementary target RNA within the genome. However, some can have multiples targets that, through their action on other global regulators, cause cascades of regulation e.g. E. coli DsrA that promotes translation of rpoS but negatively influences HNS (46, 55 and references therein).

1.2.4.3 RNA-binding proteins and translational control Rather than sRNA–mRNA pairing mechanisms, other sRNAs are involved in translational control through possession of multiple copies of a target site for a regulatory RNA-binding protein. This molecular mimicry allows sRNA of this class to function as “sponges” that mop up the regulatory protein by sequestration and thereby prevent its action on target mRNAs. This is the case for the global regulatory RNA-binding carbon storage protein CsrA, whose activity is counteracted by the sRNAs CsrB and CsrC (56). However, most RNA-binding proteins function without the involvement of small RNAs (46).

Although examples of RNA-binding proteins that promote translation exist, most serve as repressors of translational initiation. Many of these translational repressors use a displacement strategy involving direct binding across the RBS [or in its close proximity] to sterically impede access of ribosomes (reviewed in 46). The same end can be achieved by an alternative mechanism in which the repressor protein binds far upstream from the RBS in such a way that it facilitates mRNA refolding to a conformation that sequesters the SD in a RNA hairpin to blocks ribosome entry. Such a mechanism has been proposed for additional regulation of tryptophan biosynthesis in B. subtilis in response to tryptophan (57). In addition to these competition strategies, entrapment mechanisms that also prevent translational initiation have been proposed. For example, as in translational repression of the E. coli α-operon by ribosomal protein S4, which traps the 30S subunit in a dead-end complex that does not allow tRNAfMet to pair with the AUG (58).

1.2.5 The metabolically versatile pseudomonads Pseudomonas species are short rod shaped Gram-negative gamma proteobacteria that can occupy many environments due to their adaptive metabolism. Their diverse metabolic capacity is reflected in the large sizes of their genomes (http://www.pseudomonas.com) and underscores much of their medical, agricultural and biotechnological importance. For example, ubiquitous P. aeruginosa strains are major opportunistic pathogens of cystic fibrosis and burn-wound patients, while some P. syringae and P. fluorescens strains damage crops and are frequent agents of food spoilage. P. putida strains, on the other hand, are renowned for their ability to degrade organic compounds, such as aromatic (e.g. toluenes and phenols) and aliphatic compounds (e.g. alkanes and styrene), which serve as stress agents as well as growth substrates (5).

The soil and root colonizing model organism P. putida KT2440 is a long standing workhorse for biotechnological applications because it lacks virulence factors (59) and was the first organism approved for environmental release (60). With its genome size of >6 million bp, P. putida KT2440 comes equipped with ~20% of its genome committed to regulatory functions, and a large proportion of the remainder dedicated to transport and metabolism of carbon, and efflux of otherwise toxic organic compounds (59). Since P. putida strains are evolutionarily adapted with metabolic and stress-endurance traits, they represent ideal strains for many biotechnological purposes that involve chemical transitions and intermediates that would kill bacteria such as E. coli (6).

1.2.5.1 The major CRC mechanism of P. putida Pathways to degrade aromatic and other seemly exotic sources of carbon, generate intermediates that feed into central metabolism. These auxiliary pathways employ whole sets of specialized catabolic enzymes, which are metabolically expensive to produce. Therefore, the ability to assimilate such compounds is nonessential, but does provides a metabolic advantage when other more energetically favourable carbon sources are absent. Consequently, expression of these auxiliary pathways is usually controlled specifically in response to substrate availability, while being subservient to global regulatory networks that serve to optimize the energy status of the bacterium (5).

In contrast to E. coli with its preference for glucose as a carbon source (section 1.2.3.2), pseudomonads preferentially utilizes some amino acids and organic acids (61, 62). Hierarchal use of these compounds and their preferred utilization over other sources of carbon has long been

18 connected with the global regulatory protein Crc, which acts at the post transcriptional level to mediate carbon repression control (CRC; reviewed in 8).

The levels of the Crc protein do not vary much over the growth curve or under different growth conditions – rather the action of Crc is antagonised by sRNAs – e.g. CrcZ and CrcY in P. putida and CrcZ in P. aeruginosa (61, 63). These sRNAs contain multiple copies of a “catabolite activity motif” found in mRNAs targeted for translation repression by Crc and, thus, possess the features of the sRNA class that function as target mimics to sequester RNA-binding proteins (section 1.2.4.3). Production of CrcZ and CrcY is controlled by their 54-dependent promoters through the CbrA/CbrB two-component system (61, 63, 64, 65). Although the precise signal that activates the CbrA/CbrB system is unknown, the levels of CrcY and CrcZ become elevated under nutrient limiting conditions to alleviate CRC responses mediated through the Crc protein.

Until recently, Crc was thought to bind directly to the Catabolite Activity (CA) motifs of its target mRNAs to subvert catabolism of non-preferred carbon sources, and to those of CrcZ and CrcY to provide relief of Crc-mediated repression when other preferred sources of carbon become limited. However, as detailed in the Results and Discussion section of my thesis, the publication in 2013 that P. aeruginosa Crc is unable to bind RNA (66) instigated a need for reassessment of how this regulatory circuit functions. Resolving this question identified the RNA chaperone protein Hfq as an additional and critical component of CRC in P. putida. With respect to phenol catabolism, these experiments used components of the dmp (dimethylphenol) system derived from P. putida CF600.

1.2.5.2 P. putida CF600 and the dmp-system P. putida CF600 was originally isolated from coke liquor waste through its ability to grow at the expense of 3,4-dimethylphenol, and subsequently found to catabolise phenol and mono methylated phenols (cresols). This (methyl)phenol degradative ability is conferred by the dmp- system harboured on the self-transmissible Inc-P2 mega plasmid pVI150 (67). The dmp-system is composed of a regulatory gene, dmpR, and the dmp-operon, which contains fifteen structural genes (dmpKLMNOPQBCDEFGHI) for a multicomponent phenol hydroxylase and a meta-cleavage pathway for degradation of (methyl)phenols to central metabolites (68).

