Análisis genético y molecular del operon std de salmonella enterica

! ! ! Genetic!and!molecular!analysis!of!the!std!operon!! in!Salmonella!enterica! ! ! Lucía García Pastor Marzo 2017 Genetic!and!molecular!analysis!of!the!std!operon!! in!Salmonella!enterica! ! ! ! Memoria elaborada por Lucía García Pastor para optar al grado de Doctor en Biología Tesis realizada bajo la dirección del Dr. Josep Casadesús Pursals y la Dr. Elena Puerta Fernández en el Departamento de Genética de la Facultad de Biología de la Universidad de Sevilla Josep Casadesús Pursals Elena Puerta Fernández Lucía García Pastor Sevilla, Marzo de 2017 A mi familia, a Guille y a los BACT “Ve hasta donde te alcance la vista. Cuando llegues, serás capaz de ver más allá”. TOMAS CARLYLE INDEX RESUMEN ........................................................................................................................................... 13! INTRODUCTION ................................................................................................................................ 19! The genus Salmonella .............................................................................................................. 21! Evolution of Salmonella pathogenesis ..................................................................................... 22! Salmonella infection ................................................................................................................ 23! Salmonella virulence factors .................................................................................................... 25! Type III secretion systems (TTSSs) ............................................................................ 25! Flagellum and chemotaxis .......................................................................................... 28! The Salmonella enterica virulence plasmid ................................................................ 29! Adhesins ...................................................................................................................... 31! The std fimbrial operon ............................................................................................................ 33! HdfR ......................................................................................................................................... 35! RcsCDB phosphorelay system ................................................................................................. 36! Dam methylation ...................................................................................................................... 37! Phase variation systems regulated by Dam methylation .......................................................... 40! pap ............................................................................................................................... 40! agn43 ........................................................................................................................... 41! opvAB .......................................................................................................................... 41! Phenotypic heterogeneity in bacteria population ..................................................................... 42! OBJECTIVES ....................................................................................................................................... 45! MATERIALS AND METHODS ......................................................................................................... 49! Bacterial strains and strain construction .................................................................................. 51! Construction of the relevant strains ............................................................................ 58! Bacteriophages ......................................................................................................................... 59! Culture media and growth conditions ...................................................................................... 60! Bacterial transduction .............................................................................................................. 61! P22 lysates ................................................................................................................... 61! Transduction in liquid medium ................................................................................... 61! Detection of lysogenic transductants .......................................................................... 61! P22 sensitivity assay ................................................................................................... 62! DNA manipulation and transfer ............................................................................................... 62! Plasmids ...................................................................................................................... 62! La metilación Dam es el principal represor de la expresión del operón std, pero existen otros represores previamente descritos como RosE o SeqA. En colaboración con el grupo de la Dr. Delgado del Instituto Superior de Investigaciones Biológicas (INSIBIO) CONICET-UNT en Argentina, se ha identificado un nuevo represor de la expresión de std, RcsB. La expresión constitutiva de rcsB reprime la expresión de std en fondo Dam– y también en fondo silvestre. Se han realizado ensayos de retardo en gel para corroborar la unión de RcsB a la región promotora de std. El análisis bioinformático identifica un posible sitio de unión de RcsB en la región -35 del promotor de std. Con el fin de identificar otros posibles reguladores de la transcripción del operón std, se realizaron escrutinios genéticos. Los resultados obtenidos mostraron una autorregulación positiva del operón std, de la que son responsables los genes distales del operón std: stdE y stdF. Ensayos realizados mediante citometría de flujo corroboran los resultados del escrutinio. La expresión de stdE y stdF mantiene la subpoblación StdON y una expresión constitutiva de stdE y stdF aumenta el tamaño de dicha subpoblación. La expresión bimodal del operón std en fondo silvestre nos llevó a pensar en la existencia de patrones de metilación distintos para cada una de las subpoblaciones. Para demostrar la existencia de dichos patrones de metilación se han realizados ensayos de Southern Blot tras digestión del DNA con isosquizómeros de enzimas de restricción que reconocen la misma secuencia en el DNA (GATC) pero difieren en su sensibilidad al estado de metilación de la diana. Los tres sitios GATC de la región reguladora se hallan metilados cuando el promotor está inactivo (la subpoblación StdOFF es mayoritaria en un fondo silvestre). Resultados de citometría de flujo sugieren un patrón de metilación distinto para la subpoblación StdON, ya que la hipermetilación disminuye dicha subpoblación. Anteriormente se había descrito que StdE y StdF controlan la expresión de hilD a nivel postranscripcional. Teniendo en cuenta que nuestros resultados indican que StdE y StdF son reguladores autógenos de la transcripción de std, nos planteamos la posibilidad de que estas proteínas tuvieran un papel regulador. Con objeto de identificar posibles dianas de regulación del regulón StdEF se realizó un análisis transcriptómico usando un microarray de S. enterica serovar Typhimurium. Los datos obtenidos nos han permitido identificar a StdE y StdF como reguladores globales de la transcripción en Salmonella, ya que no sólo reprimen la isla de patogenicidad 1 (SPI-1) sino también genes flagelares y de quimiotaxis. Por otra parte, StdE Resumen 17 y StdF activan la expresión de genes del plásmido de virulencia