Metabolism-Dependent Taxis and Control of Motility in Pseudomonas Putida

Metabolism-dependent taxis and control of motility in Pseudomonas putida

Sofia Österberg

Department of Molecular Biology Umeå University Umeå 2013

This work is protected by the Swedish Copyright Legislation (Act 1960:729) ISBN: 978-91-7459-563-5 Cover picture: Electron microscopy image of Pseudomonas putida KT2440 Electronic version available at http://umu.diva-portal.org/ Printed by: Department of Chemistry Printing Service, Umeå University Umeå, Sweden 2013

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CONTENTS

CONTENTS ...... I ABSTRACT...... III ABBREVIATIONS ...... IV LIST OF PUBLICATIONS ...... V SAMMANFATTNING PÅ SVENSKA ...... VI 1. INTRODUCTION ...... 1

1.1 BACTERIAL ADAPTATION ...... 1 1.2 BACTERIAL TRANSCRIPTION ...... 1 1.2.1 RNA polymerase – the molecular machinery ...... 1 1.2.2 σ-factors – the specificity components ...... 2 1.2.3 The transcriptional process from start to finish ...... 3 1.2.4 Transcriptional regulation ...... 4 1.2.4.1 Riboswitches ...... 4 1.2.4.2 σ-factor control and anti-σ-factors ...... 5 1.2.4.3 σ-factor release at a morphological checkpoint ...... 6 1.2.4.4 Targeting of RNAP by the global regulator ppGpp ...... 7 1.2.4.5 Transcriptional regulation in relation to habitat ...... 7 1.3 PSEUDOMONAS ...... 8 1.4 BACTERIAL MOTILITY ...... 8 1.4.1 The flagellum ...... 9 1.4.2 Flagellum assembly ...... 9 1.4.3 Bacterial taxis ...... 10 1.4.3.1 The chemotaxis signalling pathway ...... 10 1.4.3.2 Taxis receptors ...... 12 1.4.3.3 Energy taxis and the Aer receptor ...... 13 1.4.4 Regulation of motility ...... 14 1.4.4.1 The c-di-GMP lifestyle switch ...... 14 1.4.4.2 C-di-GMP turnover enzymes ...... 15 1.4.4.3 The PAS domain ...... 16 1.4.4.4 Bifunctional c-di-GMP turnover enzymes ...... 16 1.4.4.5 C-di-GMP responsive transcriptional regulators ...... 17 1.4.4.6 C-di-GMP dependent riboswitches, RNA processing, and proteolysis ...... 17 1.4.4.7 C-di-GMP binding domains ...... 18 1.4.4.8 The flagellar brake protein YcgR ...... 19 2. AIMS ...... 20 3. RESULTS AND DISCUSSION ...... 21

3.1 METABOLISM DEPENDENT TAXIS BY PSEUDOMONAS PUTIDA CF600 ...... 21 3.1.1 P. putida CF600 shows metabolism-dependent taxis towards dmp- pathway substrates ...... 21 3.1.2 Aer2 is the metabolism-dependent and aerotaxis receptor of P. putida KT2440 ...... 22 i

3.1.3 The three Aer receptors are co-localized but differently expressed 24 3.2 TRANSCRIPTIONAL REGULATION OF THE AER RECEPTORS ...... 25 3.2.1 aer2 but not aer1 or aer3 are transcribed by σFliA-RNAP ...... 25 3.2.2 The P. putida σFliA consensus is suboptimal in vitro ...... 26 3.2.3 The aer2 UP-element controls output and σFliA specificity ...... 27 3.2.4 DksA and ppGpp affect aer2 transcription and motility ...... 28 3.3 THE BI-FUNCTIONAL C-DI-GMP TURNOVER ENZYME PP2258 AND MOTILITY CONTROL ...... 29 3.3.1 PP2258 can both synthesise and degrade c-di-GMP ...... 29 3.3.2 The in vivo activity of PP2258 is dependent on its concentration ... 30 3.4 PP4397 – THE LINK BETWEEN THE C-DI-GMP SIGNAL AND ALTERED MOTILITY ..... 32 3.4.1 PP4397 is located in a flagella associated operon ...... 32 3.4.2 PP4397 is downstream of PP2258 in the signalling pathway ...... 33 3.5 POSSIBLE AER1-PP2258-PP4397 SIGNALLING MECHANISMS AND FUTURE PERSPECTIVES ...... 33 ACKNOWLEDGEMENTS ...... 36 REFERENCES ...... 38 PAPER I-IV

ABSTRACT Bacteria living in soil and aquatic habitats rapidly adapt to changes in physico-chemical parameters that influence their energy status and thus their ability to proliferate and survive. One immediate survival strategy is to relocate to more metabolically optimal environments. To aid their movement through gradients (a process called taxis), many bacteria use whip like flagella organelles. Soil-dwelling Pseudomonas putida possesses a polar bundle of flagella that propel the bacterium forward in directed swimming motility. P. putida strains are generally fast growing, have a broad metabolic capacity, and are resistant to many harmful substances – qualities that make them interesting for an array of industrial and biotechnological application. This thesis identifies some of the factors that are involved in controlling the flagella driven motility of P. putida. In the first part of the thesis, I present evidence that P. putida displays energy-taxis towards metabolisable substrates and that the surface located Aer2 receptor (named after its similarities to the Escherichia coli Aer receptor) is responsible for detecting the changes in energy-status and oxygen-gradients that underlie this response. Aer2 is expressed simultaneously with the flagella needed for taxis responses and its expression is ensured during nutrient scares conditions through the global transcriptional regulators ppGpp and DksA. In addition to Aer2, P. putida possesses two more Aer-like receptors (Aer1 and Aer3) that are differentially expressed. Like Aer2, Aer1 and Aer3 co-localize to one cell pole. Although the signals to which Aer1 and Aer3 respond are unknown, analysis of Aer1 uncovered a role in motility control for a protein encoded within the same operon. This protein, called PP2258, instigated the work described in the second part of my thesis on the involvement of the second messenger c-di-GMP in regulation of P. putida motility. Genetic dissection of the catalytic activities of PP2258 revealed that it has the unusual capacity to both synthesize and degrade c-di-GMP. Coupling of the c-di-GMP signal originating from PP2258 to motility control was traced to the c-di-GMP binding properties of the protein PP4397. In the last part of the thesis, I present possible mechanisms for how these different components might interact to create a signal transduction cascade – from the surface located Aer1 receptor to PP2258 and the c-di-GMP responsive PP4397, and from there to the flagella motors – to ultimately determine flagella performance and the motility status of P. putida.

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ABBREVIATIONS 2D-TLC Two dimensional thin layer chromatography 5'-UTR 5’-Untranslated region ACF Alternative cellular function AMP Adenosine monophosphate CCW Counter clockwise c-di-GMP Bis-(3’-5’)-cyclic dimeric guanosine monophosphate CW Clockwise DGC Diguanylate cyclase DNA Deoxyribonucleic acid EPS Extracellular polymeric substances FAD Flavin adenine dinucleotide GMP Guanosine monophosphate GTP Guanosine triphosphate HAMP domain Histidine kinase, adenylyl cyclases, methyl-binding proteins and phosphatases domain MCP Methyl accepting protein MH1 Methylated helix 1 MH2 Methylated helix 2 mRNA Messenger RNA nt Nucleotide NTP Nucleoside triphosphate PAS domain Per-ARNT-Sim domain PDE Phosphodiesterase pGpG 5'- Phosphoguanylyl-(3'-5')-guanosine PNPase Polynucleotide phosphorylase ppGpp Guanosine pentaphosphate or tetraphosphate Ppi Pyrophosphate RNA Ribonucleic acid RNAP DNA-dependent RNA polymerase rRNA Ribosomal RNA STYKs Serine/threonine/tyrosine kinases tRNA Transfer RNA YFP Yellow fluorescence protein

LIST OF PUBLICATIONS

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

I Sarand, I., Österberg, S., Holmqvist, S., Holmfeldt, P., Skarfstad, E., Parales, R.E., and Shingler, V. (2008) Metabolism-dependent taxis towards (methyl)phenols is coupled through the most abundant of three polar localized Aer-like receptors of Pseudomonas putida. Environmental Microbiology 10: 1320-1334.

II Österberg, S., Skarfstad, E., and Shingler, V. (2010) The σ-factor FliA, ppGpp and DksA coordinate transcriptional control of the aer2gene of Pseudomonas putida. Environmental Microbiology. 12:1439-1451.

III Österberg, S.*, Åberg, A.*, Herrera Seitz, M.K., Wolf-Watz, M., and Shingler, V. Genetic dissection of a motility-associated c-di-GMP signalling protein of Pseudomonas putida. Submitted manuscript.

IV Österberg, S.*, Skoog, L.*, and Shingler, V. PP4397 provides the link between PP2258 c-di-GMP signalling and altered motility in Pseudomonas putida. Preliminary manuscript.

*Authors contributed equally

SAMMANFATTNING PÅ SVENSKA Bakterier kan leva nästan var som helst, till och med på ogästvänliga platser, exempelvis heta källor eller förorenad jord. Oavsett var bakterierna befinner sig så måste de generera energi för att överleva. Faktorer som påverkar denna förmåga varierar konstant i många miljöer och ett sätt för bakterier att anpassa sig till dessa förändringar är att förflytta sig till de för stunden mest gynnsamma platserna. Bakterier kan ändra sin rörelseriktning på mindre än en sekund, vilket gör den här typen av reaktion till en omedelbar överlevnadsstrategi. Många bakterier producerar och använder piskliknande utskott som kallas för flageller för att kunna transportera sig. Bakterierna simmar genom att rotera flagellerna som då driver bakterien framåt. Alla bakterier som kan simma är kemotaktiska. Det betyder att de kan känna av olika faktorer så som näringsämnen, syre och ljus, och röra sig mot eller ifrån dessa (processen kallas kemotaxi). Det finns en variant av kemotaxi som kallas för energitaxi och som innebär att bakterier rör sig mot den överlag mest metabolt gynnsamma miljön istället för mot specifika faktorer. Ett receptorprotein som kan känna av energinivåerna och ändra rörelsemönstret i bakterien Escherichia coli kallas för Aer. Bakterier kan reglera sin rörlighet på flera olika sätt. Ett sätt är genom att producera signalmolekylen c-di-GMP. När bakterier ökar den intracellulära mängden c-di-GMP roterar flagellerna långsammare och bakterierna rör sig därmed mindre och vice versa. Enzymerna som producerar c-di-GMP kallas för diguanylatcyklaser och de som bryter ner (tar bort) molekylen kallas för fosfodiesteraser. Trots att dessa enzymer har motsatta funktioner så är de ofta ihopkopplade till ett protein (istället för att vara två separata) och utgör då varsin domän av proteinet. I de flesta fall så är då bara ett av enzymerna fungerande medan den andra reglerar det fungerande enzymets funktion. Koppling mellan signalmolekylen c-di-GMP och flagellerna är i E. coli proteinet YcgR. YcgR kan binda c-di-GMP och bryta viktiga kopplingar i motorerna som roterar flagellerna. Jag har studerat en jord- och vattenlevande bakterie som kallas Pseudomonas putida. Denna bakterie kan använda många ämnen som ingen annan bakterieart kan utnyttja som energikälla. Den är dessutom motståndskraftig mot många substanser som är skadliga för andra bakteriearter. De här två egenskaperna gör P. putida intressant för många bioteknologiska tillämpningar. För att utnyttja bakterien optimalt krävs dock god förståelse av dess beteende. Målet med min forskning har varit att studera hur P. putida kontrollerar sin rörlighet. I den första delen av avhandlingen presenterar jag bevis för att P. putida uppvisar energitaxi mot näringsämnen och att Aer2-receptorn (har