As schematically illustrated in Fig. 5, the dmp-system is controlled by two non-overlapping promoters dependent on different -factors: namely 70-Pr and 54-Po. These promoters are

19 located within a 406 bp intergenic region and drive divergent transcription of dmpR [from 70- Pr] and the dmp-operon [from 54-Po] (38, 69 and references therein).

Figure 5. The dmp-system of P. putida CF600 Schematic representation of the P. putida CF600 dmp system (not to scale). The locations of the dmpR gene with its 5’-leader region, the binding site for DmpR- (UAS2 & UAS1, inverted arrows), and the IHF binding site, are shown relative to the dmp-operon. A model of aromatic-effector activation of DmpR and a consequence feed-forward loop (cyan stars) mediated through interplay of the Po and Pr promoters and the effects of (p)ppGpp/DksA (black star) are indicated. Adapted from (38), see text for details.

Specific regulation of this specialized catabolic pathway is achieved through the aromatic sensor-regulator DmpR – a 63.3 kDa protein that responses to the availability of phenolic compounds in the growth medium. DmpR is the master regulator of this system because it is the obligatory bEBP required for transcription from the 54-Po promoter. DmpR only takes up its transcription-promoting active form upon direct binding of phenolic compounds, which allows ATP-binding triggered multimerisation (Fig. 5; 70, 71). This aromatic effector-activated form of DmpR bind to its UASs sites located approximately 170 bp of the Po promoter. Binding of IHF to its DNA binding site, located between the UASs and 54-Po, facilitates the productive interaction between the Po-bound σ54-RNAP and active DmpR bound to the UASs (72). Hence, DmpR through limiting the activity of the Po promoter to conditions when pathway substrates are present, ultimately control the levels of the Dmp-enzymes and the (methyl)phenol degradative capacity of the bacterium.

Previous work has established that the dmp-system is intricately integrated into the (p)ppGpp / DksA network at the transcriptional level, to couple Po promoter output to low energy conditions when (methyl)phenol catabolism would be advantageous. DmpR levels are directly enhanced through the action of (p)ppGpp / DksA on 70-RNAP at the intrinsically weak Pr promoter,

20 resulting in increased binding of 70-RNAP and an accelerating the rate of open-complex formation (38, 69). More indirect input comes from the stimulatory effects of these molecules on the levels of IHF and the functional pools of 54-RNAP that are also required for 54-Po activity (37, 38, 40, 73). As a final twist, activity of the 54-Po promoter stimulates activity of the divergent 70-Pr promoter, resulting in a feed-forward loop whereby DmpR stimulates its own synthesis (38). Since (p)ppGpp levels are elevated when the cells undergo nutrient and other stresses, expression of DmpR is maximised at the transcriptional level during such condition when substrates for the pathway are present.

2. Aims

As I entered the world of regulation of (methyl)phenol catabolism by P. putida CF600, two levels of hierarchical regulation were well established – the absolute requirement of an aromatic effector molecule to activate DmpR and input of the (p)ppGpp / DksA transcriptional network. Both these converge to target the master regulator DmpR. At that time, a 5’-leader region of the dmpR gene was also known to alter the levels of DmpR by some unknown mechanism, suggesting other potential levels of control. Thus, the overall objective of my Ph.D. work was to determine the role of this 5’-leader region. As the research progressed, the following specific questions came into focus:

I] Does the DNA of the 5’-leader region affect the levels of dmpR mRNA? If so, how?

II] Are there regulatory elements within the 5’-leader region of the dmpR mRNA that alter translation?

III] How does the Crc post-transcriptional repressor fit into the picture?

IV] What additional factor does Crc require to mediate carbon catabolite repression?

V] Is Crc-mediated translational repression of dmpR all that is needed for catabolite repression of the dmp-system?

3. Results and Discussion

3.1 A promoter-distal ATAAATA DNA motif reduces the levels of mRNA transcripts The initially transcribed region of the dmpR gene encodes a 123 nt long 5’-leader region (5’-LR) of the dmpR mRNA (Fig. 6A). DNA within this region was found to mediate a reduction in the number of transcripts arising from the 70-dependent Pr promoter. This repressive effect was first detected using P. putida KT2440 harbouring a series of luciferase transcriptional reporter plasmids in which the Pr promoter and different portions of the dmpR gene were placed in front of the RBS of the bacterial luxAB genes. This analysis traced the repressive effect to the promoter-distal region of the 5’-LR DNA (Paper 1, Fig. 1).

As summarised in Fig. 6B, in close agreement with luciferase activity assays, quantitative PCR (using primers directed to the luxAB genes), revealed the presence of the 5’-LR DNA caused a 5- to 7-fold reduced number of transcripts in the stationary phase, and that this reduced number was not caused by an effect through mRNA stability.

A. 70-Pr

-35 -10 +1 TGGTTGACTTTTGCCAGATACTGAGGCTGGCTATGGGGAGCTGGCGCAGGTGAAAAAACTGCCGATTTTCCCCATGACCCCA TCTGGAATCGCCGCCTGCCTTGCGCTATAGCGGCGACCCTGATTTTCCCCATCTAAAAATAAATAGGGGCCTCGCTTACATG

B. 8 Luciferase Quantitative mRNA stability assays 8 PCR 7 100 Pr 7 6 6 80 5 + 5’-LR 5

set as 100%) - 5’-LR

0 60 4 4 3 3 40

2 2 T = Relative Transcripts Transcription from Transcription 1/2 20

1 1 ~5 min Transcripts (T Transcripts

0 0 0 5’-LR: + - 5’-LR: + - 0 2 4 6 8 10 12 14 16 Time (mins)