y del gen hdfR, que codifica el factor de transcripción esencial para la expresión del operón std. Para tratar de determinar el mecanismo regulador de estas proteínas, hemos realizado un ensayo de ChIP-seq (inmunoprecipitación de cromatina seguida de secuenciación masiva), que permite identificar sitios de unión al DNA de las proteínas analizadas. Los resultados obtenidos confirman el papel de StdE y StdF como reguladores transcripcionales de la expresión en Salmonella. StdE parece tener un papel principal, uniéndose a regiones promotoras del DNA. Por otro lado, los resultados del ChIP-seq sugieren un papel secundario o accesorio de StdF en la regulación, quizá favoreciendo la actuación de StdE. El análisis global de la unión de StdE y StdF al genoma de Salmonella ha proporcionado información acerca de la autorregulación del operón std. Como ya se ha mencionado, StdE y StdF favorecen la expresión de std, aumentando el tamaño de la subpoblación StdON. Según los datos del ChIP-seq, es StdF quien se encuentra en la región promotora, favoreciendo la expresión de std. Queda por determinar si StdF tiene un papel principal en la regulación de std o, como en otros sitios de regulación, actúa de modo auxiliar, tal vez favoreciendo la unión de HdfR o impidiendo la actividad de la metilasa Dam. Todo parece indicar que nos encontramos frente a un sistema de regulación global pero con expresión bimodal. La expresión del operón std, con la consiguiente formación de fimbrias, tiene lugar en una pequeña fracción de la población en condiciones de laboratorio, y mayoritariamente en condiciones de adherencia al epitelio intestinal. En dichas condiciones, la bacteria mantiene reprimidos los genes implicados en virulencia, expresión del flagelo o quimiotaxis, y los genes reguladores del operón std son los encargados de coordinar estos procesos. Por otro lado la expresión del operón std favorece la activación del sistema conjugativo del plásmido de virulencia y garantiza el mantenimiento del mecanismo autorregulador de expresión del propio operón std al activar el factor de transcripción HdfR y al ejercer de reguladores transcripcionales uniéndose a la región promotora del propio operón std. INTRODUCTION Introduction 21 The genus Salmonella The genus Salmonella includes facultative anaerobic Gram-negative, rod-shaped bacteria that are able to infect a variety of animal hosts including amphibians, reptiles, birds and mammals. Salmonellae are usually motile and produce peritrichous flagella. Salmonella belongs to Enterobacteriaceae family in the γ-proteobacteria subdivision and is a close relative of the genera Escherichia, Shigella and Citrobacter. Currently, the genus Salmonella is divided into 2 species, known as Salmonella enterica and Salmonella bongori (Tindall et al., 2005). Salmonella enterica comprises 6 subspecies (McQuiston et al., 2008): enterica (subsp. I), salamae (subsp. II), arizonae (subsp. IIIa), diarizonae (subsp. IIIb), houtenae (subsp. IV) and indica (subsp. VI). Salmonella subspecies are further divided into serovars (also known as serotypes). Serovars are defined by the antigen properties of the polysaccharide chain of LPS (O-antigens) and of the proteinaceous flagella (H antigens) (Grimont & Weill, 2007; McQuiston et al., 2004). There are more than 2,500 Salmonella serovars, most of them belong to the subsp. enterica (Popoff et al., 2004). Only serovars of this subspecies are able to colonize warm-blooded vertebrates (McClelland et al., 2001), accounting for 99% of human infections caused by Salmonella, while the remaining subspecies of Salmonella enterica and members of Salmonella bongori are usually associated to cold-blooded vertebrates or to the environment (Bäumler et al., 1998). Differences in host specificity and types of diseases produced are found among serovars belonging to subsp. enterica. Some serovars are able to infect a wide variety of animals, while others are host-specific (Bäumler & Fang, 2013). Serovars of subsp. enterica produce a variety of diseases that ranges from self-limiting gastroenteritis to life-threatening systemic infection, and the outcome of the disease depends on the specific serovar-host combination. For example, the human-restricted serovar Typhi produces typhoid fever, while the generalist serovar Typhimurium produces mild gastroenteritis in humans but causes a systemic infection similar to human typhoid fever when infecting mice (Velge et al., 2012). For this reason, the interaction between serovar Typhimurium and mice has been widely used as a model for human typhoid fever (Santos et al., 2001a), and most pathogenicity studies in Salmonella have employed this serovar. In this work we have used the mouse-virulent strain Salmonella enterica subsp. enterica serovar Typhimurium SL1344, from the B.A.D. Stocker collection. For simplicity, it will be often abbreviated as Salmonella typhimurium. Evolution of Salmonella pathogenesis Salmonella is a close relative of Escherichia. They diverged 120-160 million years ago (Ochman & Wilson, 1987). Almost 25% of the genetic material in the Salmonella genome is absent in Escherichia coli (McClelland et al., 2001; Porwollik & McClelland, 2003). The evolution of Salmonella pathogenicity (Figure I. 1) has involved the sequential acquisition of genetic elements, each contributing to different aspects of its lifestyle (Groisman & Ochman, 1997; Kelly et al., 2009). Among those elements are the Salmonella pathogenicity islands (SPIs), which are clusters of virulence genes in the chromosome. More than 10 SPIs have been described (Hensel, 2004) including some that are serotype-specific. These regions usually have a different G+C content than the rest of the Salmonella chromosome, and are absent in other Enterobacteriaceae, suggesting that they have been acquired by horizontal gene transfer (Kelly et al., 2009; Porwollik & McClelland, 2003). Figure I. 1 Phylogeny of the genus Salmonella. The acquisition of SPI-1 and SPI-2, and the ability to infect warm-blooded vertebrates are indicated. Modified from (Ellermeier & Slauch, 2006). subsp. enterica (I) Escherichia coli Salmonella bongori subsp. houtenae (IV) subsp. arizonae (IIIa) subsp. diarizonae (IIIb) subsp. salamae (II) subsp. indica (VI) Salmonella enterica SPI-1 SPI-2 Infection of warm-blooded vertebrates Introduction 23 Salmonella pathogenicity island 1 (SPI-1) and Salmonella pathogenicity island 2 (SPI-2) are the best characterized SPIs. SPI-1 was first acquired by the common ancestor of Salmonella bongori and Salmonella enterica, and is involved in the invasion of epithelial cells in the animal intestine (Que et al., 2013). SPI-1 acquisition provided Salmonella the ability to invade eukaryotic cells and to become an intracellular pathogen, likely associated with coldblooded vertebrates (Ellermeier & Slauch, 2006). SPI-2 allows Salmonella to survive in macrophages and colonize deeper tissues (Ochman et al., 1996), and its acquisition resulted in the split of the two Salmonella species (Ellermeier & Slauch, 2006). Consequently, only members of Salmonella enterica are able to reach deep tissues and organs producing systemic infection. The ancestor of subsp. enterica acquired the capacity to infect warm-blooded vertebrates, and different strains subsequently evolved to colonize a variety of hosts. Although the mechanism of host specificity is not fully understood, the presence of a virulence plasmid in certain serovars of subsp. enterica has suggested the potential contribution of plasmid functions (Bäumler et al., 1998). Another trait that may be involved in host specificity is the presence of different sets of fimbrial operons in different serovars (Bäumler et al., 