vi fått sitt namn eftersom den liknar Aer i E. coli) är ansvarig för att känna av de energiförändringar inuti bakterien som föranleder rörelsen (Artikel I). P. putida producerar Aer2-receptorn och flagellerna (som behövs för rörelsen) samtidigt (Artikel II). Två globala transkriptionsreglerare försäkrar dessutom att Aer2 produceras när P. putida befinner sig i näringsfattiga miljöer (Artikel II). Detta medför troligtvis att bakterien kan förflytta sig till mer näringsrika miljöer när det är av särskild vikt. Förutom Aer2 så har P. putida en liknande receptor som kallas Aer1. Signalen som Aer1-receptorn känner av är ännu okänd men under analysen av Aer1 så upptäcktes ytterligare ett protein som är involverat i kontollen av P. putidas rörlighet (Artikel I). Upptäckten av det här proteinet, som kallas PP2258, föranledde det arbete som presenteras i den andra delen av avhandlingen rörande effekten som c-di-GMP har på P. putidas rörelseförmåga. Undersökningar av PP2258s enzymatiska förmåga avslöjade att PP2258 är ovanlig i och med att det här proteinet både kan producera och bryta ner c-di-GMP (Artikel III). Kopplingen mellan c-di-GMP-signallen som PP2258 producerar och P. putidas flageller innefattar proteinet PP4397 som har vissa likheter med YcgR i E. coli (Artikel IV). I avhandlingens sista del presenterar jag en möjlig modell för hur dessa olika proteiner kan interagera för att skapa en signalväg – från ytan av P. putida där Aer1-receptorn är placerad, via PP2258 som reglerar nivåerna av c-di-GMP och PP4397 som känner av signalmolekylen, och till sist till flagellerna – för att i slutändan avgöra hur P. putida rör sig.

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1. INTRODUCTION

1.1 Bacterial adaptation Bacteria grow and proliferate almost everywhere, even in hostile environments such as hot springs and polluted soils. However, independent of where they live, continuous energy generation is one of the most important aspects for their survival. Physico-chemical parameters that influence bacterial survival and proliferation include temperature, pH, osmolarity, and nutrient sources. These parameters are constantly changing in aquatic and terrestrial habitats. One of the many ways that bacteria adapt to these fluctuations is to relocate to more favourable environments in response to environmental cues. For example, upon detection of an attractant, bacteria can change their direction of movement in less than a second (Segall et al., 1982), making this type of response an immediate survival strategy (reviewed in Alexandre et al., 2004). To aid their movement, many bacteria use whip like motility organelles called flagella. Production of these motility organelles is tightly regulated at the transcriptional level (reviewed in Aldridge and Hughes, 2002).

1.2 Bacterial transcription Control of transcriptional initiation – the first step in copying the genetic information from DNA into RNA – is a primary access point for regulating gene expression in bacteria. In addition to regulation of flagella biosynthesis, control at this level is used in many other bacterial adaptive processes. As overviewed in Fig. 1, transcription carried out by the RNA polymerase (RNAP) can be divided into three main phases: initiation, elongation, and termination. The first of these steps – initiation – involves promoter recognition, separation of the two DNA strands, and initial synthesis of a small nascent RNA. The elongation phase occurs after the RNA polymerase has left the promoter and consists of continued linking of nucleoside triphosphates (NTPs) together to form the RNA copy complementary to the DNA template. The transcriptional process ends with termination, where the DNA-RNAP-RNA complex dissociates into its component parts.

1.2.1 RNA polymerase – the molecular machinery DNA-dependent RNAP is a complex molecular machine that has to undergo multiple conformational changes as it binds and reads the DNA to make RNA. RNAP consists of six subunits, five of which are always present and make up the core enzyme, and one which is exchangeable – the σ-factor.

The core enzyme consist of the 2α, β, β’, and ω subunits and it is this part of the RNAP that has the catalytic function (reviewed in Borukhov and Nudler, 2008). The N-terminal domains of the two α subunits and the greater parts of β and β’ make up the stationary mainframe that serves three main functions: i) to bind the DNA-RNA hybrid within the main channel in the correct orientation with respect to the catalytic site, ii) to form a pore (secondary channel) through which the NTP substrate for RNA synthesis enter, and iii) to bind the σ-factor. Mobile parts of the core enzyme in and around the major channel function in RNA formation, changes in the conformation of the RNA exit channel, and movement of RNAP along the DNA as transcription progresses.

Figure 1. The bacterial transcription cycle The transcription cycle starts with a σ-factor binding to the core RNAP. Since the core RNAP is present in a lower concentration than the σ-factors, the latter have to compete for core RNAP binding. The RNAP then scans the DNA in search of a promoter to which it binds to initiate transcription. When the RNA reaches 8-11 nt, transcription is either aborted or the RNAP escapes the promoter and the elongation phase starts. The σ-factor is stochastically released during the elongation phase and returns to the σ-factor pool. The transcription ends with termination where upon the DNA, core RNAP and RNA separate. Adapted from Osterberg et al. (2011).

1.2.2 σ-factors – the specificity components The σ-factor, of which there are several different variants, directs the RNAP to the promoter regions upstream of genes. The σ-factor confers specificity to the RNAP by providing the recognition determinants for binding DNA elements within promoters (reviewed in Borukhov and Nudler, 2008). Each type of σ-factor has specific DNA motifs that it recognizes and binds to. For example, the housekeeping σ-factor (σ70) recognizes -35 and -10 elements

(where +1 is the transcriptional start site) with the consensus sequence - 35TTGACA-30 and -12TATAAT-7, respectively. The relative spacing of these elements can vary, with 17 bp being optimal. In addition to the housekeeping σ-factor that guides the RNAP to the largest bulk of promoters within all bacterial genomes, most bacterial species also possess additional σ-factors – so called alternative σ-factors – which direct RNAP to other classes of promoters for developmental and adaptive responses (reviewed in Helmann, 2002; Gruber and Gross, 2003). The number of σ-factors encoded in a genome reflects the lifestyle of the bacterium, for example enteric Escherichia coli has seven different σ-factors while free-living Pseudomonas putida has twenty-four (Shingler, 2003). The alternative σ-factors have generally lower affinity for the core RNAP than the housekeeping σ70 (Maeda et al., 2000) and the concentrations of the alternative σ-factors are also always greatly exceeded by those of σ70 (Grigorova et al., 2006).

1.2.3 The transcriptional process from start to finish As illustrated in Fig. 1, the transcription process starts with a σ-factor binding to the core RNAP to form a holoenzyme (reviewed in Hsu, 2002; Borukhov and Nudler, 2008; Nudler, 2009). The RNAP holoenzyme then scans the DNA until it encounters an appropriate promoter sequence. Once a promoter is found, the RNAP binds to it and forms a closed complex. The specific binding by the RNAP to the promoter sequence is performed by the σ-subunit, however, the binding can be strengthened through interactions between the α-subunits and A and T rich UP-elements when present (Ross et al., 1993). Within the closed complex, the DNA is destabilized and both the σ-factor and core enzyme contribute to binding of the two separate strands of the transiently melted DNA to create an open transcription bubble. When the core RNAP binds to the template DNA strand, the +1 base (where the transcription starts) gets located within the catalytic site. This RNAP-DNA structure is now called an open complex and the first NTP complementary to the +1 base can be orientated within the catalytic site. RNA synthesis per se starts with a second NTP binding to the +2 position, formation of a phosphodiester bond between the two NTPs and displacement of pyrophosphate (PPi). Next, the RNA-DNA hybrid is moved one nucleotide towards the RNA exit channel leaving the NTP insertion site free to bind a new NTP. As this process continues, the nascent RNA grows. When it reaches 8-11 nt in length, its progress to the exit channel is obstructed by parts of the σ-factor. This conflict is resolved by either abortive initiation where the small nascent RNA is released and the whole process of initiation has to start again, or by reconfiguration of the σ-factor to remove the obstruction to allow continued RNA synthesis. Up to this point in the 3

RNA synthesis process, the RNAP has not moved, and so the DNA-RNA complex within the main channel of the RNAP is at this stage causing stress on the DNA backbone. This stress ultimately leads to rewinding of upstream DNA, which breaks DNA and σ-factor interaction. This, together with the aforementioned reconfiguration of the σ-factor, enables the RNAP to escape from the promoter and continue into the elongation phase. Since several interactions between the σ-factor and core RNAP are also interrupted during these steps, the σ-factor is fully released – an event that occurs stochastically, usually during synthesis of the initial 100 to 200 nt of RNA. Transcription not only has to start at the right place, it also needs to stop at the appropriate site, i.e. where the gene or operon ends. For many genes, there are intrinsic sites encoded in the DNA that causes the RNAP to terminate the transcription. There are also specific proteins involved in terminating transcription, such as the termination factor Rho. Rho uses ATP to move along the RNA that is being synthesized until it catches up with the RNAP. It then pushes the RNAP forward, pulls the RNA from the RNAP or unwinds the DNA-RNA hybrid – all resulting in termination of transcription. The core RNAP is then free to associate with a new σ-factor and start the transcription process again.

1.2.4 Transcriptional regulation Transcription can be controlled at all steps and this control can involve different kinds of factors (e.g. proteins, small molecules, and signals encoded in the RNA) that work in various ways, see Fig. 2 (reviewed in Borukhov and Nudler, 2008; Nudler, 2009; Osterberg et al., 2011). Classical transcriptional regulators include proteins that bind to DNA within or near the target promoter or at more remote sites. These factors either increase or decrease basal transcriptional initiation rates from promoters in response to specific signals that control their activities. Three other means of transcriptional control act through i) the initially transcribed regions of RNA, ii) σ-factor regulation, and iii) by targeting the RNAP performance directly.

1.2.4.1 Riboswitches Transcription starts upstream of the coding sequence of genes (i.e. the part that is translated into protein). In some cases, the initially transcribed region lies far upstream of the translational start codon, leading to the formation of 5’-untranslated regions (5’-UTRs) in the cognate messenger RNAs (mRNAs). These 5’-UTRs can be up to hundreds of nucleotides long and can have regulatory functions, both at the transcriptional and translational level. Riboswitches form one type of regulatory element that can be found in 5’- UTRs (reviewed in Garst et al., 2011; Serganov and Patel, 2012). Riboswitches are structures in the mRNA that are able to bind small

4 molecules and affect transcription or translation, where the most common outcome is transcriptional attenuation. A typical riboswitch contains two domains: an aptamer domain and an expression platform. The aptamer is the part of the mRNA that binds the ligand and it has a specific three dimensional fold. The expression platform has a structure that is altered upon ligand binding and it is this domain that interacts with the transcriptional or translational machinery.

Figure 2. Regulatory factors that can control transcriptional output Transcriptional regulation can occur at many different steps of the transcription cycle and involve different factors, some of which are depicted in the figure. Adapted from Osterberg et al. (2011).