Figure 6. Inhibitory effect of the 5’-LR on the number of transcripts A. The sequence of the initially transcribed DNA that encodes the 123 nt 5’-LR of the dmpR mRNA (highlighted in cyan) is shown with the -35 and -10 elements of σ70-Pr promoter (74) underlined. Matches to the consensus elements of σ70-promoters are in bold, while the ATG initiation codon of dmpR is shown in bold italics. B. From left to right, the first two graphs show the luciferase activity and quantitative PCR results obtained from stationary phase LB cultures of P. putida KT2440 harbouring either a Pr-luxAB transcriptional reporter plasmid with the 5′-LR region (filled bars) or an equivalent reporter lacking the 5′- LR (open bars). The graph to the right shows the mRNA stabilities of luxAB transcripts produced from Pr reporter plasmids with (filled symbols) or without (open symbols) the 5′-LR. Time point 0 was set as 100% transcripts. 24

A similar series of Pr-luxAB transcriptional reporter plasmids, with different manipulations of the 5’-LR DNA, were used to further trace the DNA element responsible for this effect (Paper I, Fig. 2). As summarised in Fig. 7, the 5’-LR DNA could mediate its effect when placed in either orientation (compare 1 & 2 with 3 & 4 in Fig. 7), but could not be mimicked by unrelated random DNA (compare 1 with 5), attesting to its specificity. Further deletions and manipulations identified an ATAAATA motif as the DNA element that results in the reduction of transcripts arising from the Pr promoter (see 9 & 11 in Fig. 7).

70-Pr

Luciferase plate test assays +123 +1 5’-LR luxAB genes 1 1 2 7 1 + 5’-LR native 2 2 - 5’-LR 5 3 + 5’-LR reconstructed 11 8 4 +5’-LR inverted 4 3 5 + random DNA 10 9 6 + 3 5’-LR 7 + 4 5’-LR…. CATCTAAAAATAAATAGGGGCCTCGCTTACATG repressive 8 + 5 5’-LR…. AGGGGCCTCGCTTACATG non-repressive 9 + 6 5’-LR…. CATCTAAAAATAAATATG repressive 10 + 7 5’-LR…. AATAGGGGCCTCGCTTACATG non-repressive 11 GGGGCCTCGCTTAC + 8 5’-LR…. ATAAATA ATG repressive

Figure 7. The Repressive effect of the 5’-LR maps to a promoter distal ATAAATA motif Left: schematic illustration of the extent of the 5′-LR DNA in Pr-luxAB transcriptional reporter plasmids. The extent of deleted regions within the 5’-LR are shown as open regions. The oblique line indicates the presence of a non-native BglII site introduced to allow manipulation of the 5′-LR DNA. Right: Luciferase plate tests of P. putida KT2440 harbouring these plasmids (upper) and key DNA sequences of plasmids harbouring the Δ4 to Δ8 deletions. Figure adapted from Paper 1, Fig. 2.

3.1.1 Potential mechanisms underlying the action of the ATAAATA DNA motif So how does the ATAAATA DNA motif mediate its effects on the numbers of transcripts? As well as identifying the responsible DNA element, the deletion analysis shown in Fig. 7 also relocates the ATAAATA motif to different positions relative to the Pr promoter. For instance as close as 14 bp downstream (as in the large Δ8 deletion), as compared to its native location at 108 bp downstream Pr. Because the ATAAATA motif can have its effects in the absence of the vast majority of upstream and downstream dmpR-derived DNA (e.g. Δ6), this makes any riboswitch or transcriptional attenuation mechanism based on restructuring of the mRNA highly unlikely.

Two alternative possible mechanisms that would impede progression of σ70-RNAP from the Pr promoter were experimentally addressed. The first of these was suggested by the similarity of the ATAAATA motif to the -10 element of σ70-promoters (consensus TATAAT) – such -10 mimics 25 can cause pausing by binding of partially detached 70 in the early stage of transcriptional elongation to result in lower transcript levels (75-77). However, using transcriptional reporter gene fusions to other promoters, the repressive effect of the ATAAATA motif of the 5’-LR was found to operate with both strong and weak promoters dependent on 70 and with promoters dependent on alternative -factors with very different recognition motifs, thus eliminating this possibility (Paper I, Fig. S1). However, although we consider it unlikely due to the magnitude of the effect, these experiments do not eliminate the possibility that progression of σ70-RNAP from Pr is impeded by transient binding of another σ70-RNAP complex to the ATAAATA motif.

A second, and more likely, possibility we considered was that the ATAAATA motif represents a binding site for a transcriptional repressor. The repressive effect of the ATAAATA motif was also found when using the same transcriptional reporter plasmids (1 and 2, Fig. 7) in a heterologous E. coli host, implying that such a potential repressor would have to be available in both E. coli and P. putida. HNS and HNS-like proteins drew our attention as candidates because the ATAAATA motif bears similarity to the consensus sequence of E. coli HNS (TCGATAAATT). HNS is one of the main nucleoid-associated proteins in enterics that is now recognized as a general silencer of many stress-response genes as well as genes acquired by horizontal transmission (78, 79). Transcriptional reporter assays using strains with single null mutations of each of the five known HNS-like proteins of P. putida refuted the idea of HNS-like proteins as the potential repressors acting through this motif (data not shown). However, HNS- like proteins possess the AT-rich DNA binding hook motif [Q/RGR] and this Q/RGR motif is functional in otherwise structurally unrelated proteins (80). Thus, it still remains possible that some other protein may bind to the ATAAATA-binding to mediate transcriptional repression.

Comparative analysis of luciferase activity profiles across the growth curve in rich media revealed similar repression effects of the 5’-LR DNA on transcription in both the exponential- and stationary- phases of growth (Paper I, Fig. 1B). It is worth noting here that the increase in transcription at the exponential-to-stationary phase transition, observed irrespective of the presence or absence of the 5’-LR, is due to elicitation of (p)ppGpp synthesis that directly stimulate Pr activity (37, 38). These findings suggest that if such a potential repressor exists, then its levels and activity remain relatively constant under the single experimental condition tested so far. Notably, whatever mechanism underlies the inhibitory effect of the ATAAATA DNA motif of the 5’-LR, it acts counteractively to the stimulatory effect of (p)ppGpp seen as the cells enter the stationary phase. Such simultaneous negative and positive regulatory input has been found in

26 so called “incoherent feed-forward loops” that have been proposed to function as pulses- generators and/or mediate accelerated replies in response to regulatory signals (81).