1998; Townsend et al., 2001). Salmonella infection Salmonella enterica is found mostly in the intestine of animal hosts and the infection is usually transmitted by the fecal-oral route. Infection normally starts via ingestion of contaminated water or food. Salmonella must endure adverse conditions along the digestive track, which serves as a protective barrier against bacterial infections. The acid pH in the stomach destroys the majority of microorganisms (Tennant et al., 2008). However, Salmonella responds with the activation of the acid tolerance response, a complex adaptive system, which allows the bacterial population to endure periods of severe acid stress (Foster & Hall, 1990; Lee et al., 1995). High concentrations of bile are secreted in the small intestine, in particular in the duodenum, during digestion. Bile has two main antibacterial actions: as a detergent disrupting the cell envelope (Gunn, 2000) and as DNA damaging agent producing DNA rearrangements and point mutations (Prieto et al., 2004). Nevertheless, like other enteric bacteria, Salmonella is intrinsically resistant to high concentrations of bile (Gunn, 2000). Once in the distal small intestine, Salmonella has the ability to penetrate inside epithelial cells. In a first step, adhesins and fimbriae are necessary to mediate the adherence (Wagner & Hensel, 2011); afterwards the bacterial population is able to invade the intestinal epithelium through three different routes (Figure I. 2): (i) by inducing a phagocytosis-like process in non-phagocytic enterocytes, (ii) through specialized epithelial M cells, and (iii) through dendritic cells that intercalate epithelial cells by extending protrusions into the gut lumen (Finlay & Brumell, 2000; Grassl & Finlay, 2008). The two first routes are mediated by the virulence-associated type III secretion system encoded on Salmonella pathogenicity island 1 (SPI-1) (Zhou & Galán, 2001), with invasion of M cells being the predominant route of intestinal traversal (Watson & Holden, 2010). Figure I. 2 Diagram of Salmonella infection. The three main routes of Salmonella invasion of the intestinal epithelium are represented: direct invasion of intestinal epithelial cells (1), adhesion and translocation through M cells (2), and capture by dendritic cells (3). Adapted from (Fàbrega & Vila, 2013; Sansonetti, 2004). Introduction 25 Depending on the serovar and the host, Salmonella infections may have three different outcomes after invasion of intestinal epithelial cells: gastroenteritis, systemic infection, or asymptomatic chronic carriage. In gastroenteritis, the infection is localized in the intestine. Invasion of intestinal epithelial cells triggers an inflammatory reaction in the intestinal mucosa. Liquid accumulation in the intestinal lumen leads to diarrhea (Nunes et al., 2010; Santos et al., 2001b). In systemic infection, the pathogen crosses the epithelial barrier and is able to survive inside phagocytes due to functions encoded in SPI-2. Salmonella disseminates through the lymphatic system reaching deep tissues and colonizing target organs, particularly the spleen, the liver, the gall bladder and the bone marrow, where bacteria can proliferate, eventually causing death upon septic shock (Bäumler et al., 2011). A fraction of individuals that has survived systemic infection becomes asymptomatic, lifelong carriers of Salmonella, acting as reservoirs for future infections. In humans, serovar Typhi can undergo chronic carriage in the gall bladder (Bäumler et al., 2011). Salmonella virulence factors The interaction between Salmonella and the host requires a complex bacterial machinery to colonize and succeed in a variety of host environments, all of them challenging. This machinery includes genes required for chemotaxis, motility, adhesion, invasion, replication and survival within host cells covering the whole pathogenic process from the intestinal step to systemic dissemination. The virulence factors employed by the pathogen to achieve the infection process are described below. Type III secretion systems (TTSSs) Type III secretion systems (TTSSs) are specialized organelles whose main role is to inject bacterial proteins, called effectors, into eukaryotic cells (Cornelis & Van Gijsegem, 2000; Galán, 1999; Galán & Collmer, 1999), representing the principal virulence factors for Salmonella pathogenicity (LaRock et al., 2015). TTSSs are evolutionarily related to the flagellar export apparatus and are present not only in bacteria pathogenic for animals and plants but also in symbionts for plants or insects (Galán, 2001). Non-fimbrial adhesins This category comprises an heterogeneous group of proteins either secreted via type I secretion systems (SiiE and BapA) or autotransported across the bacterial membranes (ShdA, MisL, SadA) (Wagner & Hensel, 2011). Fimbrial adhesins Among the adhesion systems, fimbrial adhesins include the majority of Salmonella adhesins. Fimbriae are proteinaceous structures extending form the surface of a bacterial cell, and appear in several copies per cell. These structures facilitate adhesion to abiotic and biotic surfaces. Three pathways for the assembly of fimbrial adhesins have been described. I. Chaperon-usher dependent assembly Pathway (CUP). Fimbrial subunits are directed to the periplasm through the general secretion pathway via an N-terminal secretion sequence that is cleaved off during transport. In the periplasm, chaperones prevent the premature assembly of fimbrial subunits and their degradation by proteases, and direct fimbrial subunits to the usher, which coordinates the assembly of the fimbriae. On the tip of the completed fimbriae, a receptor-binding site interacts with its host receptor. Based on the usher protein (the most conserved fimbrial protein), fimbriae belonging to the chaperone-usher pathway can be divided into 6 groups (γ, κ, π, β,  and σ) (Nuccio & Bäumler, 2007). Several chaperon/usher-type fimbrial operons present in S. enterica serotype Typhimurium belong to this group. II. Nucleator-dependent assembly pathway. The fimbrial subunits are transported into the periplasm via the general secretion pathway. Polymerization of the fimbrial subunits occurs outside of the bacterial cell envelope (Wagner & Hensel, 2011). In Salmonella, the fimbrial adhesins assembled through this pathway are designated as “thin aggregative fimbriae” or “curli”, and are encoded by the operons csgDEFG and csgBA. These fimbriae are known to mediate binding to proteins like fibronectin (Olsén et al., 1989) and are involved in biofilm formation (Brombacher et al., 2003; Gerstel & Römling, 2003; Prigent-Combaret et al., 2001) III. Assembly pathway for type IV fimbriae. In S. enterica, type IV pili are only encoded by SPI-7 in the strictly human-adapted serovar Typhi (Zhang et al., 1997; 2000). The subunits for the pilus formation are assembled in the periplasm at the inner membrane, and Introduction 33 are extruded through a secretin in the outer membrane as an intact pilus (Wolfgang et al., 2000). As it was mentioned before, fimbriae expression involves crosstalk and participates in the coordination of the sequential expression of SPI-1, flagellar and fimbrial systems. Nonfimbrial adhesins are also subject to crosstalked regulation, as it happens with the SPI-4- encoded sii operon which is regulated by the SPI-1 SprB regulator (Saini & Rao, 2010). Hybridization analyses have shown high presence of fimbrial DNA sequences in multiple Salmonella serovars. Additionally, each serovar harbors a particular repertoire of fimbrial genes (Townsend et al., 2001). The selective pressure responsible for this uneven distribution is unknown (Humphries et al., 2001) and several