1.2.4.2 σ-factor control and anti-σ-factors Since σ-factors are the determinants for promoter recognition and compete for core RNAP to form the holoenzyme, altering their relative concentrations, activity, and/or availability is one mean of regulating basal transcription rates. In addition to transcription and translational control, other mechanisms that regulate the activities of σ-factors includes targeted degradation (Zhou et al., 2001), σ-factor activation by proteolytic cleavage (Hilbert and Piggot, 2004), and generation of σ-factor variants that have a high turnover (Kim et al., 2009). However, a predominant mode of regulating the availability of σ-factors is by anti-σ-factors (reviewed in Osterberg et al., 2011). Anti-σ-factors bind their cognate σ-factors tightly to mask the σ-factor interfaces that normally interact with the core RNAP. Thus, anti-σ-factors sequester their cognate σ-factors and prevent them from binding to core RNAP. The anti-σ-factor mechanism allows for a rapid

5 accumulation of σ-factors when they are required, since no de novo synthesis steps are needed. Signals that lead to σ-factor release include different physico-chemical stresses, iron limitation and organelle biogenesis.

Figure 3. Regulation of flagella biosynthesis Transcription of genes involved in flagella biogenesis is hierarchical. The genes in the last level of the hierarchy are transcribed by the σFliA-RNAP. Up until completion of the flagellar basal body and hook structures, the specialised σFliA is sequestered by its anti-σ-factor FlgM. Once the hook is in place, FlgM is secreted and σFliA is free to associate with the core RNAP. The last genes can thus be transcribed and the flagellum structure completed.

1.2.4.3 σ-factor release at a morphological checkpoint Transcription of the genes needed for flagella biogenesis is hierarchical to ensure correct assembly. One checkpoint in this process is complete assembly of the hook basal body structure that anchors the flagellum filament to the cell. Up to this point the alternative σ-factor σFliA is bound to its anti-σ-factor FlgM. However, once the hook basal body is completed, FlgM is secreted through this structure with σFliA acting as a chaperon, see Fig. 3 (reviewed in Aldridge et al., 2006). This leaves σFliA free to associate with core RNAP and initiate transcription of the last genes needed for completion of the flagellum structure as well as proteins involved in chemotaxis (see section 1.4.2 Flagellum assembly). Unlike most other σ- factor/anti-σ-factor pairs, FlgM and σFliA are not transcribed together within the same operon. In Salmonella, the flgM gene is regulated by two promoters where one promoter is σFliA-dependent – creating a feedback loop

6 that aids in maintaining a balance between σFliA-dependent transcription and transcription dependent on other σ-factors (Gillen and Hughes, 1993).

1.2.4.4 Targeting of RNAP by the global regulator ppGpp When bacteria encounter physico-chemical stresses, they produce a nucleotide called ppGpp. This small molecule is an “alarmone” that reprograms the transcriptional machinery from expressing genes needed for growth to those needed for adaptation and survival (reviewed in Dalebroux and Swanson, 2012). This is achieved both directly by ppGpp-targeting the RNAP to alter its properties, and indirectly by changing σ-factor competition, and thus the composition of the RNAP holoenzyme pool. Direct targeting of RNAP by ppGpp alters its performance at susceptible promoters to either decrease or increase their output. The effect of ppGpp is often amplified by the regulatory protein DksA, which sensitizes RNAP to ppGpp by binding to the secondary channel of the RNAP (reviewed in Gourse et al., 2006). ppGpp vastly decreases output from powerful σ70-dependent promoters such as those that drive transcription of ribosomal RNA (rRNA) and transfer RNA (tRNA) genes (reviewed in Dalebroux and Swanson, 2012). This, in turn, is thought to result in more available core RNAP that alternative σ-factors can compete for, and consequently to indirectly facilitate expression of genes whose products are crucial for survival. DksA and ppGpp have been shown to negatively regulate motility in E. coli (Aberg et al., 2008; Lemke et al., 2009). These two transcriptional regulators affect flagellar biosynthesis by decreasing the output of genes in the two top levels of the transcriptional hierarchy, and thus also genes in the last level (transcribed by σFliA-RNAP) indirectly, resulting in non-flagellated cells under very poor nutritional conditions (Lemke et al., 2009). However, there are contradicting results that instead indicate that ppGpp leads to flagella production (Magnusson et al., 2007).

1.2.4.5 Transcriptional regulation in relation to habitat Free-living bacteria, in contrast to those that are intracellular pathogens or endosymbionts, have a large proportion of their genomes dedicated to encoding transcriptional regulators. Free-living bacteria are faced with constantly changing environments and it is probably this fact that is reflected in the evolution and selection of a greater number of regulators within their genomes (Cases et al., 2003). These regulators likely help the bacteria to integrate and respond to the many physico-chemical parameters that influence their survival. Two environments where fluctuations are considerable are water and soil and in both these types of habitats bacteria belonging to the Pseudomonas genus can be found.

1.3 Pseudomonas Pseudomonas species are rod shaped gram-negative gamma Proteobacteria. This genus is composed of omnivorous saprophytes and, due to their simple nutritional needs, they can be found in most temperate, aerobic and semi- aerobic waters and soils. In these habitats, they are involved in element cycling, degradation and recycling of biotic and xenobiotic organic compounds. The Pseudomonads are considered as specialists in metabolising diverse organic compounds, even toxic ones, and they are often resistant to harmful substances. These qualities make them attractive for various biotechnological applications such as bioremediation (Dejonghe et al., 2001), production of biocatalysts (Schmid et al., 2001), and de novo synthesis of chemicals (Poblete-Castro et al., 2012). P. putida strains are frequently isolated from polluted soils and their metabolism has been studied due to their ability to degrade aromatic compounds. P. putida strains generally grow fast and have a high biomass yield in addition to its low maintenance demands and high tolerance against toxins and growth inhibiting substances, which has made it an interesting organism for industrial applications (reviewed in Poblete-Castro et al., 2012). To be able to thrive in diverse environments and under harsh conditions, the Pseudomonads have some of the largest genomes amongst free-living bacteria (Cases et al., 2003), and a high proportion of their genes are dedicated to carbon metabolism, transport and efflux of organic compounds as well as regulation (Nelson et al., 2002). However, to efficiently grow and proliferate, the Pseudomonads also need to move to environments that are optimal for catabolism and energy-generation.

1.4 Bacterial motility The ability to move is an important property for many bacteria. For some, it is a part of their virulence strategy, while for others it enables symbiosis with their host organism. A third aspect is that it allows the bacterium to transport itself to beneficial environments, e.g. towards nutrient sources or away from toxic compounds. Several different types of bacterial motility has evolved, these include gliding, twitching, swarming, and swimming. Gliding is unaided by pili or flagella (reviewed in Nan and Zusman, 2011), twitching is dependent on type IV pili (reviewed in Mattick, 2002), while flagella are required for swarming and swimming (reviewed in Patrick and Kearns, 2012). The difference between the latter two is that the bacteria either move as individuals in aqueous environments (swimming) or in groups of cells over moist surfaces (swarming), which requires secretion of wetting agents (reviewed in Patrick and Kearns, 2012). 8

1.4.1 The flagellum Flagella – the organelles that mediate the movement in swimming and swarming – are long filamentous structures that are rotated like propellers to drive the bacteria forward. The proton driven flagellar motor rotates amazingly fast with a speed of up to 18,000 rpm (Chen and Berg, 2000) – the same maximum rpm as for a Formula one race car engine. There are also more rare Na+ driven flagellar motors and these can rotate even faster, with up to 100,000 rpm (Magariyama et al., 1994). This results in movement at speeds of up to 100 body lengths per second! Different bacteria species have different numbers of flagella and they also locate them differently, everything from one polar localized flagellum to several dispersed all over the surface of the bacterium can be found – spirochaetes even have internal flagella located within the periplasm. The flagellum can be divided into three basic parts: the basal body, hook, and filament, see Fig. 3 (reviewed in Jarrell and McBride, 2008). The basal body anchors the flagellum filament to the cell and also contains the motor that drives its rotation. The basal body is mainly made up of a series of protein rings located in different planes of the bacterial envelope: in the outer membrane – the L ring, in the peptidoglycan layer – the P ring, in and above the inner membrane – the MS ring, and in the cytoplasm – the C ring. The C-ring, which is also called the switch complex, is a part of the flagellar motor and consists of three proteins (FliG, FliM, and FliN) that control the rotational direction of the flagellum. The other part of the motor is the stator that confers energy to the motor by passing protons over the inner membrane. The stator is made up of two proteins called MotA and MotB, where MotB holds the stator in place by binding to the peptidoglycan layer and MotA interacts with FliG and turns the C-ring around. A rod-like protein structure is located in the centre of the basal body rings. This rod extends to the outside of the cell where it is attached to a flexible hook that changes the angle of rotation. Torque is passed from the flagellar motor, via the rod to the hook, and finally to the filament which is the propeller that ultimately pushes the bacterium forward.

1.4.2 Flagellum assembly The flagellum is assembled in a highly ordered manner from the inside out – the basal body first, then the hook, and finally the filament. Genes encoding the flagellar proteins are under hierarchical control, with either three or four tiers of regulation, to ensure that the proteins are expressed at appropriate times (Chilcott and Hughes, 2000; Dasgupta et al., 2003). There are some common themes to the regulation irrespective of the number of regulatory levels (Dasgupta et al., 2003; reviewed in Jarrell and McBride, 2008). One or more master regulators lie at the top of the 9 regulation hierarchy. These are transcriptional regulators that react to quorum sensing signals, global regulators, and small molecules and initiate transcription of genes encoding basal body and hook components as well as other regulatory proteins required for subsequent transcription of other flagellar genes. Another common feature is that the last genes in the hierarchy are transcribed by σFliA-RNAP. The FliA protein is sequestered until needed (see Fig. 3), but once associated with the core RNAP, the genes encoding hook associated proteins, flagellin to make up the flagella filament, motor proteins (MotA and MotB), and proteins involved in the chemotaxis signalling system are expressed (see section 1.2.4.3 σ-factor release at a morphological checkpoint). A number of regulatory proteins are involved in the flagella assembly. For example, in Pseudomonas aeruginosa FleN controls the number of flagella produced (Dasgupta et al., 2000), FlhF determines the polar localization of the flagellum (Pandza et al., 2000), and FleL functions as a ruler and determines the length of the flagella (Dasgupta et al., 2003).

1.4.3 Bacterial taxis Bacteria that can swim do so by rotating their flagella counter clockwise (CCW) to move forwards and clockwise (CW) to change their swimming direction – in bacteria that have multiple flagella this causes tumbling and a random change in the direction of movement. Almost all motile bacteria, including those that can swim, are chemotactic (reviewed in Szurmant and Ordal, 2004). This means that they can sense chemoeffectors and alter their otherwise random movement either to move away or towards them. Chemoeffectors consists of nutrients, toxins, oxygen, pH, osmolarity, and light (reviewed in Wadhams and Armitage, 2004). Since bacteria are small, they sense chemoeffector concentrations temporarily, instead of along the length of their body. When the bacteria swim in a favourable direction, they elicit longer periods of CCW flagellar rotation and smooth swimming. This in turn leads to an overall movement towards optimal environments and is called a biased random walk.

1.4.3.1 The chemotaxis signalling pathway The signalling system that underlies chemotaxis involves the Che proteins CheA, CheB, CheR, CheY and CheZ in addition to the chemoeffector receptors [see section 1.4.3.2 Taxis receptors and Fig. 4] (reviewed in Porter et al., 2011). When the receptors sense decreased amount of attractants, they activate autophosphorylation of the CheA histidine kinase. The phosphoryl group is then transferred to response regulator CheY, which diffuses to the flagellar motor and interacts with the switch complex proteins FliM and 10

FliN. This interaction promotes the flagella to rotate CW and the bacterium will thus change its swimming direction. This signal is terminated once CheY-P becomes dephosphorylated, a process that is aided by specific phosphatases, e.g. CheZ in E. coli.