3.2 The CA-motif within the 5’-LR results in translational inhibition of dmpR Next we addressed whether the 5’-LR is also involved in controlling translation of dmpR. As a starting point, we generated a Pr-dmpR expression plasmid with the Δ8 deletion of 5’-LR because it was i) the largest 5´-LR deletion which preserved wild-type transcript levels and ii) maintained the integrity of the RBS of dmpR (see schematic in Fig. 8). DmpR levels produced from otherwise identical Pr-dmpR expression plasmids, either having the full length 5’-LR or its Δ8 variant, were monitored in two ways. Firstly, by introduction of the plasmids into a P. putida KT2440 reporter strain carrying a Po-luxAB transcriptional fusion on its chromosome – which provides a sensitive genetic system to assay intracellular DmpR levels as reflected by DmpR- dependent Po promoter activity, and secondly, by Western analysis of DmpR in extracts from cells cultured to the exponential and stationary phases of growth in LB containing a potent aromatic effector for DmpR activity (Paper I, Fig. 3).

In contrast to the wild-type counterpart, the Δ8 derivative produced elevated levels of DmpR in the exponential phase, as assessed by both the biological Po-activity read-out and Western analysis (Paper I, Fig. 3B and C). Given that transcription from Pr was unaffected (Paper I, Fig. 3D), this data provided strong support for a role of the 5’-LR mRNA in mediating translational repression of dmpR. As expanded on in later sections, this exponential phase muting of DmpR levels would be anticipated to contribute to down-regulation of the auxiliary (methyl)phenol catabolic pathway when preferred carbon sources are present.

A. RBS AUAAAUAGGGGCCUCGCUUACAUG 8 ...UUUCCCCAUCUAAAAAUAAAUAGGGGCCUCGCUUACAUG WT ...UUUCCCCAUCUCCCCAUAAAUAGGGGCCUCGCUUACAUG 4C substitution mutant AAnAAnAA–catabolite activity motif (CA-motif) B.

U U A C U C-GA O G- -C Potential 2 structure of the 5’-LR mRNA C-G C-G G-C C-G AU-A 8 A C Two alternative substructures G -C G -C U - U C -G U -A - A U U UU C AU UA A AA A 4C A A A A U A G C AAAU GA- 5’ CG

Figure 8. A functional CA-motif is found within the 5’-LR of dmpR. A. Alignment showing the key RNA sequences of the 5’-LRs of dmpR variants with manipulations in the potential CA-motif (consensus 82; orange). The shaded box illustrates the ribosome binding site (RBS) that overlaps the CA-motif. The ATG initiation codon of dmpR is highlighted in cyan italics. B. Predicted secondary structure of the 5’-LR mRNA of dmpR with two alternative substructures shown with the presumed CA-motif and overlapping RBS highlighted. Figure adapted from Paper I, Fig. 7.

As overviewed in the introduction, the Crc protein is a key player in catabolite repression control in P. putida. Previously identified targets of Crc-mediated translational repression possess a Catabolite Activity (CA) motif (consensus AAnAAnAA where ‘n’ is any ribonucleotide) in proximity to, or overlapping, the RBSs (62, 82, 83, 84). Three observations prompted us to assess the role of Crc as a likely candidate involved in translational repression of dmpR: I) the presence of a CA-motif (AAAAAUAA, orange in Fig. 8) overlapping the presumed RBS of dmpR; II) that removal of half of this site (as in the Δ8 Pr-dmpR derivative) resulted in elevated DmpR levels in the exponential phase where carbon repression control by Crc is at its highest (8); and III) that there was little effect on cellular DmpR levels in the stationary phase, where Crc activity would be counteracted by the two sRNAs, CrcZ and CrcY, proposed to act in concert to sequester Crc to block its activity (63).

To verify the presumed CA-motif within the 5’-LR was the target through which translational repression was exerted, we constructed another mutant, denoted 4C (Fig. 8). This mutant of the CA-motif has four consecutive C residues substituting the first half of the CA-motif, but maintains the remainder of the 5’-LR and the RBS intact. The 4C mutant showed the same effect on DmpR levels as the Δ8 deletion mutant (Paper I, Fig. 3), defining this region as a critical component of the target site.

3.2.1 Crc exerts translational regulation on dmpR in vivo The next question became, does Crc mediate translational repression in vivo? To answer this question, we monitored the effect of artificial manipulations of Crc on dmpR translation. These experiments used a Po-luxAB transcriptional reporter plasmid carrying Pr-dmpR in its native configuration with respect to Po (Paper I, Fig. 4A), introduced independently into wild-type, a Crc-null, and a double CrcZ/CrcY-null derivative of P. putida KT2440. Strains were again assayed in LB containing a potent aromatic effector required for DmpR activity.

Although the growth rate was badly affected by the removal of Crc, presumably as a consequent of its global regulatory role in metabolism, the absence of Crc resulted in relieved translational inhibition of DmpR levels in the exponential phase where Crc is normally available (Paper I, Fig. 4B compared to 4C). As anticipated, lack of the sRNAs which would otherwise counteract Crc activity (CrcZ and CrcY), resulted in decreased DmpR levels in the stationary phase (Paper I, Fig. 4D). Consistent with these findings, we found that artificially decreasing the available level of Crc, by overexpressing CrcZ or CrcY from an IPTG inducible promoter, enhanced DmpR levels, as assessed by its biological read-out through the activity of the Po-promoter (Paper I, Fig. 5A and B). This effect of CrcZ or CrcY over-expression was not seen in strains already lacking Crc (Paper I, Fig. 5C and D), confirming that Crc activity is antagonized by both CrcZ and CrcY.