tentative explanations have been proposed. On top of their role in attachment, it has been shown the contribution of fimbriae to biofilm formation (Ledeboer et al., 2006). Competitive infection experiments have revealed the participation of fimbriae in long-term colonization in mice (Weening et al., 2005). Each fimbrial operon appears to attach a specific target cell type (Bäumler et al., 1996), enabling the bacterial cell to distinguish among different epithelial cell types. Redundancy of fimbrial tools may counterbalance the loss of a single fimbrial operon, since the inactivation of an individual fimbrial operon results in only moderate attenuation (van der Velden et al., 1998). On the other hand, fimbrial appendages could, at the same time, pose a problem for the bacterium to evade the host immune system, since the fimbrial subunits likely act as antigens during host-pathogen interaction. This aspect could explain why fimbrial expression is commonly regulated by phase variation, generating fimbriated and non-fimbriated subpopulations (Humphries et al., 2001). Phase variation will be described below in more detail. The std fimbrial operon In S. enterica serotype Typhimurium, the elaboration of fimbriae on the surface of cells grown in nutrient broth and their ability to agglutinate yeast or red blood cells, was described in 1966 (Duguid et al., 1966). So far, laboratory-grown cultures of S. enterica have only been shown to elaborate type 1 fimbriae (Duguid et al., 1966), encoded by the fim operon (Clegg et al., 1987), and thin curled fimbriae (Grund & Weber, 1988; Stolpe et al., 1994), encoded by the csg operon (Römling et al., 1998). However, nine additional fimbrial operons, named bcf (Tsolis et al., 1999), stf (Emmerth et al., 1999; Morrow et al., 1999), saf (Folkesson et al., 1999), stb, stc, std, sth, sti, and stj (McClelland et al., 2001), have been later identified by sequence analysis. The std fimbrial operon was first identified in S. enterica serovar Typhi (Townsend et al., 2001) and later found in other Salmonella enterica serovars, including Typhimurium (Anjum et al., 2005; Chan et al., 2003; Humphries et al., 2001; Porwollik et al., 2004). However, it has not been detected in related enterobacterial species, suggesting an acquisition mediated by horizontal transfer (Porwollik et al., 2002). The std operon belongs to the π-fimbriae group. Members of the same group are well-characterized virulence factors such as pyelonephritis-associated (P) fimbriae of E. coli and the Mannose-resistant/Proteus-like (MR/P) fimbriae of Proteus mirabilis (Nuccio & Bäumler, 2007). Std fimbriae contribute to cecal colonization by binding to α (1,2) fucose residues, which are abundant in the cecal mucosa (Chessa et al., 2009), facilitating Salmonella attachment to the intestinal epithelium. Expression of the std operon, as many of the putative fimbrial operons, is tightly repressed under laboratory conditions (Humphries et al., 2003), and derepression occurs in the intestine of infected animals, where Std fimbriae are synthesized. Deletion of stdAB results in a competitive defect in colonizing the caecum of mice and in being shed with the faeces (Weening et al., 2005). Results from adhesion assays of Salmonella enterica serovar Enteritidis to intestinal epithelial cells as well as to the small intestine and caecum of poultry reveal decreased adhesion of the stdA mutant (Shippy et al., 2013). Recently, the std operon (Figure I. 6) was defined to contain 6 genes (stdA-F), all of them transcribed by the promoter located upstream the first gene (stdA; (López-Garrido & Casadesús, 2012)). Figure I. 6 Diagram of the std operon. std operon contains 6 genes (stdA-F), all of them transcribed by the promoter located upstream the first gene (PstdA). stdE stdFstdDstdCstdB yohM PstdA stdA SL1344_3009 std operon Introduction 35 The mechanisms that prevent std expression outside the animal environment remain unknown, but some of the bacterial functions involved in this regulation have been identified. Lack of RosE, an homolog to the E. coli ArgR repressor, allows std expression under laboratory conditions (Chessa et al., 2008). Dam methylation also prevents expression of the std operon, and Dam– mutants were found among Std-expressing isolates induced by transposon mutagenesis (Chessa et al., 2008). It was previously shown that the GATC- binding protein SeqA is a repressor of the std operon, and that the poorly known HdfR protein (a LysR-like protein) is required for std expression in S. enterica Dam– mutants (Jakomin et al., 2008). The possibility that HdfR might activate std transcription in a Damdependent manner is supported by the existence of three GATC sites upstream the std promoter, and unpublished results from our group suggest that HdfR binds the std promoter in a Dam methylation-dependent manner. Flow cytometry experiments in Dam- and SeqA- mutants show the existence of two subpopulations of Std cells: Std+ and Std– (Jakomin et al., 2008). The authors suggest the possibility of bistable or phase-variable std expression in the animal environment, and speculate that the competition between HdfR and SeqA or RosE might be the responsible for subpopulation formation. As in other fimbrial operons, structural and regulatory fimbrial proteins are encoded in the same operon. Recently, it was described that the Dam-dependent regulation of SPI-1 requires the last two genes of the std operon: stdE and stdF. Both genes repress SPI-1 expression through HilD at the postranscriptional level (López-Garrido & Casadesús, 2012). Lowered expression levels of the SPI-1 activator HilD would explain the reduced capacity of Dam– mutants to invade epithelial cells (García-Del Portillo et al., 1999). HdfR HdfR is a poorly known protein that belongs to the widely distributed LysR-type family of transcriptional regulators (LTTRs). LTTRs are the largest group of prokaryotic DNA-binding proteins (Maddocks & Oyston, 2008), and act as homodimers or homotetramers (Zaim & Kierzek, 2003). LTTRs control the transcription of multiple operons involved in disparate functions such as amino acid metabolism, oxidative stress, nitrogen fixation, degradation of aromatic compounds, and bacterial virulence (Schell, 1993). Based on the secondary structure, a helix-turn-helix (HTH) motif has been proposed to conform the DNA-binding domain (Kullik et al., 1995). This type of transcriptional regulators often needs a small ligand acting as a coinducer. The binding site of the inducer has been mapped to the C-terminal part (Lahiri et al., 2009). Many LTTRs activate the expression of target genes but repress their own expression, frequently by the use of divergent promoters (Zaim & Kierzek, 2003). In E. coli, HdfR was shown to negatively control flhDC expression through direct binding to its promoter, while hdfR expression was negatively regulated by H-NS (Ko & Park, 2000). In Salmonella few information is available aside from their role as an activator of std transcription in Dam– and SeqA– mutants (Jakomin et al., 2008). RcsCDB phosphorelay system The Rcs phosphorelay system appears to be conserved in the family Enterobacteriaceae (Pescaretti et al., 2009) and it is implicated in gene expresssion of a huge variety of genes, such as those controlling the biosynthesis of colanic acid (Stout & Gottesman, 1990), regulation of flagellum synthesis (Francez-Charlot et al., 2003), motility (Cano et al., 2002) or O-antigen chain length determination (Delgado et al., 2006), among others. Rcs regulon is activated by cell interaction with surfaces (conditions that lead to biofilm formation) and plays a role in later stages of biofilm formation (Danese et al., 