Figure 4. The classical chemotaxis signalling pathway The chemotaxis receptors (MCPs) are, together with CheA and CheW, located in polar clusters. The MCPs sense chemoeffectors (attractants or repellents) and transfer the sensed signal to CheA. This alters the autophosphorylation activity of CheA. The phosphoryl group on CheA can be transferred to CheY, which in turn sets the rotational direction of the flagella to CW. The system is reset by CheZ, which dephosphorylates CheY. CheZ also localizes to the polar clusters via CheA and CheW (Cantwell et al., 2003), but is here depicted free in the cytosol for simplicity. The CheB methylesterase and CheR methyl transferase make up the memory of the chemotaxis system. The phosphoryl group of CheA can also be transferred to CheB, which then removes methyl groups from the MCPs and make them less prone to activate CheA. The action of CheB is counteracted by CheR, which adds methyl groups to the MCPs.

The phosphoryl group of CheA can also be transferred to CheB, a phosphorylation-responsive methylesterase. CheB demethylates glutamic residues on the chemotaxis receptors [CheB-P performs this 100 times faster than CheB (Anand and Stock, 2002)], which decreases the ability of the receptors to activate CheA and adapts the system to the current chemoeffector concentration. The action of CheB is counteracted by the methyltransferease CheR. This circuit thus functions as a primitive memory for the bacteria and allows them to sense changes in a few molecules of a chemoeffector against background concentrations that can differ with at least five orders of magnitude (Wadhams and Armitage, 2004).

Many different variations of chemotaxis have evolved in different bacteria. Different bacterial species have different numbers of the chemotaxis components, alternative adaptation mechanisms exist, and there are receptors located in the cytoplasm with different subcellular localization in addition to the classical receptors found in the inner membrane (reviewed in Krell et al., 2011; Porter et al., 2011).

1.4.3.2 Taxis receptors The receptors that sense chemoeffectors are called methyl accepting proteins (MCPs) and the number of MCPs encoded in bacterial genomes correlates with their lifestyle – strict pathogens have few while bacteria with complex life styles have many (reviewed in Krell et al., 2011). As illustrated in Fig. 5, a typical MCP has a periplasmic ligand binding domain and a cytosolic signalling and adaptation domain (reviewed in Baker et al., 2006). The latter domain is made up of an α-helix that extends away from the inner membrane into the cytoplasm where it forms a hairpin structure. The cytoplasmic domain can be divided into four subdomains: closest to the membrane is a histidine kinase, adenylyl cyclases, methyl-binding proteins and phosphatases (HAMP) domain, then follows methylated helix 1 (MH1), the signalling domain at the hairpin turn, and methylated helix 2 (MH2) extending towards the membrane juxtaposed to MH1. The MH1 and MH2 together contain the glutamic acids (four or more) that are substrates for CheB and CheR. MCPs form dimers, which in turn forms trimers with each other. In E. coli, these trimers are situated the cell pole where they are part of signalling clusters together with the histidine kinase CheA and the coupling protein CheW – both of which are required for the clustering to occur (reviewed in Baker et al., 2006). This clustering of receptors is believed to allow for allosteric interactions, amplification of signals and sensing of small changes in chemoeffector concentrations (reviewed in Porter et al., 2011). When a ligand is bound by one receptor subunit, it causes several conformational changes: the periplasmic surface of the receptor is altered, the orientation of the receptor subunit is changed relative to its dimeric partner, and the receptor dimer changes orientation with respect to the membrane (reviewed in Baker et al., 2006). The new conformation of the receptor dimer is believed to cause disorder in the receptor cluster, which results in an altered phosphorylation status of CheA.

Figure 5. Classical MCP versus the Aer receptor The classical MCPs consist of a periplasmic ligand binding domain, a HAMP domain and two helixes that can be methylated. The Aer receptor, on the other hand, contains a FAD binding PAS domain in the cytoplasm, a HAMP domain and two helixes that cannot be methylated. Both types of receptors form homodimers that in turn can form heterotrimers.

1.4.3.3 Energy taxis and the Aer receptor There is a variant of chemotaxis called energy taxis (or metabolism- dependent taxis) where the bacteria move towards the metabolically most favourable environment (reviewed in Taylor et al., 1999; Alexandre et al., 2004). Energy taxis receptors monitor either the redox state of the electron transport chain or the proton motive force across the inner membrane and enable the bacteria to sense metabolizable substrates, electron acceptors, and metabolic inhibitors. Although energy taxis has been found in a wide variety of bacterial species, sensing in energy taxis is only understood for a few (reviewed in Schweinitzer and Josenhans, 2010). The most studied energy taxis receptor is the Aer protein of E. coli. Aer is a low abundance taxis receptor constituting approximately 2.5% of the total receptor pool. It can function without the aid of other MCPs (Bibikov et al., 2004), however, in the normal setting it forms homodimers that then form trimers with the classical MCPs in the receptor clusters (Gosink et al., 2006). Unlike the ligand binding MCPs, Aer has its sensing domain facing the cytoplasm where it senses redox changes in the cell, see Fig. 5 (Edwards

13 et al., 2006). Signal sensing occurs through a FAD-binding Per-ARNT-Sim (PAS) domain (see section 1.4.4.3 The PAS domain) that is connected to the HAMP domain via a transmembrane anchor. It is not known exactly how Aer senses redox changes but it might involve exchange of the bound FAD with the cytoplasmic FAD/FADH pool, since the cofactor is loosely associated with the receptor (reviewed in Taylor, 2007). Once the signal is sensed by the PAS domain, this domain directly interacts with the HAMP domain in the cytoplasm (reviewed in Taylor, 2007) – this contrasts classical MCPs where the signal is propagated to the HAMP domain via the transmembrane region. Another difference between Aer and the classical MCPs is that Aer cannot be regulated by CheR and CheB because it lacks the methylation sites needed (Bibikov et al., 2004).

1.4.4 Regulation of motility Flagella driven motility can be regulated in several different ways. The first site where regulatory mechanisms act is on flagellum assembly. For example, master regulators of flagella gene expression is regulated posttranscriptionally in E. coli and P. aeruginosa. In E. coli the global regulator CsrA activates translation of the master flagella genes (flhDC) (Wei et al., 2001) while expression of the master regulator FleQ in P. aeruginosa (see section 1.4.4.5 C-di-GMP responsive transcriptional regulators) is repressed by Vfr – a transcriptional regulator activated by the second messenger cyclic AMP (Dasgupta et al., 2002). Once the flagella are formed another second messenger called bis-(3’-5’)-cyclic dimeric guanosine monophosphate (c-di-GMP) can influence the swimming behaviour of bacteria.

1.4.4.1 The c-di-GMP lifestyle switch C-di-GMP was first discovered in the context of cellulose synthesis in Gluconacetobacter xylinus in the late 1980’s (Ross et al., 1987). This small molecule has since then been shown to have a major role in development, virulence, motility, and biofilm formation. The involvement of c-di-GMP in biofilm formation has been extensively studied. Biofilms, which are formed on surfaces, are multi-cellular communities where the bacteria interact with each other and create a three dimensional structure that enables them to better cope with their environment (reviewed in Boyd and O'Toole, 2012). Although the biofilm is advantageous for the bacteria, it can cause severe problems for humans since it can confer higher antibiotic resistance and cause biofouling of medical implants and industrial implements. The formation of the biofilm starts with bacteria swimming to a surface where they attach. This initial attachment is regulated by the c-di-GMP

14 responsive SadB protein in P. aeruginosa, where high levels of the second messenger promote attachment (Merritt et al., 2007). In P. aeruginosa both type IV pili and Cup fimbria also have roles in the formation of the biofilm and expression of all of them is c-di-GMP regulated (reviewed in Harmsen et al., 2010). To form the three dimensional structure of a biofilm, bacteria produce extracellular polymeric substances (EPS). The matrix that the EPS form is important for attachment, interactions between the bacteria, tolerance, and exchange of genetic material (reviewed in Harmsen et al., 2010). The EPS consists, among other things, of different polysaccharides whose production is controlled by c-di-GMP (reviewed in Harmsen et al., 2010). Under certain conditions, e.g. when nutrients are depleted, the biofilm is dispersed and the bacteria become free living once again. Dispersal of P. aeruginosa biofilms is triggered by the BdlA regulator that lowers the intracellular levels of c-di-GMP (reviewed in Harmsen et al., 2010). In Pseudomonas fluorescence and P. putida, dispersal involves three Lap proteins – LapA, LapG and LapD. LapA is an adhesive protein that mediates attachment to surfaces and it is released via proteolysis by LapG when low c- di-GMP levels are detected by LapD [see section 1.4.4.6 C-di-GMP dependent riboswitches, RNA processing, and proteolysis] (Gjermansen et al., 2010; Newell et al., 2011).

1.4.4.2 C-di-GMP turnover enzymes C-di-GMP is synthesized by enzymes called diguanylate cyclases (DGCs) that contain the GGDEF motif within their catalytic A-site. The DGCs form homodimers and convert two molecules of GTP into one c-di-GMP (Paul et al., 2004; Ryjenkov et al., 2005; Wassmann et al., 2007). In addition to the A-site, some DGCs also contain an inhibitory site (I-site) that can bind c-di- GMP and hinder further synthesis (Chan et al., 2004; Christen et al., 2006). Enzymes called phosphodiesterases (PDEs) have opposing function to the DGCs and degrade c-di-GMP. This class of enzymes can either have an EAL or an HD-GYP domain. The EAL containing PDEs degrade c-di-GMP to linear pGpG (Christen et al., 2005; Schmidt et al., 2005) while the HD-GYP containing ones can convert c-di-GMP to two molecules of GMP (reviewed in Ryan et al., 2006). Genes encoding potential DGCs and PDEs are exclusively found in bacteria and they are abundant within most bacterial genomes, e.g. Pseudomonas putida KT2440 contains 42 such genes (Galperin et al., 2010; Ulrich and Zhulin, 2010). With the large number of DGCs and PDEs found in some bacterial species, there must be regulatory mechanisms to avoid unwanted cross talk between different c-di-GMP responsive signalling pathways. One such mechanism is sequestration of the c-di-GMP turnover

15 enzymes to specific compartments of the cell, as is the case for the complex that inhibits the E. coli PNPase [see section 1.4.4.6 C-di-GMP dependent riboswitches, RNA processing, and proteolysis] (Tuckerman et al., 2011). Regulation of the amount of enzymes expressed might also avoid cross talk between different signalling pathways. A third mean of control lays within the sensory domains that most of the DGC and PDE proteins are coupled to. These domains can perceive external or internal signals (e.g. light, oxygen, and the energy status of the bacterium), and change the overall activity of the protein. One sensory domain that is frequently linked to c-di-GMP turnover proteins is the PAS domain.