3.2.2 Crc and Hfq act as co-partners in formation of riboprotein complexes in vitro Does Crc need another factor to be able to form of riboprotein complexes through CA-motifs? We were prompted to ask this question in the light of the finding that P. aeruginosa Crc is unable to bind RNA (66), and the suggestion that previously observed RNA-binding activity with Crc-His preparations was due to low level contamination by the naturally His-rich E. coli RNA chaperone Hfq. Rather than interpreting this as an in vitro artefact per se, we considered that Hfq may indeed be a genuine co-partner required for Crc involvement in RNA binding to CA-motifs and, therefore, catabolic repression. Hence, our next approach was to determine if this was the case using purified P. putida proteins prepared either without a His tag (Crc) or from an E. coli Hfq-null strain (wild-type Hfq-His and substituted derivatives thereof).

Hfq is an abundant protein that forms homohexameric ring with three RNA-binding surfaces. The proximal surface binds short U-rich sequences typically found in sRNAs. The distal surface of Hfq binds A-rich longer ARN repeats of mRNAs [where A is an adenine, R is a purine, and N can be any ribonucleotide] (85, 86) that are very similar to the CA-motif described in the 29 preceding section. An additional, less defined surface, consisting of the outer rim or lateral RNA binding surface which connects the proximal and distal RNA-binding sites, may be important for binding internal U-rich sequences of sRNAs (87). Since P. putida and E. coli Hfq proteins are highly similar with more than 90% identity in their N-terminal 72 residues, in addition to wild- type P. putida Hfq-His, we also generated and purified a Y25D substitution derivative (predictably defective in binding A-rich RNA) and a K56A substitution derivative (predictably defective in binding U-rich RNA).

Using these preparations in RNA electro-mobility shift assays (EMSA), we found that: 1) consistent with findings from P. aeruginosa, P. putida Crc alone was unable to bind to the CA- motif of dmpR (Paper I, Fig. 6A; 66), 2) that Hfq could only bind the CA-motif at very high concentrations to form an unstable complex (Paper I, Fig. 6B), and most dramatically, 3) that although Crc could not bind RNA alone, it triggered formation of a distinct stable riboprotein complex in the presence of very low levels of Hfq that on their own were unable to elicit formation of any detectable complex (Paper I, Fig. 6D and 6E). Fig. 9 illustrates the kind of data obtained.

(0.8 µM) (0.7 µM) (0.8 µM) (0.7 µM) 0 0 0 0 Crc/Hfq Co-complex

WT RNA 4C RNA probe probe (0.01 µM) (0.35-1.4 µM) (0.8-3.2 µM) (0.35-1.4 µM) (0.8-3.2 µM) (0.01 µM)

UUCCCCAUCUAAAAAUAAAUAGGGGC UUCCCCAUCUCCCCAUAAAUAGGGGC

Figure 9. Crc and Hfq work in partnership to form a stable riboprotein complex

In agreement with this in vitro data, lack of Hfq was found to have a similar effect on DmpR levels as lack of Crc (Paper 1, Fig. 4E and F), supporting the idea that these two protein cooperate in mediating their effect. That this was through the identified CA-motif of the 5’-LR was corroborated by the findings that 1) no co-complex could be formed by Crc-Hfq when using probes carrying the 4C substitution that abolishes repression in vivo (Paper 1, Fig. 3 and Fig. S2), and 2) that co-riboprotein complex formation required the A-rich binding properties of Hfq, but not its U-rich RNA binding property (Paper 1, Fig. 6C & F). 30

Co-participation of Crc and Hfq has implications for all Crc-targeted mRNA and also for how the sRNAs CrcZ and CrcY, each with their six CA-motifs, function to control Crc activity through its availability. Similar Crc-Hfq co-dependent stable complex formation was obtained with other RNA oligonucleotides containing CA-motifs previously considered as Crc targets present in the Crc regulated alkS mRNA, benR mRNA, phaC1 mRNA and CrcZ carrying its six CA-motifs (Paper II: Fig. 2A, B, C and Fig. 4D).

The higher numbers of CA-motifs of CrcZ also allowed a different approach to monitor Crc and Hfq behaviour using Hfq substitution derivatives. This approach involved a pull-down strategy to purify Hfq-His associated complexes in the presence or absence of Crc and CrcZ (or tRNA as non-specific structured RNA) by affinity chromatography using Ni-NTA agarose beads. Subsequent analysis by the SDS-PAGE electrophoresis and western blotting revealed that Crc and Hfq can form a ternary co-complex with CrcZ in vitro, but that these two proteins made no detectable co-complex unless there is a specific RNA to be bound (Paper II: Fig. 5A and B). As found for the CA-motif of dmpR, formation of the ternary riboprotein complex relied on the A- rich binding properties of Hfq [intact in the wild-type and the K56A substitution derivative, but not in the Y25D derivative] (Paper II: Fig. 5C). These data are all constant with the idea that RNA binding by the distal face of Hfq initiates the events that lead to the co-participation of Crc to form a stable complex capable of inhibiting translation of mRNAs on one hand, and sequestering of Crc on the other, as schematically illustrated in Fig. 10.

U U A C U CrcZ & C-GA G- -C C-G CrcY C-G G-C C-G AU-A A C G -C G -C U - U C -G U -A - A U U UU C Hfq AU UA A AA A A A A A U A G C AAAU Crc GA- 5’ CG

Two alternative potential substructures of the dmpR 5’-LR

Figure 10. Translational inhibition of mRNA for less preferred substrates versus sequestering via sRNAs 31

3.3 Crc and Hfq participation in catabolic repression Catabolite repression control is strong under conditions such as exponential phase growth in rich LB medium or in minimal medium containing two carbon sources – one preferred and one less preferred. To ascertain the levels of all the players in the Crc-Hfq/CrcZ-CrcY circuitry, we determined protein levels by western analysis and CrcZ and CrcY levels by quantitative PCR under different growth conditions (Paper II, Fig. 9). The results corroborated previous finding that Crc levels remain relatively constant, while CrcZ and CrcY levels become highly elevated when catabolic repression needs to be relieved (61,63); and established that in P. putida, Hfq levels, like those of Crc remain relatively constant under different conditions.