2000; Majdalani & Gottesman, 2005). Perturbation in the cell surface induces the Rcs system. Mutations in rfa genes (synthesis of lipopolysaccharide) or mdo genes (membrane derived oligosaccharides or periplasmic glucans) implicate induction of Rcs system (Ebel et al., 1997; Majdalani & Gottesman, 2005). In Salmonella typhimurium, mutations in tolB (periplasmic component of Tol system) were found to increase Rcs regulon activity (Mouslim & Groisman, 2003). The Rcs system is composed by three proteins: RcsC, RcsD, and RcsB. Environmental signals are transmitted to RcsC, the sensor kinase, which in turn transfers a phosphate to a conserve aspartate on RcsB, the cognate response regulator. RcsD is the intermediary in the phosphoryl transfer (Majdalani & Gottesman, 2005). The main rol of RcsB is as a positive transcriptional regulator of a wide range of genes (capsule synthesis, membrane proteins), but Introduction 37 it has also been reported to behave as a negative regulator (flagellar synthesis) (Francez- Charlot et al., 2003). RcsB may act alone or in combination with other regulators, such as RcsA. RcsA is an unstable regulatory protein that acts with RcsB as an auxiliary regulator in some processes such as the capsule (cps loci) synthesis in E. coli, where the overexpression of rcsB is sufficient to increase cps levels in the absence of RcsA (Brill et al., 1988). On the other hand, RcsB is essential, independendly of RcsA levels (Majdalani & Gottesman, 2005). Dam methylation Base methylation at specific DNA sequences by DNA methyltransferases is a DNA modification common in all kingdoms of life. In α-Proteobacteria and γ-Proteobacteria, postreplicative N6 methylation in adenosine moieties lowers the thermodynamic stability of DNA (Engel & Hippel, 1978) and alters DNA curvature (Diekmann, 1987), influencing the interaction between DNA-binding proteins and their cognate DNA sequences (Wion & Casadesús, 2006). The methylation state of such sites can be used as a signal, thereby permitting spatial or temporal control of the interaction between DNA binding proteins and DNA (Messer & Noyer-Weidner, 1988). In bacteria, two types of DNA methyltransferases perform base modifications: DNA methyltransferases associated with restriction-modification systems (Bickle & Krüger, 1993; Loenen et al., 2014) and solitary DNA methyltransferases (Wion & Casadesús, 2006). Dam methylase of γ-proteobacteria is a solitary methyltransferase. Dam methylation is found in the orders Enterobacteriales, Vibrionales, Aeromonadales, Pasteurellales, and Alteromonadales (Løbner-Olesen et al., 2005). In E. coli and Salmonella, dam mutations produce pleiotropic defects but viability is not (or very little) impaired (Torreblanca & Casadesús, 1996). However, in Vibrio cholerae and in some strains of Yersinia Dam methylation is essential (Julio et al., 2001; Marinus, 1996). The Dam methylase transfers a methyl group from S-adenosyl-methionine to the N6 amino group of the adenosine moiety of 5’-GATC-3’ sites (Marinus, 1996). Methylation occurs shortly after DNA replication, and methylates transiently hemimethylated GATC sites. Nevertheless, the Dam enzyme can methylate both hemimethylated and nonmethylated GATC sites with similar efficiency (Marinus, 1996). Flanking sequences of the GATC sites influence the DNA binding ability of the Dam methylase and/or the efficiency of methyl group transfer (Peterson & Reich, 2006). The Dam protein shares significant identity with DpnII, MboI and other DNA methyltransferases that are part of restriction-modification systems (Low et al., 2001; Løbner-Olesen et al., 2005). This relatedness suggests that Dam has evolved from an ancestral restriction-modification system. An essential difference, however, is that the Dam methylase is highly processive, being able to achieve several methylation reactions without dissociating from the DNA molecule, while restriction-modification methylases are distributive (Urig et al., 2002). N6-methyl-adenine is used as a signal for genome defense, DNA replication and repair, nucleoid segregation, regulation of gene expression, control of transposition, and hostpathogen interactions (Low et al., 2001; Løbner-Olesen et al., 2005; Marinus, 1996) (Figure I. 5). Figure I. 7 Roles of N6-methyl-adenine in enteric bacteria. Methylation-sensitive DNA-binding proteins involved in each process are indicated. Adapted from (Wion&&&Casadesús,&2006). Dam methylation plays an important role in the initiation of chromosome replication in E. coli. Chromosome replication starts when DnaA, the initiator protein, binds the oriC Introduction 39 replication origin, and separates the two strands of the double helix. DnaA binding is only possible if the oriC GATCs are methylated; a hemimethylated origin is inactive (Boye et al., 2000). The oriC region contains 11 GATC sites in 254 bp, a density ten fold higher than the average expected in the E. coli chromosome (Zyskind & Smith, 1992). The GATC sites of the oriC remain hemimethylated for up to one-third of the cell cycle after DNA replication (Campbell & Kleckner, 1990). The protein SeqA avoids further rounds of replication by binding to hemimethylated GATC sites (Taghbalout et al., 2000). SeqA also binds to newly synthesized hemimethylated GATC sites along the chromosome, organizing the daughter chromosomes into nucleoid domains (Han et al., 2003). During DNA replication, mismatched base pairs errors are common. Dam methylation has an important role in mismatch repair by permitting the identification of the template DNA strand (Hsieh, 2001). The new strand is temporarily nonmethylated. MutS protein recognizes the mismatched base pair and recruits and assembles the MutHLS complex. This assembly activates the endonucleasa activity of MutH, which cleaves the nonmethylated DNA strand at the nearest hemimethylated GATC site, ensuring that the parental template strand is not cut (Friedhoff et al., 2003). As it was mentioned before, in E. coli and Salmonella, dam mutations produce pleiotropic defects such as lower colonization capacity, reduced motility, envelope instability, ectopic expression of fimbriae, sensitivity to bile salts, lower expression of virulence genes, and altered LPS O-antigen chain length (Cota et al., 2015; López-Garrido & Casadesús, 2010; Marinus & Casadesús, 2009). These pleiotropic virulence-related defects may explain the extreme attenuation of Dam– mutants upon oral inoculation in the mouse model (Badie et al., 2007; García-Del Portillo et al., 1999). Gene regulation by Dam methylation can be divided into two main categories (Low & Casadesús, 2008) : I. Clock-like controls, in which the methylation state of the DNA is used as a signal to couple gene expression to a specific stage of the cell cycle. Hemimethylation may activate transcription (conjugal transfer gene, traJ (Camacho & Casadesús, 2005)) or repress it (chromosome replication gene, dnaA (Kücherer et al., 1986). II. Switch-like controls, in which differential methylation patterns of GATC sites, typically found at or near the promoters of phase variation systems, modulate gene expression (van der Woude, 2011). Binding of proteins to DNA may prevent Dam methylation of certain GATCs, generating hemimethylated GATCs, and nonmethylated GATCs if the block persists during two consecutive DNA replication rounds (Casadesús & Low, 2006). DNA sequences surrounding specific GATCs sites may also decrease the processitivity of Dam methylation (Peterson & Reich, 2006). Phase variation systems regulated by Dam methylation As mentioned above, some envelope structures such as fimbriae, flagella or the LPS O- antigen show phase variation and are under epigenetic control (Cota et al., 2015; Cummings et al., 2006; Kingsley et al., 2002; Nicholson & Low, 2000). In phase variation systems, gene expression alternates between active (ON phase) and inactive (OFF phase) states, and often depends on differential DNA methylation patterns (Switch-like controls). Binding of certain transcriptional regulators hinders Dam methylation and creates heritable DNA methylation patterns (Figure I. 8). pap The pap operon encodes pyelonephritis-associated pili that mediate adhesion of uropathogenic E. coli to the urinary mucosa (Hernday et al., 2004). Synthesis of Pap pili is subjected to phase variation, and Dam methylation controls the switching between ON and OFF states (Hernday et al., 2002). The regulatory region of the pap operon contains six binding sites for the leucine-responsive regulatory protein, Lrp. Sites 5 and 2 contain GATC motifs known as GATCdist and GATCprox, respectively. In the OFF state, Lrp binds cooperatively and with high affinity to sites 1-3 avoiding RNA polymerase binding (Casadesús & Low, 2006; Hernday et al., 2002; 2004). Lrp binding at sites 1-3 reduces the affinity of Lrp for sites 4-6, and preserves the nonmethylated state of GATCprox while GATCdist remains methylated. The high affinity of Lrp for nonmethylated GATCprox and its incapability to bind a methylated GATCdist generates a feedback loop that propagates the OFF state (Hernday et al., 2002; 2004). Switching to the ON state needs translocation of Lrp to sites 4-6. Translocation involves the auxiliary protein Introduction 41 PapI. The PapI/Lrp complex has higher affinity for GATC sites 4-6 than for 1-3 GATCs (Hernday et al., 2003). Binding of Lrp to sites 4-6 hinders methylation of GATCdist and permits methylation of GATCprox, favoring the propagation of the ON state (Hernday et al., 2004). agn43 The agn43 gene of E. coli encodes Agn43, an autotransporter protein whose expression is subjected to phase variation under control of Dam methylation and OxyR (Henderson & Owen, 1999; Henderson et al., 1997). Three GATC sites are contained within the binding site of OxyR at agn43 regulatory region (Wallecha et al., 2002). OxyR binding to nonmethylated agn43 GATCs represses transcription and prevents GATC methylation. On the other hand, methylation of GATC sites avoids OxyR binding (Haagmans & van der Woude, 2000; Henderson et al., 1997; Waldron et al., 2002). agn43 expression thus depends on the competition between OxyR binding and Dam methylation, and switching may occur upon DNA replication (Waldron et al., 2002; Wallecha et al., 2002). opvAB The Salmonella enterica opvAB operon shows phase variation and creates bacterial lineages with standard (OpvABOFF) and shorter (OpvABON) O-antigen chains in the lipopolysaccharide. Transcription of OpvAB is controlled by the LysR-type factor OxyR and by Dam methylation. The opvAB regulatory region contains four sites for OxyR binding (OBSA-D), and four GATC motifs (GATC1-4) which are targets for Dam methylation. OpvABOFF and OpvABON cell lineages present opposite DNA methylation patterns in the opvAB regulatory region: (i) in the OpvABOFF state, GATC1 and GATC3 are non-methylated, whereas GATC2 and GATC4 are methylated; (ii) in the OpvABON state, GATC2 and GATC4 are non-methylated, whereas GATC1 and GATC3 are methylated (Cota et al., 2015). The OBSA and OBSC sites of opvAB are identical to the consensus sequence for OxyR binding while OBSB and OBSD are not. This difference may explain the higher stability of the OpvABOFF lineage, resulting in a ∼600-fold difference in the ON→OFF and OFF→ON transition rates. The predominant OFF state involves binding of OxyR to the OBSA and OBSC sites, which protects GATC1 and GATC3 from methylation. In this configuration, MATERIALS AND METHODS Materials and Methods 51 Bacterial strains and strain construction Salmonella enterica and Escherichia coli strains used in this Thesis are listed in Table M1. Salmonella enterica strains belong to serovar Typhimurium and derived from the mousevirulent strain SL1344 or ATCC14028. For simplicity, Salmonella enterica serovar Typhimurium is often abbreviated as S. enterica. Table M 1. Strains of Salmonella enterica and E.coli used in this study. Strain Name Genotype Reference S. enterica SL1344 Wild type (Hoiseth & Stocker, 1981) SV5367 SL1344 ∆dam231 (Balbontín et al., 2006) SV8479 SL1344 stdA::gfp This study SV8480 SL1344 ∆dam231 stdA::gfp This study SV8481 SL1344 ∆hdfR stdA::gfp This study SV9311 SL1344 ∆dam231 ∆hdfR stdA::gfp This study SV8528 14028 rcsC11 stdA::gfp This study SV5182 14028 stdA::lacZ (Jakomin et al., 2008) SV5183 14028 ∆dam231 stdA::lacZ (Jakomin et al., 2008) SV9312 14028 ∆dam231 rcsB56N stdA::lacZ This study SV9313 14028 ∆dam231 rcsC11 stdA::lacZ This study SV9314 14028 ∆dam231 ∆tolB stdA::lacZ This study SV9315 14028 ∆dam231 p-rcsB stdA::lacZ This study SV5264 14028 ∆dam231 (López-Garrido & Casadesús, 2010) SV5815 14028 ∆dam231 rcsB56N This study SV9316 14028 ∆dam231 rcsC11 This study SV9317 14028 ∆dam231 ∆tolB This study SV9318 14028 ∆dam231 p-rcsB This study SV8487 SL1344 ∆dam231 hdfR-3xFLAG This study SV7889 SL1344 hdfR::lacZ This study SV9319 SL1344 ∆dam231 hdfR::lacZ This study SV8188 SL1344 stdA::lacZ This study SV8449 SL1344 PLtetOhdfR This study SV8477 SL1344 PLtetOhdfR stdA::lacZ This study SV9292 SL1344 PLtetOhdfR stdA::gfp This study SV9123 SL1344 pIC552 This study SV9124 SL1344 /pIZ2320 This study SV9125 SL1344 ∆hdfR /pIZ2320 This study SV8192 SL1344 stdA::lacZ /pBR328 This study SV8194 SL1344 stdA::lacZ /pIZ2318 This study SV9320 SL1344 stdA::lacZ /pIZ2319 This study SV8189 SL1344 ∆hdfR stdA::lacZ This study SV8193 SL1344 ∆hdfR stdA::lacZ /pBR328 This study SV8195 SL1344 ∆hdfR stdA::lacZ /pIZ2318 This study SV9321 SL1344 ∆hdfR stdA::lacZ /pIZ2319 This study SV9322 SL1344 PLtetOstdEF stdA::gfp This study SV9323 SL1344 ∆stdEF stdA::gfp This study SV9279 SL1344 ∆dam231 ∆stdEF stdA::gfp This study SV9001 SL1344 ∆dam231 PLtetOstdEF stdA::gfp This study SV8507 SL1344 PLtetOstdEF stdA::lacZ This study SV9324 SL1344 ∆dam231 stdE-3xFLAG This study SV9325 SL1344 ∆dam231 stdF-3xFLAG This study SV8526 SL1344 ptp166 stdA::gfp This study SV9205 SL1344 stdE::gfp This study SV9207 SL1344 ∆dam231 stdE::gfp This study SV9206 SL1344 ∆hdfR stdE::gfp This study SV9208 SL1344 ∆pstdA-stdC stdE::gfp This study SV6503 14028 PLtetOstdEF (Cmr) (López-Garrido & Casadesús, 2012) SV6634 14028 PLtetOΔstdEF (Cmr) (López-Garrido & Casadesús, 2012) SV8141 SL1344 PLtetOstdEF (Cmr) This study Materials and Methods 53 SV8142 SL1344 PLtetO∆stdEF (Cmr) This study SV7553 SL1344 PLtetOstdEF (KmR) This study SV7552 SL1344 PLtetO∆stdEF (KmR) This study SV7554 SL1344 PLtetOstdEF spvA This study SV7555 SL1344 PLtetOΔstdEF spvA This study SV7556 SL1344 spvA::tn5dKm This study SV4938 SL1344 trg::mudQ pSLT- (García-Quintanilla & Casadesús, 2011) SV7551 SL1344 traB::lacZ This study SV7550 SL1344 PLtetOstdEF traB::lacZ This study SV7549 SL1344 PLtetOΔstdEF traB::lacZ This study SV8152 SL1344 ∆ygiD This study SV9326 SL1344 PLtetOstdEF ∆hdfR This study SV9327 SL1344 PLtetO∆stdEF ∆hdfR This study SV9288 SL1344 flhC::lacZ This study SV9289 SL1344 PLtetOstdEF flhC::lacZ This study SV9290 SL1344 PLtetO∆stdEF flhC::lacZ This study SV9328 SL1344 ∆hdfR flhC::lacZ This study SV9329 SL1344 PLtetOstdEF ∆hdfR flhC::lacZ This study SV9320 SL1344 PLtetO∆stdEF ∆hdfR flhC::lacZ This study SV9331 SL1344 PLtetOstdEF ∆hdfR spvA This study SV9332 SL1344 PLtetOΔstdEF ∆hdfR spvA This study SV9333 SL1344 ∆hdfR spvA This study SV8185 SL1344 sipB::lacZ This study SV8186 SL1344 PLtetOstdEF sipB::lacZ This study SV8187 SL1344 PLtetO∆stdEF sipB::lacZ This study SV8109 SL1344 ∆flhC sipB::lacZ This study SV8110 SL1344 PLtetOstdEF ∆flhC sipB::lacZ This study SV8111 SL1344 PLtetO∆stdEF ∆flhC sipB::lacZ This study SV8103 SL1344 fliC::lacZ This study SV8104 SL1344 PLtetOstdEF fliC::lacZ This study SV8105 SL1344 PLtetO∆stdEF fliC::lacZ This study SV8106 SL1344 ∆flhC fliC::lacZ This study SV8107 SL1344 PLtetOstdEF ∆flhC fliC::lacZ This study SV8108 SL1344 PLtetO∆stdEF ∆flhC fliC::lacZ This study SV8023 SL1344 ∆flhC traB::lacZ This study SV8024 SL1344 PLtetOstdEF ∆flhC traB::lacZ This study SV8025 SL1344 PLtetO∆stdEF ∆flhC traB::lacZ This study SV9287 SL1344 PLtetOstdEF-3xFLAG This study SV9109 SL1344 PLtetOstdEF ∆flhC This study SV9110 SL1344 PLtetOΔstdEF ∆flhC This study SV9309 SL1344 PLtetOstdEF PLtetOhilD This study SV9310 SL1344 PLtetOΔstdEF PLtetOhilD This study SV9113 SL1344 PLtetOstdEF invH::lacZ This study SV9114 SL1344 PLtetOΔstdEF invH::lacZ This study SV9119 SL1344 PLtetOstdEF PLtetOhilD invH::lacZ This study SV9120 SL1344 PLtetOΔstdEF PLtetOhilD invH::lacZ This study SV9273 SL1344 ∆dam231 ∆stdEF This study SV7890 SL1344 PLtetOstdEF hdfR::lacZ This study SV7891 SL1344 PLtetO∆stdEF hdfR::lacZ This study TR5878 galE496 r(LT2)– m(LT2)– r(S)+ ilv-542 metA22 trpB2 Fels2– fliA66 strA120 xyl-404 metE551 hspL56 hspS29 J.R. Roth E. coli DH5α supE44 ∆lacU169 (Φ80 lacZ∆M15) hsdR17 recA1 endA1 gyrA96 thi-1 reA1 (Hanahan, 1983) BL-21 F- dcm ompT hsdS (rB- mB-) gal (malB+) K-12 (λS) Stratagene BL-21 pET28a-stdE This study Materials and Methods 55 Targeted gene disruption was achieved by Datsenko and Wanner method (Datsenko & Wanner, 2000). This method is based in the λ Red recombination system. The strategy consists in replacing the chromosomal sequence by an antibiotic resistance marker (contained in pKD3, pKD4 or pKD13) that is generated by polymerase chain reaction (PCR) using oligonucleotides that harbor 40 nucleotides of homology with the sequence to be replaced. λ Red recombination gene expression is carried out under an inducible promoter inside a thermosensitive low copy number plasmid (pKD46). Cultures harbored pKD46 plasmid (this plasmid expresses the λ Red system under the araB promoter which is inducible by arabinose) grown in LB with ampicillin at 30ºC were diluted 1:100 into LB with ampicillin and arabinose 1 mM and incubated in a shaker at 30ºC until O.D.600 ~ 0.8. The competent cells were prepared and electroporation was done as further described. After selection, antibiotic resistance cassettes introduced during strain construction can be excised by recombination with plasmid pCP20 (Datsenko & Wanner, 2000). pCP20 plasmid can be transduce with P22. This transduction is incubated at 30ºC for 1h and spread in LB with ampicillin. To eliminate the plasmid, green plates were prepared without antibiotic and incubated at 37ºC. To confirm the excision of the marker, the strains were streaked in plates of LB ampicillin and plates of LB with chloramphenicol or kanamycin. The excision of the antibiotic marker was checked by colony PCR with external oligonucleotides. The oligonucleotides used for gene disruption (named as UP and DO) and allele verification (named as E1 and E2) by the polymerase chain reaction (PCR), are listed in Table M2. For construction of lac fusions in the Salmonella chromosome, FRT sites generated by excision of KmR cassettes with pCP20 (Datsenko & Wanner, 2000) were used to integrate either pCE37 or pCE40 (Ellermeier et al., 2002). For strain construction operations involving chromosomal markers we used transduction with P22 HT 105/1 int201 ((Schmieger, 1972) and G. Roberts, unpublished). The recipient strain was transduced using a P22 lysate from a strain with the desirable genetic marker. All the markers used in this thesis were selected directly by spreading the transduction mixture on selective media.!Transduction protocol is described below. Table M 2. Oligonucleotides used in thi study. Oligonucleotide name sequence (5’!3’) stdA129GFP UP tttcactggtaccatcaccaactcaccctgtgatatcgcataagaaggagatatacatatg stdAGFP DO ccgtggacggcttctccctgtcgttatttaccgcgtgaaattatcacttattcaggcgta stdA E1 ggaaagttcaggtgcttcg stdAE2 gctttcggtgttgtcgtcc PLtetOhdfR UP atatcgattagtcgtcgtaaagctttccgccatcctgcacaggcttacccgtcttactgtc PLtetOhdfR DO ggaaagtttttaacaattccgtatccactgtgctctccacaaaacctctccataactggg hdfR E1 ggagagcacagtggatacgg hdfR E2 gattatctgatcaggtaatc promotorhdfRBglII FOR agctagatctccgtgacgagagaatccacg promotorhdfRXhoI REV agctctcgaggatacgaaagctcaccgcgg PLtetOstdEF pStd UP gttttctgctgcaatacccgttactgttacctataactaaaggcttacccgtcttactgtc PLtetOstdEF pStd DO tcagggcacataaaacctctccataactgggtaaatgatgtgctcagtatctctatcactgatag stdE21GFP UP ccagttatggagaggttttatgtgccctgataatacacactaagaaggagatatacatatg stdEGFP DO ttaccgacccggcgttttgataccagcggcggtccggcttttatcacttattcaggcgta stdE E1 tgctgcaatacccgttactg stdE E2 caggctgcctgtatgcg PLtetO sense ttggaacctcttacgtgcc mutpStdA-stdC UP ccttttacttgcattatatccagttttttatgctaccagtgtgtaggctggagctgcttc mutpStdA-stdC DO gatgtcgcccctgcgtccttaaacgcgcagttcagggcatcatatgaatatcctccttag Km PLtetOUP ttaatgtcatgataataatggtttcttagacgtcgatatcgtgtaggctggagctgcttc Km PLtetODO cttggattctcaccaataaaaaacgcccggcggcaaccgacatatgaatatcctccttag NdeIstdE-FOR ttttcatatgtgccctgataatacaca EcoRIstdE-REV ttttgaattcttatgttccgtcatcctcatcc std-PLtetOUP attcacccgcttataattac std southern DO cagctaatgcctaatcagttg Materials and Methods 57 EMSA-std300 FOR attcacccgcttataattac EMSA-std735 FOR cgcattaatatcccccagcc EMSA-std REV attacgcatagataatatgtc 8032 EMSA-Fwd Pdps gcgctattacttcgtc 8033 EMSA-Rev Pdps cgggatccctcatatcctcttgatgtttgtgt pBR328-Fw actgtccgaccgctttgg pBR328-Rv gccagcaaccgcacctg ygiD UP ggttatctgagttcttcctctgtgaagaaagcgtatggtggtgtaggctggagctgcttc ygiD DO cgccatcgggcaatcattcagcgccttatccggcctaccccatatgaatatcctccttag ygiD E1 tatcaatttgccgtccgaacc ygiD E2 tcgaaggatagattctcctg RT-hilD FOR agtttgctttcggagcggta RT-hilD REV agcaccaacatcccaggttc RT-promotorstd FOR cattaaaaagtatttctttgatg RT-promotorstd REV gaaaattcttattcaaattaaaac RT-flgE DIR gacggtacgacaacgaacac RT-flgE REV aaaaccgttctggctaatcg RT-hilA DIR gaatcttttcatggctggtca RT-hilA REV gggtccaattttaaacactcgt RT-motA DIR taggggcgttcattgtcg RT-motA REV acgggatagctttcatcgtg RT-sipB DIR gtaatggtggccgatgaaat RT-sipB REV cgcctgctgaataaacgac RT-traA DIR gtacggtcaaggcgacattt RT-traA REV ccgcgagaataacccactt RT-trg DIR cggggtcgtacaaacgat RT-trg REV gcggtaatttccgagattttt For ligation of DNA fragments with the appropriated vector (properly digested), 1 U of T4 DNA ligase (1 U/µl, Roche Diagnosstics) was used in the buffer supplied by the manufacturer. Routinely, the mixture was incubated at 16ºC overnight. Electrophoresis in an 0.8% agarose (Low Electro Endosmosis agarose-Pronadisa, Conda, España) gel made up in TAE 1x buffer (40 mM Tris-acetate and 10 mM EDTA pH7.7) was used to test the quality of DNA extraction, to determine DNA fragments after plasmid restriction, to estimate the efficiency of endonuclease restriction, etc. The loading buffer used was 10X Loading buffer supplied by Takara. The 1 Kb ladder (GIBCO BRL, Life Technologies, New York, USA) was used as molecular weight marker. Ethidium bromide (0.5 µg/ml final concentration) was added to the gels to make bands visible. Gels were illuminated with UV and pictures were taken with Gel DocTM Imager- BioRad. Bacterial transformation High efficiency E. coli transformation Competent cells were prepared using a variation of the Inoue method (Inoue et al., 1990), which guarantees high transformation efficiency (5x107 to 5x108 transformants per µg of plasmid DNA). An E. coli DH5α/BL-21 overnight culture was diluted 100-1000 times in 200 ml of SOB (20 g/l tryptone, 5 g/l yeast extract, 0.5 g/l NaCl, 0.19 g/l KCl, adjust with NaOH to pH7 and 5 ml MgCl2 2 M after autoclaving), and incubated at 22ºC and 200 rpm until OD600 ∼0.5 was reached. The culture was chilled quickly on ice and kept on ice for 10 minutes. Cells were harvested by centrifugation at 2,500 g and 4ºC for 10 min. The pellet was resuspended in 20 ml of cold filtrated TB (10 mM PIPES, 15 mM CaCl2, 250 mM KCl, adjust to pH6.7 with KOH, 55 mM MnCl2 ), and 1.5 ml of dimethyl sulfoxide (DMSO) was added. After 10 min incubation on ice, aliquots of 0.2-0.5 ml were prepared, freezed in liquid nitrogen, and stored at -80º C. For transformation, an aliquot of competent cells was slowly thawed on ice and was mixed with the plasmid. The mixture was incubated on ice for 30 minutes, subjected to heat shock (42oC, 45 s), and cooled on ice for 1 min. One ml of LB was then added. The mixture was incubated at 37oC for 1 h; finally, the cells were concentrated in 100 µl and spread on selective media. Materials and Methods 65 Salmonella electroporation An overnight culture was diluted 1/100 in LB and, depending on the strain, was grown at 37oC or 30oC (strains with a thermolabile plasmid like pKD46) until an OD600 ∼0.8 was reached. The cultured was chilled on ice and kept on ice for 5 minutes. 