1.4.4.3 The PAS domain The PAS domain can be found in all kingdoms of life, but 87% of all predicted PAS domains are found in prokaryotic genomes (reviewed in Henry and Crosson, 2011). The PAS domain can either be part of single domain proteins or linked together with other domains; there can even be several PAS domains in the same protein. When constituting part of a multi domain protein, PAS domains are often found at the N-terminal of both enzymatic and non-enzymatic effector domains (reviewed in Galperin, 2004; Henry and Crosson, 2011). The effector domains that the PAS domain is coupled to are part of a wide variety of classes, implicating it in many different signalling systems, not only those involving c-di-GMP (reviewed in Henry and Crosson, 2011) The PAS domain can bind a wide spectrum of ligands and cofactors and function as a sensor for different signals (e.g. gases, redox potential, and light) and it can also mediate protein-protein interactions (reviewed in Moglich et al., 2009; Henry and Crosson, 2011). However, most of the PAS domains seem to have a role in signal transduction, e.g. in nucleotide cyclases and phosphatases (reviewed in Galperin, 2004). When coupled to c- di-GMP turnover DGCs and PDEs, it is most often found at the N-terminal (reviewed in Henry and Crosson, 2011).

1.4.4.4 Bifunctional c-di-GMP turnover enzymes Although the DGCs and PDEs counteract each other, they are often found linked together in the same protein. In most cases studied, only one of the domains is enzymatically functional – the other is non-enzymatic but instead regulates the function of the protein (reviewed in Schirmer and Jenal, 2009). So far, a few proteins in various bacterial species have been found to both synthesise and degrade c-di-GMP. The first bifunctional enzyme identified was the unusual Rhodobacter sphaeroides bacteriophytochromes BphG1 (and BphG2). Tarutina et al. (2006) showed that the BphG1 EAL domain functions as a light independent PDE, while the GGDEF domain requires

16 that the EAL domain is proteolytically cleaved off and that light of the right wavelength is sensed. Since then six other bifunctional enzymes have been found, including a response regulator (Levet-Paulo et al., 2011), two enzymes affecting long-term survival (Kumar and Chatterji, 2008; Gupta et al., 2010), one biofilm regulator (Ferreira et al., 2008), and as shown in this thesis, a regulator of motility (Paper III). Although structures of DGCs and PDEs exist and the processes of synthesising and degrading c-di-GMP are fairly well understood, the mechanisms of downstream targeting have not been as elucidated. Nevertheless, some c-di-GMP binding domains and downstream targets have been discovered.

1.4.4.5 C-di-GMP responsive transcriptional regulators Three major transcription factors – VpsT, Clp, and FleQ – are all responsive to c-di-GMP. In Vibrio cholera, VpsT down regulates motility associated genes and up regulates biofilm matrix ones. For VpsT to be able to exert its control, it needs to undergo c-di-GMP dependent dimerization (Krasteva et al., 2010). Clp is a global transcriptional regulator that directly and indirectly controls many genes associated with diverse cellular functions (e.g. motility and EPS synthesis) in Xanthomonas campestris (He et al., 2007). Binding of c-di-GMP to Clp disrupts the DNA-binding capacity of Clp and thus its ability to up and down regulate genes (Leduc and Roberts, 2009; Chin et al., 2010). FleQ is not only the master regulator of flagella synthesis in P. aeruginosa, it also regulates genes involved in biofilm formation. One operon that FleQ regulates is necessary for formation of Pel exopolysaccharides (Hickman and Harwood, 2008). In the absence of c-di- GMP, FleQ blocks transcription of pel, however, binding of c-di-GMP to FleQ relieves transcriptional repression and converts FleQ to an activator of pel transcription (Baraquet et al., 2012).

1.4.4.6 C-di-GMP dependent riboswitches, RNA processing, and proteolysis Two classes of riboswitches that can bind c-di-GMP have been discovered (reviewed in Ryan et al., 2012). Upstream of some genes coding for DGCs and PDEs, a conserved RNA domain can be found called GEMM (Sudarsan et al., 2008). This type of domain can form two different structures that regulate the expression of their downstream genes in response to c-di-GMP (Sudarsan et al., 2008). A second type of c-di-GMP dependent riboswitch controls a self-splicing ribozyme in Clostridium difficile (Lee et al., 2010). In E. coli, a protein-RNA complex exists that has oxygen dependent c- di-GMP signalling properties (Tuckerman et al., 2011). The complex consists of the two O2 sensors DosC and DosP, which both have a sensing domain 17 coupled to DGC and PDE domains respectively, a polynucleotide phosphorylase (PNPase), several other proteins, and RNA. The Dos proteins respond to the O2 levels and change the amount of c-di-GMP in the vicinity of the PNPase. This accelerates the activity of the PNPase in the presence of the second messenger, which in turn results in different tailing of RNA under anaerobic and aerobic conditions. As described under section 1.4.4.1 The c-di-GMP lifestyle switch, a c-di- GMP dependent proteolysis mechanism controls dispersal of biofilms in P. fluorescences. A model for this has been proposed by Newell et al. (2011): Under biofilm favourable conditions, when the c-di-GMP levels are high, the inner membrane spanning LapD binds c-di-GMP via its degenerate EAL domain in the cytoplasm (Newell et al., 2009). This allows LapD to sequester LapG in the periplasm. Sequestered LapG is hindered from cleaving the surface adhesive protein LapA and the bacteria stay attached to the biofilm. When phosphate levels become limiting in the biofilm, the PDE RapA degrades c-di-GMP, reducing levels of c-di-GMP in the cytoplasm (Monds et al., 2007). With its EAL domain now empty, LapD can no longer sequester LapG, which is therefore free to cleave LapA releasing the bacteria to the surroundings (Navarro et al., 2011).

1.4.4.7 C-di-GMP binding domains Two c-di-GMP binding motifs (in addition to EAL and HD-GYP) have been found in several proteins. These are the I-site of DGCs and the PilZ domain. In enzymatically active DGCs, the I-site binds c-di-GMP and allosterically inhibits further production of the second messenger – this allows for tight control of the c-di-GMP concentrations (reviewed in Schirmer and Jenal, 2009). However, the I-site can also have a role in enzymatically inactive DGCs. For example, the Caulobacter crescentus PopA protein (involved in cell cycle progression) has a degenerate DGC domain that enables correct cellular localization of PopA (Duerig et al., 2009). Membrane bound PelD is another example of a protein with an I-site, although it lacks a DGC-GGDEF domain. PelD is, in addition to FleQ (see section 1.4.4.5 C-di-GMP responsive transcriptional regulators), also able to regulate P. aeruginosa Pel production in response to c-di-GMP signals via the I-site (Lee et al., 2007). The PilZ domains can either exist by themselves or together with other types of domains in multi domain proteins. A common feature of PilZ domains is that they undergo a conformational change when binding c-di- GMP, which leads to altered protein-protein interactions and allosteric effects. In contrast to this common feature, PilZ domains can have different binding modes, stoichiometries, and quaternary structures (Benach et al.,

2007; Ryan et al., 2012). These variations may contribute to different actions by the PilZ domain proteins upon c-di-GMP binding.

1.4.4.8 The flagellar brake protein YcgR E. coli encodes two different PilZ domain containing proteins, one of which is the flagellar brake protein YcgR. YcgR has two domains – PilZN and PilZ. One molecule of YcgR can bind one c-di-GMP dimer via its PilZ domain with high specificity. This binding changes the conformation of YcgR to a more condensed form without changing its monomeric status (Ryjenkov et al., 2006).

Figure 6. Possible mechanism of YcgR in c-di-GMP responsive control of flagella In the absence of c-di-GMP, YcgR binds to FliM in the flagellar switch complex/C- ring, which functions as the rotor of the flagellar motor. Once YcgR binds c-di-GMP, it undergoes a conformational change that leads to interaction with FliG. This breaks interactions between FliG and the stator protein MotA, which leads to lost energy transfer between the stator and the rotor. Adapted from Paul et al. (2010).

YcgR is situated at the flagellar motor where it interacts with FliM and FliG in the C-ring (Boehm et al., 2009; Fang and Gomelsky, 2010; Paul et al., 2010). A model (Fig. 6) has been proposed by Paul et al. (2010), where YcgR interacts with FliM. Once bound to c-di-GMP, YcgR also interacts with FliG. This interaction is proposed to displace the C-terminus of FliG that normally interacts with MotA in the stator, thus preventing energy transfer from the stator to the rotor. YcgR-FliG could also block neighbouring FliG molecules (that are not interacting with YcgR) from switching the rotation direction to CW. YcgR would hence both hinder torque generation by blocking FliG- MotA contacts and lock the motor in a CCW state, hindering the bacterium from changing direction when it encounters obstacles.

2. AIMS How does Pseudomonas putida sense where to go – or if to go at all? Answering this question summarizes the general aim of my research over the last five years. When I started as a graduate student, an initial goal was to elucidate how the P. putida strain CF600 controls its taxis response towards phenolic compounds. The project then unexpectedly took a turn from phenol catabolism sensing to a more general control of motility by c-di-GMP and is currently, as I pass the project on to our newest group member, returning to how P. putida controls its motility in response to environmental cues. During the evolution of my research, the following specific questions came into focus:

− What type of taxis does P. putida use to move towards phenolics?

− Which one of several potential receptors mediates this directed motility response?

− How are the three similar Aer-like receptors of P. putida differentially regulated?

− What are the c-di-GMP turnover properties of the motility-associated PP2258 signalling protein?

− How does c-di-GMP signalling from PP2258 couple to motility control?

3. RESULTS AND DISCUSSION

3.1 Metabolism dependent taxis by Pseudomonas putida CF600 The starting point of my thesis work was to determine the underlying chemotactic mechanism that directs Pseudomonas putida CF600 towards phenolics. P. putida CF600 is a soil bacterium with approximately 6 flagella located at one pole of the bacterium. This P. putida strain harbours a plasmid (pVI150) that allows it to use phenol, monomethylated phenols, and 3,4-dimethylated phenol [hereafter collectively called (methyl)phenols] as carbon sources (Shingler et al., 1989). The enzymes that metabolise these compounds are encoded in the dmp-operon, which is controlled by the transcriptional regulator DmpR that is encoded close by on pVI150. DmpR is an aromatic-responsive protein which requires binding of dmp-pathway substrates or structural analogues (effectors) to take up its active form capable of promoting transcription of the dmp-operon. Consequently, DmpR ultimately controls the capacity of the dmp-system to degrade (methyl)phenols (reviewed in Shingler, 2003).

3.1.1 P. putida CF600 shows metabolism-dependent taxis towards dmp-pathway substrates We started our analysis by elucidate which compounds that serve as chemoattractants for P. putida CF600. Four different types of phenolic compounds were considered: i) (methyl)phenols that are substrates for the dmp-pathway, ii) intermediates of the pathway, iii) compounds that are DmpR effectors but are not pathway substrates, and iv) 2,4-dichlorophenol as a phenolic compound which is neither a DmpR effector nor a dmp- pathway substrate. However, P. putida CF600 only displayed taxis towards the first and second type of phenolics – i.e. compounds that it can metabolize (Paper I, Fig. 2) – providing the first indication that the taxis response was dependent on the bacteria’s catabolism. This was initially surprising to us since other catabolic plasmids involved in metabolism of aromatic compounds encode classical chemotaxis receptors or mediators that facilitate movement towards any compound recognised, irrespective of whether it was subsequently metabolised or not (Grimm and Harwood, 1999; Hawkins and Harwood, 2002). To follow up on this initial indication of metabolism-dependent behaviour, we utilized P. putida KT2440 that lacks the pVI150 plasmid and does not innately exhibit taxis towards phenol or (methyl)phenols. We found that simply introducing the capacity to degrade phenolics (via the dmp- system) likewise rendered this strain capable of taxis responses towards the

21 same range of phenolics as P. putida CF600, and that this phenotype was dependent on the presence of DmpR. Both these findings supported the idea of metabolism-dependent taxis, but did not resolve if DmpR was required per se or simply because it is needed to allow metabolism of phenolics. Therefore, to uncouple catabolism of phenolics from DmpR, we expressed different parts of the dmp-operon using a heterologous promoter. This strategy enabled DmpR-independent growth on phenol and methylated phenols (when expressing the entire suit of enzymes) or just phenol but not methylated phenols (when expressing just the phenol hydroxylase genes). The results confirmed the metabolism-dependent taxis phenotype, i.e. that P. putida KT2440 can only mediate taxis towards phenolics that it can metabolise and that no other gene derived from the pVI150 plasmid was required (Paper I, Fig. 2). In this sense, the pVI150 plasmid differs from other catabolic plasmids for aromatic compounds, such as NAH7 of P. putida G7 and pJP4 of Ralstonia eutropha JMP134, which encode a specific chemotaxis receptor and mediator, respectively (Grimm and Harwood, 1999; Hawkins and Harwood, 2002).