As alluded to in the previous section, co-dependence on Hfq and Crc for down regulation of DmpR levels was verified in vivo using single and double null mutants of P. putida. Inactivation of the hfq gene relieved Crc-mediated repression of DmpR levels to a similar extent as achieved when Crc or both Crc and Hfq were lacking (Paper I, Fig. 4E, F). P. putida Crc-mediated translational repression of transcriptional regulators has previously been shown to result in down-regulation of mRNA of cognate pathways for less preferred amino acids. To gain further information on the cooperative influence of Crc and Hfq on hierarchical carbon metabolism in P. putida, quantitative PCR was used to determine the mRNA levels of selected gene in wild-type, Crc-null and Hfq-null cells cultured to the exponential phase in LB medium. These genes included hpd and phhA [both involved in the assimilation of aromatic amino acids], bkdA1 and ivd [both involved in the assimilation of branched-chain amino acids, and aatJ [involved in the transport of acidic amino acids] (62, 88). In each case, lack of Crc or Hfq resulted in similar elevation of the levels of these mRNAs.

Taken together, the combined data described above, firstly re-establishes the previously proposed model for Crc-mediated catabolic repression with the critical inclusion of the RNA chaperone Hfq; and secondly identified the Crc-Hfq/CrcZ-CrcY circuitry as a control check point in expression of the DmpR master regulator of (methyl)phenol catabolism.

3.3.1 Crc-Hfq input on DmpR levels and (methyl)phenol catabolism Because DmpR controls expression of the Dmp-enzymes, the negative input of Crc-Hfq on DmpR levels would be anticipated to influencing growth at the expense of (methyl)phenols in the presence of preferred substrates. Therefore, the next question to address was if this level of control through DmpR was sufficient to impose catabolic repression on the native dmp-system carried on the pVI150 plasmid of P. putida CF600. 32

As a first step, we confirmed that P. putida CF600, like P. putida KT2440, had all the components of the Crc-Hfq/CrcZ-CrcY circuitry (Paper III, Figs. S1 and S2). Having established that this was the case, we next engineered our two Crc-Hfq target-site-deficient mutations (4C and 8) within the native pVI150 plasmid in P. putida CF600 and compared the ability of the resulting strains to grow in minimal media at the expense of phenolics. As anticipated, no effect of these manipulations were seen under these non-catabolite repression growth conditions (Paper III, Fig. 1B).

We next tested for effects through the Crc-Hfq target of dmpR under catabolic repression conditions and monitored depletion of 2.5 mM 2-methyphenol over growth. Using 50% LB as the repressive medium, we could not detect any effect of the 4C and 8 mutations on 2- methylphenol catabolism, which remained restricted to the stationary phase (Paper III, Fig. 2). This was surprising since the 4C and 8 strains, as expected, had de-repressed DmpR levels in the exponential phase (Paper III, Fig. 2 western blot image). Likewise we found no detectable effect of de-repression of DmpR via the 4C and 8 mutations in minimal medium containing 10 mM glutamine as the preferred carbon source and 2.5 mM 2-methylphenol as the non-preferred substrate (Paper III, Fig. 3A). However, in this case, growth in the two carbon sources displayed a distinct diaxie lag phase as the bacteria switched from growth at the expense of glutamine to growth at the expense of 2-methylphenol (Paper III, Fig. 3A).

To be able to amplify any potential differences between the adaptation and/or fitness properties of these strains, we first differentially marked them with fluorescence tags to facilitate identification of strains in co-culture competition experiments. However, once again we were unable to detect any differences mediated by the 4C and 8 mutations under catabolite repression conditions – minimal medium containing glutamine and 2-methylphenol – even after repeated cycles of sequential dilution and growth every 12 hours for four days (Paper III, Fig. 3B). Our interpretation of these results is that Crc-Hfq mediated repression of DmpR levels is insufficient to abolish catabolic repression on dmp-system, and that other Crc-Hfq target sites within the dmp-operon likely also contribute to subverting (methyl)phenol catabolism when preferred substrates are present.

3.3.2 The dmp-system has multiple additional CA-motifs for targeting by Crc-Hfq In silico analysis of the dmp-operon identified four additional potential CA-motifs within the polycistronic mRNA, each in close proximity to the cognate translation initiation codons (Paper

III, Fig. 4A). Three of these genes encoding subunits of the multicomponent phenol hydroxylase (dmpL, dmpN, and dmpO) for the first step of the pathway, and one encodes the ring cleavage enzyme catechol 2, 3-dioxygenase (dmpB) for the second step.

We subjected these potential sites to the same battery of in vitro EMSA tests used to define the co-dependence of riboprotein complex formation on Crc and Hfq at the CA-target site of dmpR. All showed the same properties, namely: dependence on the co-addition of Crc and Hfq for stable complex formation and the requirements of the A-rich RNA binding property of Hfq. However, titrations of Crc (paper III, Fig. 4) and competition experiments using probes with or without a functional CA-motif (paper III, Fig. 5 & Fig. S3), revealed different apparent affinities. Based on these data, we classified the CA-motifs of dmpR, dmpO and dmpB as high-affinity sites and those dmpL and dmpN as lower affinity sites. However, it should be emphasised that these comparative affinities were determined with short (26 nt) RNA probes, and that relative affinities might differ when presented in the context of full length mRNAs.

Taken together, these data support the idea that targeting of translation of the enzymes for the first two steps in (methyl)phenol degradation works alongside control of DmpR levels to subvert (methyl)phenol catabolism in the presence of preferred substrates.

3.4 Current model for hierarchical regulation of (methyl)phenol catabolism by the dmp- system The main findings presented in this thesis can be summarised as follows:

I] The ATAAATA DNA motif of the initially transcribed region of the dmpR results in lower levels of transcripts. While we have been unable to trace the underlying mechanism, I would speculate that this motif serve as a DNA-binding site for an as yet unknown repressor. Signal-responsive control of such a repressor could potentially add an additional level of regulation to feed into the dmp-system to control (methyl)phenol catabolism via transcriptional regulation. Such regulation would work alongside aromatic-effector activation of DmpR and (p)ppGpp regulatory inputs to couple metabolism of phenols to the presence of substrate and nutritional stress.

II] The CA-motif of dmpR and other CA-motifs implicated in hierarchical assimilation of different carbon sources in P. putida are all co-targeted by Crc and Hfq to form a stable riboprotein complex in vitro. Where tested, lack of Crc or Hfq have similar repercussions on 34

Crc-targeted mRNAs in P. putida. This places Hfq firmly alongside Crc in the carbon repression control circuity employed to optimise use of available carbon sources. This previously unknown translational regulatory input into the dmp-system would work hand-in-hand with transcriptional responses to i) provide robust expression of the pathway enzymes when need and ii) help subdue expression when preferred carbon sources are available.

III] The CA-motif of dmpR alone is insufficient to account for carbon catabolite repression on the dmp-system. Rather, multiple Crc-Hfq CA-target sites within dmpR and the dmp-operon likely mediate a two-tiered simultaneous translational control – one at the level of the DmpR transcriptional regulator, and the other at the level of Dmp-enzyme production. I would speculate that removal of any individual one of the five targets identified would be insufficient to alleviate catabolic repression in the face of a preferred carbon source.

Like others before me, my research so far leaves many questions only partially answered and has generated some new ones. Below I bring up three of what I consider the most immediate questions arising from my work on carbon repression control and (methyl)phenol catabolism in P. putida.

How does the Crc-Hfq riboprotein complex form and inhibit translation? The simplest interpretation of the in vitro and in vivo data with these two proteins is a direct competition model whereby the Crc-Hfq co-complex occludes binding of the ribosome (as depicted in Fig. 10). However, other possible scenarios, including locking the ribosome or the 30S complex in a non-functional state, could also account for translational repression. Hfq is an extremely adaptable protein. Although it usually functions as a chaperone for base-pairing between sRNAs and their mRNA targets, it can perform non-canonical regulatory roles by binding directly to the mRNA (89). Hfq can also interact with other proteins, either directly or through association with a common RNA template. For example E. coli Hfq has been found to interact with at least 30 proteins or large protein complexes (54, 90). Given this adaptability of Hfq, a minimum of three different mechanisms can be envisaged to underlie the interaction between Crc, Hfq and RNA: i) the Hfq protein recognizes the CA-motif first to recruit Crc through restructure the RNA and/or direct protein:protein interactions, and only then can Crc- bind the RNA, ii) Hfq and Crc form a pre-complex with the RNA and then Hfq leaves to fulfil its functions elsewhere, or iii) Crc-Hfq protein:protein interaction stimulate the RNA binding property of Hfq while Crc never binds the RNA at all.

Does the 5’-LR of the dmpR mRNA impose other regulatory inputs? The CA-motif involved in Crc-Hfq mediated control only covers 8 nt (7%) of the 123 nt 5’-LR region of the dmpR mRNA. In comparison with other chromosomally- or plasmid-encoded phenol catabolic systems of P. putida strains, this 5’-LR region is more highly conserved than other regulatory or protein coding regions (Paper I, Fig. S4). This raises the question of whether the remaining regions of the 5’-LR are simply required for correct presentation of the CA-motif on an exposed loop, or if they might also be involved in mediating other regulatory inputs. As schematically illustrated in Fig. 11, the latter possibility is suggested by two other potential regulatory features – a small upstream ORF (uORF) and a target site for the PhrS sRNA.

U U A C U C -GA G- -C An upstream ORF (uORF) C-G A target site for PhrS C -G G-C 5’-LR dmpR 5’ UGCC-UUGCGCUAUG C -G AU- A PhrS-creg 3’ ACGG-AACGCACGAG A C G -C G -C 5’-LR pqsR 5’ AGCCCUUGCGCUUCC U -U C -G U -A - A U U UU C AU UA A A A A A A A A U A G C AAAU GA- 5’ CG

Figure 11. Additional potential regulatory features of the 5’-region dmpR Two alternative initiation codon and the termination codon of the uORF (grey) are shown in red, while that for dmpR is highlighted in cyan. Complementarity of the potential PhrS target site (blue) within the uORF is compared with its known target on the right. Figure adapted from Paper I.

The uORF might be involved in translational coupling by ribosomal read-through of the weak UAG termination (91) and scanning to the downstream AUG initiation codon of dmpR. In doing so, it could increase the translational efficiency by increasing the frequency of translational initiation events directly. In addition, passage of the ribosome may alter the secondary structure of the mRNA and could also serve as a means of disrupting the Crc-Hfq riboprotein complex when catabolism of phenols is needed. It is notable that the potential target site for PhrS within this uORF bears greater complementarity to the 5’-LR of dmpR than it does to its known target – the pqsR mRNA of P. aeruginosa. This regulatory RNA mediates pairing through a discreet region (denoted creg) and its levels increase when cells enter stationary phase (52). Hence PhrS could provide yet another regulatory link between nutritional cues and metabolism of phenolic compounds by playing upon translation of DmpR.