3.1.2 Aer2 is the metabolism-dependent and aerotaxis receptor of P. putida KT2440 The metabolism-dependent taxis response towards (methyl)phenols is coupled to the central chemotaxis system since inactivating the cheA gene abolished the response (Paper I, Fig. 2). Next we wanted to identify which receptor was responsible for signalling to CheA during this response. In addition to mediating aerotaxis, the Escherichia coli Aer receptor is also able to direct the bacterium towards readily oxidizable substrates by metabolism- dependent taxis (Bibikov et al., 1997; Zhulin et al., 1997; Greer-Phillips et al., 2003). This suggested to us that a similar receptor of P. putida may be responsible for the metabolism-dependent taxis seen. Out of 27 predicted MCPs of P. putida KT2440 (Nelson et al., 2002), three have very similar domain architecture as Aer: Aer1-3, although all of them lack the HAMP domain (Paper I, Fig. 3A, also see Fig. 7). We also considered another potential receptor, PP3414, but this protein lacks key features indicative of membrane association and energy-sensing and so I leave it out of the following discussion. To determine which, if any, of the Aer-like receptors plays a role in metabolism-dependent taxis, we created individual null mutations by replacing an internal portion of each gene with a kanamycin (Km) resistance cassette. The Aer2 null strain was the only mutant that showed impaired motility on minimal media soft agar plates with phenol as the carbon source (Δaer2::Km; Paper I, Fig. 3D). Lack of Aer2 also decreased taxis towards other non-phenolic carbon sources implicating it as a general receptor for

22 metabolism-dependent responses of this organism. These defects were readily complemented by expressing aer2 in trans but not by aer1 or aer3, indicating that these receptors may have distinct functions (Paper I, Fig. 3D). Since E. coli Aer mediates aerotaxis responses and O2 is required for catabolism of phenolics by the dmp-system, we also tested if any of the three Aer-like receptors were involved in aerotaxis to guide P. putida through O2 gradients. As with the metabolism-dependent taxis, only the strain lacking Aer2 appeared to be defective in aerotaxis (Paper I, Fig. 3F). Thus, under the experimental conditions tested, Aer2 appears to be the primary receptor for aerotaxis and general metabolism-dependent taxis responses of P. putida KT2440.

Figure 7. Domain architecture of Aer, Aer1-3, and PP2258 and alignment of their PAS domains At the top is a schematic illustration of the predicted domain organisation of E. coli Aer, the three Aer-like receptors of P. putida and PP2258. TMD stands for transmembrane domain. Below is an alignment of the PAS domains of the five proteins. Residues found in all proteins are labelled with stars and residues important for FAD binding are in bold (Repik et al., 2000; Campbell et al., 2010). The alignment was done with Clustal Omega (Goujon et al., 2010; Sievers et al., 2011).

The function(s) of the Aer1 and Aer3 receptors remain an open question. Aer1 and Aer3 may serve similar roles as Aer2 under different experimental conditions. However, lack of Aer2 could only be complemented by a plasmid independently expressing the Aer2 receptor and not by 23 plasmids expressing either Aer1 or Aer3, suggesting that some part(s) of Aer2 may provide unique specificity for its signal transduction role. The regions exhibiting highest similarity between the P. putida Aer-like proteins and Aer of E. coli are the PAS domain and the signalling domain that interacts with CheA. This might indicate that specificity lies in the PAS to transmembrane region or in interactions between these domains. We never constructed hybrid Aer receptors, but such an approach could be one way to dissect the roles of the different domains, which might give clues as to the functions of Aer1 and Aer3.

3.1.3 The three Aer receptors are co-localized but differently expressed It had not been determined if P. putida possesses chemoreceptor polar rafts like those present in E. coli and there are also chemotaxis receptors in other species that have different localizations (reviewed in Krell et al., 2011; Porter et al., 2011). Therefore, to gain further insights to the roles of the three different Aer-like receptors, we studied their cellular localization. To address where the Aer receptors are situated, we individually tagged them with yellow fluorescence protein (YFP). We found that all three Aer receptors are localized to one pole of P. putida and that their localization is dependent on CheA, indicating that P. putida also probably has its taxis receptors arranged in polar rafts (Paper I, Fig. 5). Using an alternative fluorescence protein fusion partner (DsRed), we were also able to show that the three Aer-like receptors co-locate to the same pole. However, we did not investigate if other MCPs encoded in the P. putida genome also localize to this pole. P. putida is a rod shaped lophotrichous bacterium with the flagella located at one end of the cell. One currently unresolved, but intriguing question, is if the receptors co-localize at the same pole as the flagella or if they are positioned at opposite poles. I would think it likely that a mechanistically optimal arrangement would be to have the receptors and flagella in close proximity (i.e. at the same pole). Such an arrangement should minimise the response time from signal-reception by the receptors since phosphorylated CheY would not have to diffuse as far to reach the flagellar motors. When comparing the YFP-signal arising from the different Aer receptor fusions, it was most frequently observed in cell populations expressing Aer2- YFP as compared with Aer1-YFP and especially Aer3-YFP (Paper I, Fig. 5). This indicated that the three aer genes are differentially expressed, a possibility that we further analysed with transcriptional reporters. These reporters showed that all three receptors have different transcription profiles in rich medium (Paper I, Fig. 6). In minimal medium, however, the profiles were more similar but transcription of aer2 was higher than that of aer1 and

24 aer3 (Paper I, Fig. 6). This latter condition is energy limiting for the bacteria and taxis towards carbon sources would be critical in this situation, implying that Aer2 is the most important receptor under this circumstance.

3.2 Transcriptional regulation of the Aer receptors Since the three aer genes are differentially regulated, we thought that dissecting the promoters that drive their expression might give insights to their functions. We therefore analysed aer1, aer2 (in particular), and aer3 transcription both in vivo and in vitro.

Figure 8. Genomic context of the three aer genes Schematic illustration of the aer genes of P. putida. The σ-factors responsible for promoter recognition are depicted as well as identified 5’-UTRs and factors influencing the transcription of the aer genes.

3.2.1 aer2 but not aer1 or aer3 are transcribed by σFliA- RNAP The aer1 and aer3 genes are both located in operons while aer2 is monocistronic (Fig. 8). We could find no obvious promoter immediately upstream for the operons but the DNA upstream of aer2 contains a sequence with two mismatches from the P. putida σFliA promoter consensus (Paper II, Fig. 3). A transcriptomics analysis had also shown that aer2 is down regulated in a σFliA deficient strain (Rodríguez-Herva et al., 2010), which strongly indicated that the Paer2 promoter is transcribed by σFliA-RNAP. We verified this in vivo with aer2 transcriptional reporters, which showed that aer2 transcription is abolished in a P. putida σFliA null strain (Paper II, Fig. 1). A complementary in vitro approach using reagents purified from P. putida showed that σFliA-RNAP promoted transcription from the Paer2 promoter in a concentration dependent manner, while σ70-RNAP only mediated trace levels of transcripts (Paper II, Fig. 2). Hence, expression of the metabolism- dependent and aerotaxis receptor Aer2 is coupled to flagella biosynthesis through dependence on the specialized σ-factor σFliA. Since no obvious promoter could be found for the aer1 and aer3 operons, transcriptional reporters for these two operons were introduced into 13 different σ-factor deficient P. putida strains – among these the σFliA

25 null derivative. In no case was the transcription from the aer1 or aer3 regulatory regions decreased. Although P. putida encodes an additional 10 alternative σ-factors, in silico analysis suggests that they are all probably involved in specialised systems for iron acquisition. Thus, the house-keeping σ70 appeared the most likely candidate σ-factor for the promoters of the aer1 and aer3 operons. Analysis in vivo with the transcriptional reporters in a strain with elevated σ70 levels also increased transcription in both cases and in vitro transcription with σ70-RNAP (but not σFliA-RNAP) gave transcripts from regulatory regions of both operons (Paper II, Fig. S1). In the case of the aer1 operon, subsequent deletion and mutagenesis analysis identified that this operon has a 5´-UTR with the promoter far (~160 bp) upstream of the GTG start codon (Paper III, Fig. S2). Thus, in contrast to aer2, transcription of both the aer1 and aer3 operons appears to be directed by σ70-RNAP.

3.2.2 The P. putida σFliA consensus is suboptimal in vitro Because the σFliA consensus of P. putida differs from that defined for E. coli, we were interested in using the Paer2 promoter to probe the significance of this difference (Paper II, Fig. 3) (Ide et al., 1999; Rodríguez-Herva et al., 2010). Therefore, we extended our analysis to include two additional aer2 promoter derivatives – one with the Paer2 mutated to the P. putida σFliA consensus sequence and one to the E. coli consensus sequence. Since the native Paer2 promoter only differs at two positions relative to the P. putida consensus, the output from both these promoters were unsurprisingly more or less the same (Paper II, Fig. 3). However, more intriguing we found that the E. coli consensus derivative gave over 2-fold more transcripts than P. putida consensus derivative in vitro despite the use of P. putida derived reagents (Paper II, Fig. 3). The P. putida consensus sequence deviates from the E. coli consensus by having a C instead of an A at the penultimate position in the -35 motif: TCAAGT and TAAAGT, respectively. All but one of the eight sequences that were used to derive the P. putida consensus sequence, which included Paer2, possess this C. Transcription dependent on σFliA has been investigated in both E. coli and C. trachomatis and in both organisms it has been shown that substituting the penultimate A to a C decreases promoter output (Yu et al., 2006). Hence, we concluded that the previously identified P. putida σFliA consensus is likely to be sub-optimal for maximal transcription by σFliA- RNAP in this species. The one P. putida σFliA promoter that has a penultimate A is the one for the flgM-PP4396-PP4397 operon. These genes encode the anti-σ-factor FlgM, a FliN-like protein (a chaperon for flagellar hook associated proteins) and a YcgR-like protein (also see 3.4.1 PP4397 is located in a flagella associated operon). The promoter for this operon, in addition to a penultimate A instead of the usual detrimental C, also has only one

26 mismatch from the E. coli consensus (the T in the -10 motif is an A). As such, the PflgM is likely to have close to maximal output – a property that might be important for a feed-back loop to maintain appropriate balance of σFliA- dependent transcription when FlgM is being actively secreted. For the other genes, possession of sub-optimal σFliA promoters would likely permit a appropriate background for transcriptional regulators since promoter output could be both decreased or increased from the basal levels.

3.2.3 The aer2 UP-element controls output and σFliA specificity Rodríguez-Herva et al. (2010) suggested that P. putida σFliA dependent promoters need AT-rich UP-elements in order for recognition by σFliA-RNAP. AT-rich regions upstream of σFliA dependent promoters have also been shown to increase transcription in other species (Fredrick et al., 1995; Shen et al., 2006). We again tested this issue in the context of the Paer2 promoter. The sequence upstream of the aer2 promoter is rich in A and T bases and transcriptional reporters in vivo showed that this region is indeed required for aer2 transcription and further that the promoter activity could be elevated by changing the upstream region to the E. coli UP-element consensus (Paper II, Fig. 4). We also examined the importance of the AT- rich region upstream of Paer2 in vitro. Again we found differences in promoter output when coupled with different upstream regions, although they were not as distinct in vitro as in vivo (Paper II, Fig. 4). This is a common phenomenon that is thought to reflect the fact that there is no competition between σ-factors for core RNAP and the holoenzyme is usually close to saturating for the promoter in the test tube setting. However, the transcriptional reporter with the UP-element consensus gave markedly increased transcription in vivo, a difference that was almost absent in the in vitro transcription analysis. This led us to examine if the σ-factor specificity had been altered when introducing the UP-element consensus. We found that in vitro transcription using σ70-RNAP gave notably higher amount of transcripts when using the template with the UP-element consensus as compared to the native aer2 upstream region (Paper II, Fig. 4). The strong interaction between the α-subunit of the RNAP with the UP-element thus seems to overcome poor binding by σ70 to the promoter sequence. Hence, the large difference seen between the different transcriptional reporters is probably (at least in part) due to a changed σ-factor dependence of the promoter from elusively σFliA to dual σFliA/σ70. The native AT-rich upstream region of the aer2 promoter acts as an UP- element and is thus likely to be engaged by the α subunits of σFliA-RNAP. Our analysis demonstrates that it provides interactions that are strong enough to compensate for the sub-optimal promoter sequence but weak enough to 27 allow for σ-factor specificity. When the upstream regions of the σFliA dependent promoters used to derive the σFliA consensus sequence were analysed, no UP-element consensus could be determined (Rodríguez-Herva et al., 2010). One explanation to this, in the light of the aer2 promoter, is that the composition of the UP-element has evolved to fine-tune the output of these sub-optimal promoters since the promoters themselves are all quite similar, deviating at most by one nucleotide from the consensus.

3.2.4 DksA and ppGpp affect aer2 transcription and motility To further dissect the regulation of Paer2 promoter, we looked at the effects of the global transcriptional regulators ANR, ppGpp and DksA – all three of which have been implicated in control of aer genes in other species. P. aeruginosa has two genes encoding Aer-like receptors, one of which requires the Pseudomonas FNR homologue ANR that is involved in anaerobic stress responses (Hong et al 2004). In P. putida, lack of ANR had no effect on motility or aer2 transcription under the conditions tested (Paper II, Fig. 5). However, these did not include analysis under low oxygen levels. Loss of ppGpp in P. putida resulted in a similar reduced motility phenotype as the Aer2 null mutant, while the absence of DksA had an even more severe effect on the motility (Paper II, Fig. 5). Since both ppGpp and DksA are global regulators, the phenotypes probably reflect both specific and pleiotropic effects. When we looked at promoter activity using transcriptional reporters in DksA or ppGpp deficient strains, Paer2 output was reduced in both mutants, but only in the stationary phase (Paper II, Fig. 5). Since DksA is present at constant levels throughout the growth curve and ppGpp is produced at the exponential-to-stationary phase transition, it seemed likely that DksA exerts its effect together with ppGpp. In vitro transcription from Paer2 was independently stimulated by the addition of ppGpp or DksA alone (Paper II, Fig. 5). However, addition of the two together resulted in the highest level of stimulation, indicating that DksA sensitizes σFliA-RNAP to ppGpp (Paper II, Fig. 5). To my knowledge, Paer2 remains the only example where σFliA-RNAP has been shown to be regulated directly by DksA and ppGpp. In E. coli, transcription of aer is inhibited by ppGpp and DksA (Aberg et al., 2009) under energy limiting conditions and lack of DksA results in increased flagellation and motility (Aberg et al., 2009; Lemke et al., 2009). Contradictory to this, Magnusson et al. (2007) reported that lack of either ppGpp or DksA results in decreased E. coli motility. However, the growth medium used differed between the former and latter research groups, which might underlie the opposing results. Our results with P. putida aer2 are consistent with those of Magnusson et al, and we have also used the same growth medium.

Lemke et al. (2009) showed that in E. coli, genes in the two first levels of the flagellar biosynthesis hierarchy with σ70 dependent promoters are directly regulated by ppGpp and DksA. In this organism the hierarchy is three tiered, however, in P. aeruginosa it has been shown to be four tiered. The master regulators also differ with FlhDC at the top in E. coli and FleQ in P. aeruginosa. Whether fleQ or genes downstream in the Pseudomonas hierarchy are controlled by ppGpp and DksA remains to be elucidated but it is possible that the different life-styles of Pseudomonas and E. coli require that production of their motility apparatus to be regulated differently in response to environmental cues.

3.3 The bi-functional c-di-GMP turnover enzyme PP2258 and motility control Having established that Aer2 is the major metabolism-dependent and aerotaxis receptor and that it is produced simultaneously with the flagella, we next turned our attention to the function of the PP2258 protein. We serendipitously stumbled across the PP2258 gene during our initial deletion approach designed to identify which receptor is responsible for the metabolism-dependent taxis, and found that the PP2258 gene, which is encoded just downstream of aer1, influenced general motility of P. putida. In contrast to the decreased motility found with the Aer2 null strain, the Aer1 null derivative unexpectedly resulted an increased spreading on motility plates as compared with the wild type (Paper I, Fig. 3). Aer1 has 94% identity with Aer of P. putida PRS2000, in which a gene-block insertion leads to decreased spreading on succinate plates (Nichols and Harwood, 2000). We found this discrepancy intriguing and it led us to examine the genomic context of aer1. We found that aer1 and PP2258 are co-transcribed in a bi-cistronic operon (Paper I, Fig. 4). The gene block insertions used by us and Nichols and Harwood (2000) differ: the one we used might allow some transcription of a downstream gene, while the one used for the P. putida PRS2000 aer gene is flanked by strong transcriptional termination sites, so would not. We followed up these observations by generating a single PP2258 null and a double Aer1 and PP2258 null strain. Both derivatives showed reduced spreading on motility plates independent of the growth substrate used (Paper I, Fig. 4). This indicated to us that PP2258 is involved in general motility control rather than a taxis response per se.

3.3.1 PP2258 can both synthesise and degrade c-di-GMP PP2258 is a tripartite protein with a PAS-GGDEF-EAL domain organization, thus possessing two domains with potential to influence c-di-GMP levels in P. putida (Fig. 9). This suggested that the motility control exerted by PP2258 was through c-di-GMP signalling. However, both the GGDEF and the EAL 29 motif of PP2258 lacks complete consensus. Nevertheless, structural modelling revealed that key functional features are present within PP2258 (Paper III, Fig. 1), indicating that both were likely to be enzymatically active. In addition, this modelling also suggested that the PAS domain might be involved in protein dimerization – a property that would be required for diguanylate cyclase activity of the GGDEF domain. To get further insights into the enzymatic properties of PP2258, we generate derivatives with substitutions within key motifs in the context of the full length PP2258 protein and within PP2258 truncates. The motility phenotypes of these mutants when over-expressed in a PP2258 null P. putida strain were correlated with quantification of cellular c-di-GMP levels as determined by two dimensional thin layer chromatography (2D-TLC). Using this approach we could experimentally confirm that both the GGDEF and the EAL domains mediate cognate DGC and PDE activities, and that the DGC c-di-GMP synthesising activity is held in check by a regulatory I-site (Paper III, Fig. 3 and 4). We also found that the GGDEF domain requires the PAS domain for its DGC activity, probably because it facilitates dimerization (Paper III, Fig. 3).

Figure 9. Catalytic activities of PP2258 Schematic illustration of the PP2258 protein and the functions of its two enzymatic domains. Note that the protein needs to form a homodimer for the DGC domain to be active.

3.3.2 The in vivo activity of PP2258 is dependent on its concentration The dual enzymatic potential of PP2258 raises questions concerning which activity is predominant and if the two activities of PP2258 can be modulated. During our analysis we found that the effect of PP2258 depends on its dosage. The null mutant has reduced motility, indicative of elevated c-di- GMP levels, suggesting that PP2258 predominantly functions as a PDE (via its EAL domain) in the wild type setting (Paper III, Fig. 2). However, over- expression of PP2258 leads to vastly increased c-di-GMP levels and

30 decreased motility (Paper III, Fig. 2), suggesting that at sufficient levels its DGC activity predominates. One likely explanation for this is that under this artificial condition PP2258 is forced into a dimeric (DGC competent) configuration. Alternatively, it is also plausible that the artificially elevated levels caused by PP2258 over-expression additionally causes malfunctioning of other c-di-GMP turnover proteins. Any potential switching of the activities of PP2258 would require some form of regulation. The first site where regulation could take place is at the level of transcription. As mentioned previously, the σ70-dependent aer1- PP2258 operon promoter is located approximately 160 bp upstream of the operon start, which creates a 5’-UTR (Paper III, Fig. 5). Since c-di-GMP dependent riboswitches have been found for genes encoding some c-di-GMP turnover enzymes (Sudarsan et al., 2008), we investigated if the 5’-UTR of aer1-PP2258 might have this function. However, we concluded that it does not because neither removing the 5’-UTR nor altering the c-di-GMP levels in the bacterium had any effect on the transcriptional output (Paper III, Fig. 5). During this analysis, we did note that the transcriptional profile from monocopy reporters (Paper I, Fig. 6) differed from that observed with multicopy reporters (Paper III, Fig. S3), suggesting some mode of regulation of the Paer1-PP2258 promoter, but this was likewise independent of c-di- GMP levels. Another layer of regulation might occur at the protein level – either by signal transduction to the PAS domain (as expanded upon in the final section of my thesis), or through direct modulation of the GGDEF domain. Unlike most enzymatically active DGCs, PP2258 does not have the full consensus GGDEF motif but instead has a serine at the first position (SGDEF). Another protein capable of synthesising c-di-GMP (ECA3270) also has a SGDEF motif and Perez-Mendoza et al. (2011) speculated that the serine could be subjected to regulation by e.g. phosphorylation. One bifunctional enzyme involved in c-di-GMP turnover that has been reported to be regulated by phosphorylation is the Legionella pneumophila Lpl0329 (Levet-Paulo et al., 2011). However, in the case of Lpl0329, it is phosphorylation of its response regulator domain that alters the DGC activity and not phosphorylation of the GGDEF domain directly. P. putida encodes 14 potential serine/threonine/tyrosine kinases (STYKs) (Krupa and Srinivasan, 2005), and it would be interesting to see if one or more of these are involved in regulating the DGC activity of PP2258. However, a P. putida phosphoproteome analysis performed by Ravichandran et al. (2009) did not identify PP2258 or any other c-di-GMP turnover enzymes.