Is the ability to adapt to growth on phenolics an individualistic event? Much recent research has focussed on heterogeneity of gene expression in genetically identical cells as an evolutionary selected property that ensures the presence of a sub-population cells ready to respond to a new environmental condition (92). As emphasised throughout this thesis, the dmp-system has multiple known (and potential) regulatory inputs at both the transcriptional and translational level. Each of these has the capability, through stochastically events, to generate cells with higher- or lower- than any average content of different components. For example, (p)ppGpp levels that appear to underlie the sub-population of metabolically inactive “persisters” that are less susceptible to antibiotics (reviewed in 20). Generation of heterogeneity, or so called “bet-hedging”, has been observed in many bacterial processes, including adaption from growth on one carbon source to another (93). In contrast to the Monod concept of metabolic adaptation of an entire population during the diauxic lag phase (94), this phase can also be due to the period of time it takes a sub-population to grow to sufficient levels as to be delectable with a spectrophotometer (95). I would speculate that the multiple regulatory inputs to the dmp-system would result in a heterogeneous population of genetically identical cell, only a sub-proportion of which could actually be capable of adapting to growth on toxic phenolics.

Acknowledgements

This is a very good opportunity and worth to take some time to send my heartiest thanks to everyone those who have supported and helped me during my PhD studies both within and outside of work. First, I would like to express my deepest appreciation to my supervisor Vicky (Professor Victoria Shingler) for your great guidance and supervision on my PhD work and for always trying your best with good organisation and time-management abilities to optimise the research progress. And also I would like to thank Prof. Martin Gullberg for your excellent supportive ideas for some of my experimental approaches.

Enormous gratitude goes to my first met lab partners Eleonore, Teresa and Sofia, yet my words are not enough to thank you all and I wish to be lucky again to work with people like you at my next carrier life. You all made my life easier and happier during my PhD. Eleonore, I truly felt you like my own Mother in Sweden who was always so touched and I like the way you behave as a calm and patient lady. Teresa, you are my Best friend as this friendship will never end wherever I go. You had so much good qualities and I really couldn’t believe such a person exist in real life! Sofia, I have been following you as my life Guider, so that I could balance my professional life with my family life. Hope your son, Rasmus is a big boy now. I want to appreciate your prompt support for me at any time when technical problems came across and also for that you didn’t miss to go with me to every friday & wednesday seminars. Big thanks also go to my present group members, Lisa and Henrik for being such good friends whom I have shared lab with during the last years.

I am very thankful to all my past and present journal club members including Prof. Maria Fällman, Dr. Matthew Francis, Assoc. Prof. Åke Forsberg, Tomas Edgren, Anna fahlgren for giving me inputs and comments for building up my self-confidence. I also take this opportunity to thank Prof. Staffan Bohm, Prof. Sun Nyunt Wai, Prof. Anders Byström, Prof. Jan Larsson and Prof. Bernt Eric who involved in evaluating my progress reports every year and Prof. Jörgen Johansson, for go-ahead approval for my Thesis and Prof. Sven Bergström with his ever-smiling face for being a humble professor. I wold like to thank my collaboration group in Spain including Fernando Rojo, Reneta, Sofia whom I met in the Pseudomonas conference 2013.

I would like to thank Mikael Wikström (for helping me with the ladok registrations), Marek (for helping me with all IT solutions). I want to thank past & present nicest administrative staff Berit, Helena, Maya Sandow, Ethel, Ingella, Maria and Selma for supporting me with official work and also to Jonny, to Siv, to Media room staff, and to Jessica and Marianella for making my working life easier.

I would like to thank those with whom I shared teaching responsibility -Reine and Stefanie Mangold, I was lucky to have you two as my Guiders for teaching, so I learnt lot from you to execute well planned and organized practical sessions to assist the undergraduates with my best. I also Thank Jessica (for your best effort even when you were sick), Frederik Hugosson, Kristoffer, Dharmesh and Anne-Laure (calm and helpful) and for all being good teaching partners.

I also want to thank you all the present and past nicest people (project students, doctoral students, post docs) I met at the department of Molecular Biology from the ground floor to 3 floors up including Christine Grundstrom, Kamaraj, Soffie, Rajdeep, Neyyer, Pramod, Mitesh, Annika, Marylise, Monika, Christopher Andersson, Sakura, Teresa tiensuu, Stefanie, Svitlana, Hassan, Tony, Hao Xu, Gunilla, Laura, Sara, Akhilesh, Junfa, Jyoti,

Kristina, Ievgen, Tracy, Susanne, Mridula, Damini, Helena, Christian Spoerry, Margerida and Philge Philip.

I never ever forget my very good and nice friends whom I had fun with. Thank you so much for having good times with me- Roshani, Ala, Nicole, Akbar, Lena, Anne-Laure, Geetanjali, Steffi, Hande & Frederic, Bernard, Tomas & Helen, Sofia Håglin, Saskia, Sabine, Connie, Ummehan & Kemal, Sarp, Nikola, Nabil, Bhupender, Edvin, Patricia & Tiago, Swarupa and Ikenna. Thank you very much Edvin for sharing all your past experience in preparations for the Thesis and the Defence. Further, I truly appreciate your words that helped me to put my stress down to some extent. Big thanks for my Sri Lankan best friends- Gangani & Chammi akki for your nice friendship and also to my Sri Lankan neighbours in Umeå- specially Nilupa akki for sharing your experience in many aspects.

I am deeply indebted to my lovely parents (Sujatha & Upasena) and my one and only lovely brother (Sheron) for their love and affection and their proper leading and motivation which brought me this far not only to reach this level of education but also as a good mannered citizen to the world. Special Thanks to my father who took care of my little daughter while I was busy with writing the Thesis. My loving husband (Madawa), without you I would never have managed to get this far to complete my PhD studies. I truly appreciate your full dedications and patience for understanding me with my busy working life. However, you taught me the importance of a balanced life, so that I was lucky to be a mother of two lovely daughters (Sanuji & Vinuji) with whom I tried to forget the hard times. Your huge support during my long days of experiments while providing good protection and care for me and our children is unbelievable. And also I would like to thank all my other family members- Parents-in-Law (Nanda & Lionel), Brother-in-Law (Nilanga ayya), Sisters- in-Law (Nisansala nangi, Jeevani akki and Warunika akki) and our little ones (Chenuki duwa, Dinuki duwa & Manuth putha) for all their supportive talks and activities and being with me both at difficult days and joyful days.

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