3.4 PP4397 – the link between the c-di-GMP signal and altered motility The above results established that PP2258 is a c-di-GMP signalling protein of P. putida, however, the link between PP2258 and motility control of the flagella was still missing. In E. coli and S. enterica, the PilZ containing protein YcgR has been shown to bind c-di-GMP and alter interactions in the flagellar motor, thereby decreasing torque generation and promoting a CCW rotation bias. The P. putida protein that is most similar to YcgR is called PP4397 and shows 22% and 24% identity to E. coli and S. enterica YcgR, respectively. PP4397 has been studied in silico and in vitro were it has been shown that it can bind c-di-GMP and undergoes a dimer to monomer transition upon this binding (Ko et al., 2010). Matilla et al. (2011) looked at the phenotype of a PP4397 transposon insertion mutant in a high c-di-GMP background. They did not see any alteration compared with the wild type for many different c-di-GMP influenced traits. However, they did not look at the motility phenotype (or did not report their results if they did). PP4397, therefore, remained our top candidate as the protein responsible for transferring the PP2258 c-di-GMP signal to the flagellar motor.

Figure 10. Genomic context of PP4397 Schematic illustration of the operon in which PP4397 is situated. Transcription from both the σ54 and the σFliA promoters result in transcripts with PP4397. Adapted from Paper IV; Osterberg et al. (2013).

3.4.1 PP4397 is located in a flagella associated operon What we first noticed when considering PP4397 as the potential link was that the genomic context in which its gene is situated differed from that of E. coli and S. enterica ycgR. Unlike ycgR (which is monocistronic), PP4397 seemed to be located in an operon together with the σFliA anti σ-factor gene flgM and a gene encoding a protein belonging to the FlgN family (FlgN is a chaperon for flagellar hook associated proteins). This genomic organization was experimentally confirmed and in addition to these three genes (flgM, PP4396 and PP4397), which are controlled by a σFliA-dependent promoter, a fourth gene (flgA, encoding a protein involved in flagellar basal body P-ring formation) upstream of flgM was also co-transcribed from a σ54-dependent 32 promoter (Paper IV, Fig. 1, also see Fig. 10). This organisation of flgA, flgM and flgN controlled by dual promoters can also be found in Salmonella but in this case no additional gene encoding a PilZ protein is present (reviewed in Chilcott and Hughes, 2000).

3.4.2 PP4397 is downstream of PP2258 in the signalling pathway The coupling of PP4397 transcription with that of flagellar associated genes further strengthened our hypothesis that PP4397 is involved in motility control. To test if PP4397 was downstream of PP2258 in the c-di-GMP signalling pathway, a null PP4397 mutation (ΔPP4397::Tc) was introduced in both the wild type and the PP2258 null strain (ΔPP2258::Km). We could not see any motility difference between the wild type and the PP4397 single mutant but the reduced motility phenotype of the ΔPP2258 single mutant was rescued by a ΔPP2258::Km ΔPP4397::Tc double mutant (Paper IV, Fig. 2). The ΔPP2258::Km reduced motility phenotype was partially restored by introducing PP4397 into the double mutant, but not variants incapable of binding c-di-GMP (Paper IV, Fig. 3). This showed that PP4397 acts downstream of the c-di-GMP signal from PP2258. If PP4397 is the direct link between the c-di-GMP signal and the flagellar motor is left to be determined. If this is the case then PP4397 should co-localize with the flagella. On-going approaches to determine if this is the case are fluorescence microscopy studies, co-immunoprecipitation experiments and two-hybrid systems.

3.5 Possible Aer1-PP2258-PP4397 signalling mechanisms and future perspectives During our work over the last years we have identified some of the multitude of components that regulates P. putida motility. We started off by looking at three Aer-like receptors and found Aer2 to be the major metabolism- dependent and aerotaxis receptor, and that it is i) produced together with the flagella, and ii) that its expression is stimulated under conditions of nutrient scarcity (Papers I and II). However, we have not been able to identify the signals sensed by Aer1 or Aer3 (Fig. 11 Q1). Are they also sensing energy fluctuations inside the cell as Aer2 but under different circumstances? Or are they perhaps sensing surfaces like WspA in P. aeruginosa? WspA is the MCP in the alternative cellular function (ACF) chemotaxis complex called Wsp. WspA is not polar localized like classical MCPs and the Aer receptors of P. putida, because of its weak trimer interactions. This property, which is mainly due to its C-terminal region located after the HAMP domain, enables WspA to sense surfaces and cell-cell interactions (Guvener and Harwood, 2007; O'Connor et al., 2012). The P. putida Aer1 and Aer3 receptors both 33 have approximately 30% identity to the C-terminal region of WspA and about 40% identity to the same part of the E. coli Aer, suggesting that they perhaps are more likely to function as energy receptors than surface sensors.

Figure 11. Current model and open questions Illustration of the herein identified components that regulate P. putida motility. For simplicity, only one out of approximately six flagella is depicted. It is also unclear at what pole the Aer-like receptors are localized – it might well be the same as the flagella. Questions: Q1. What do Aer1 and Aer3 sense? Q2. Are signals sensed by Aer1 and/or Aer3 transferred to the central chemotaxis system? Q3. Do Aer1 and PP2258 interact? Q4. Can the PP2258 PAS domain detect a signal? If so, what kind of signal? Q5. Is PP4397 directly interacting with the flagellar motors? If so, with what protein(s)?

Although we do not know the signals that Aer1 and Aer3 sense, it seems like they are able to signal through the classical chemotaxis pathway because of their polar localization via CheA (Fig. 11 Q2; Paper I). An alternative possibility is that they are only held in place by CheA and that they do not change the autophosphorylation activity of this kinase. Another potential interaction partner for Aer1 is the c-di-GMP turnover enzyme PP2258 (Fig. 11 Q3). The genes for these two proteins are localized in the same operon and thus simultaneously transcribed (Paper I), indicating that their functions are likely to be interconnected. Since both Aer1 and PP2258 possess PAS domains and this type of domain is known to be involved in protein-protein interactions, one possibility is that Aer1 transfers a sensed signal to PP2258 via the PAS domains. Aer1-PP2258 interactions could render PP2258 incapable of forming homodimers, something that is needed for DGC activity and which we believe is dependent upon the PP2258 PAS domain (Paper III). This would lead to the PP2258 EAL domain activity being dominant and PP2258 would thus function as a PDE, degrading c-di-GMP.

In P. aeruginosa, the Aer-like protein BdlA has been shown to interact with the DipA enzyme (Petrova and Sauer, 2012a). DipA has a GAF-PAS- GGDEF-EAL domain organization but only displays PDE activity (Roy et al., 2012). Interactions between BdlA and DipA stimulate the DipA PDE activity and promote biofilm dispersal. This signalling network has recently been shown to be regulated in several different ways (Petrova and Sauer, 2012b). Petrova and Sauer (2012b) suggest a model where BdlA is inactive in planktonic P. aeruginosa. Upon biofilm formation and elevated c-di-GMP levels, BdlA is phosphorylated and its amino-terminal PAS domain cleaved of by ClpP. The PAS domain then re-associates with BdlA, possibly with aid of ClpD. BdlA is now capable of responding to conditions inducing biofilm dispersion, which it does by increasing the PDE activity of DipA. This leads to decreased c-di-GMP levels and dispersal of the biofilm. The decreased c- di-GMP levels also results in BdlA returning to an inactive form. It is tempting to speculate that the Aer1-PP2258 signalling system might have some similarities to the BdlA-DipA one. An alternative to Aer1 being the receptor that transfers a signal to PP2258, is that the PAS domain of PP2258 itself can sense and modulate the c-di-GMP turnover activity of the enzyme (Fig. 11 Q4). The PP2258 PAS domain lacks important residues needed for FAD-binding (Fig. 7) so any potential signal sensed by PP2258 is probably not the same as the one sensed by Aer1 (which appears to be FAD-binding, see Fig. 7). Independent of the regulation of PP2258, the c-di-GMP produced by PP2258 is bound by the YcgR-like protein PP4397, which affects the flagella – either directly or indirectly. Since PP4397 seems to have a different c-di-GMP binding mode than YcgR, it would be interesting to find out if PP4397 binds and affects the flagellar motors in the same way as YcgR (i.e. via FliM and FliG). YcgR not only decreases the energy transferred between the flagellar stator and motor, it also seems to lock the flagella in a CCW rotation state. Since taxis receptors also affect the rotational direction of flagella, the Aer1 receptor and PP2258 via PP4397 might have synergistic effects on the flagella rotation direction either in response to one type of signal (if Aer1 signals to PP2258) or in response to different signals (if PP2258 has sensing capabilities in itself). Although the work I present here has given us a better understanding of P. putida motility control, perhaps the most intriguing question left to be answered is under what circumstances P. putida stops moving.

ACKNOWLEDGEMENTS First of all I would like to thank my supervisor Vicky. Thanks for all your help and guidance through the years! Your memory is better than an elephant’s when it comes to experiments done – someone once called you Vickypedia and I think that is a very fitting nickname. In addition to your admirable scientific thinking, I also appreciate your organisational skills which make life in the lab so much easier! Life in the lab and office wouldn’t have been the same without my fellow co-workers. Eleonore you are a rock and always willing to help with everything! I have enjoyed working with you and all the talks we have had. Teresa, I have really liked our conference trips, thanks for letting me be a control freak and plan most of them :) Linda H, you left big shoes to fill when you moved to the states and I did my best to fill them. You are a true scientist and will go far! Anjana, you will now be the senior PhD student in the group and you will do great! Take good care of Sanodji :) Lisa, I happily leave my project to you. With your enthusiasm you will take it far. And even though I won’t see you at work, you are always welcome to come and visit your neighbour! Maybe we can do that baking sometime. Thanks also to all people who have come and gone during my years in the lab, especially Sofie, Karina, and Anna, whom I shared projects with. Thanks also to everybody else at the department for making it a nice place to spend six years in. Thanks all of you who have shared the lunchtime with me. Many, many, thanks to you who have made my working life so much easier: Johnny, the lab-service ladies, the administrators, and Jessica and Marianella. What would we do without you? Before I started my PhD I had some amazing class mates who made my undergraduate studies to a great time. Some of you are still a part of my little Umeå family. Annika, who came with me from Sundsvall. Hope you enjoy your new little family and the big house of yours. Thank you for not moving to Sävar but staying within walking-distance ;) Annasara and Andréas, as well as your girls, for always caring and giving us nice dinners and fika! Marie and John with your boys, it’s always nice to visit you and your fika isn’t too bad either ;) I miss our board game nights! Linda W, for always being easy to talk to and arranging the pyssel evenings at work. And girls: we need to have syjunta again – soon! Jag skulle aldrig ha hamnat i Umeå om det inte hade varit för min käre bror Patrik, så tack Bossa för att du är den bästa bror man kan tänka sig och för att du alltid tar hand om mig! Jag älskar inte bara dig utan resten av din familj också. Lisa och Elin är precis lika smarta som du, om inte smartare, och de finaste brorsdöttrar man kan tänka sig! Mamma Lena och pappa Christer, vad skulle jag göra utan er?! Ni är de bästa föräldrar man kan ha

36 och jag hade inte varit särskilt morsk om det inte hade varit för att ni alltid finns där och stöttar mig. Allra sist vill jag tacka min fina lilla familj! Min älskade Reine för att du förstår mig bättre än någon annan, inklusive mig själv. Tack för att du alltid är vettig och vet hur du ska få mig att må bra. Jag vet inte vad jag skulle göra om jag inte hade dig! Och Rasmus, mammas älskade lilla gogubbe. Gud vad tråkigt livet hade varit utan dig! Tack för att du fick mig att tänka på något annat än avhandlingsskrivandet och för alla fina kramar och skratt som jag får varje dag!

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