20 ml cells were harvested by centrifugation at 4000 rpm for 5 min at 4oC. The supernatant was discarded and the bacterial pellet was softly re-suspended in 1 ml of cold ddH2O. Once cells were resuspended, 19 ml of cold water were added, and cells were washed again. Finally, cells were harvested and re-suspended in 250 µl of water. Electroporation was performed by mixing 1 µl of plasmid DNA or 10 µl of PCR product with 50 µl of competent cells. The mixture was transferred to a cuvette that had 2 mm gap between the plates. The cuvette was subjected to an electric discharge in the electroporator (2.5 KV, 200 Ω and 25 µF). The electroporator employed was a BTX Electrocell Manipulator 600 (Harvard Apparatus, Holliston, Massachusetts, USA). After the discharge, 1 ml of LB was added to the cells, and the mixture was transferred to a 10 ml tube, which was incubated at 37o C with shaking for 1 h. Finally, cells were concentrated in 100 µl and spread on selective media. pBR328-based plasmid library genetic screen pBR328-based multicopy plasmid library of the Salmonella enterica genome was used to search for factors that might activate expression of the std operon in a wild type (Dam+) background stdA::lacZ strain (SV5182). This library was previously constructed and described (López-Garrido & Casadesús, 2012). Briefly, Salmonella enterica Serovar Typhimurium SL1344 was partially digested with Sau3A. DNA fragments ranging between 7-11 kb long were ligated to the pBR328 vector, previously digested with BamHI and dephosphorylated. Salmonella strain TR5878 was transformed with the ligation products, and ampicillin–resistant clones were selected on LB with Ap plates. Pools of approximately 1000 independent transformants were collected and lysed with phage P22 HT 105/1 int201. As a quality control, the ability of the library pools to complement null mutations in araA (required for growth with L-arabinose as the sole carbon source) or xylA (required for growth with D-xylose as the sole carbon source) were tested. Lysates that permitted successful complementation were stored and used for plasmid delivery to recipient strains in subsequent genetic screens. The genetic screen was done by mixing 100 µl of a saturated stdA::lacZ culture with 10 µl of each P22 lysate of the pBR328 plasmid library pools. After 45 minutes, the transductions were plated in LB Cm and X-gal agar plates and incubated at 37ºC overnight. 10 transductions were made for each of the 9 different pools (90 transductions total). The following day, the stdA::lacZ background colonies with increased X-gal activity were culture in LB Cm liquid medium. Minipreps were done to recover the pBR328 plasmid derivatives. Positive plasmids were re-transformed into SV5182 to corroborate the positive phenotypes. β-galactosidase activity of the selected candidates was tested and the plasmids were sent for Sanger DNA sequencing of the inserts using pBR328 specific oligonucleotides that flanked the region of the insertion. pIZ2318 and pIZ2319 obtained from the genetic screen are listed in Table M4. β-galactosidase assays Levels of β-galactosidase activity were assayed using the CHCl3-sodium dodecyl sulphate permeabilization procedure (Miller, 1972). β-galactosidase activity data are the averages and standard deviations from ≥3 independent experiments. Motility assays Motility assays were carried out in motility agar plates, containing 10 g/l tryptone (Difco), 5 g/l NaCl, and 0.25% Bacto-agar (Cano et al., 2002). A sterile stick was soaked in saturated bacterial cultures grown in LB, and used to inoculate motility agar plates. Bacterial motility halos were compared after growth at 37ºC for 6 hours. Phenotypic assays for biofilm formation Salmonella strains (SL PLtetOstdEF, SL PLtetOΔstdEF and SL ΔygiD) were tested for their ability to produce biofilm in LB medium (Latasa et al., 2012). The culture was grown at 22ºC Materials and Methods 67 for 7-10 days in static. For better visualization biofilm was stained with a 0.1% solution of crystal violet. Matings Cultures of the donor and the recipient were grown overnight in LB broth. Cells were harvested by centrifugation and washed with LB. 1 ml aliquots of both strains, were sucked onto a membrane filter 0.45 mm pore size. The donor/recipient ratio was 1:1. The filters were placed on LB plates and incubated during 4h at 37ºC in GasPak microaerophilic jars (Camacho & Casadesús, 2002) and microaerophilic conditions were obtained by a GENbox Anaer bag, supplied by BioMériex (Marcy l’Etoile, France). An anaerobic indicator was used to monitor microaerophilic conditions. Filters were place in 10 ml tubes with 1 ml of fresh LB and vortex for recovering the cells. Serial dilutions were done before plating. Conjugation frequencies were calculated per donor cell, as previously described (Camacho & Casadesús, 2002; García-Quintanilla et al., 2008). Subcellular fractionation Subcellular fractionation was performed as previously described (Pucciarelli et al., 2002), with some modifications. Briefly, bacteria were grown in LB medium at 37ºC and spun down by centrifugation at 15,000 g for 5 min at 4ºC, then resuspended twice in cold phosphatebuffered saline (PBS pH 7.4, 0.137 M NaCl, 2,7 M KCl, 4.3 mM Na2HPO4.7H20, 1.4 mM KH2PO4). Unbroken cells were further removed by low-speed centrifugation (5,000 g, 5 min, 4ºC). The supernatant was centrifuged at high speed (100,000 g, 30 min, 4ºC) and the new supernatant was recovered as the cytosol fraction. The pellet containing envelope material was suspended in PBS with 0.4% Triton X-100 and incubated for 2 h at 4ºC. The sample was centrifuged again (100,000 g, 30 min, 4ºC) and divided into the supernatant containing mostly inner membrane proteins and the insoluble fraction corresponding to the outer membrane fraction. An appropriate volume of Laemmli buffer (SB4x: 0.000125% bromophenol blue, 200 mM Tris-HCl pH 6.8, 20% β-mercaptoethanol, 40% glycerol and 8% SDS) was added to each fraction. After heating (100ºC, 5 min) and clearing by centrifugation (15,000 g, 5 min, room temperature), the samples were analysed for protein content by Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) described by Laemmli (Laemmli, 1970). Protein extracts and Western blotting analysis Total protein extracts were prepared from bacterial cultures grown at 37ºC in LB medium until stationary phase (O.D.600 ~2). A volume containing ~2.5 x 108 cells were collected by centrifugation and suspended in 50 µl of Laemmli sample buffer (SB4x: 0.000125% bromophenol blue, 200 mM Tris-HCl pH 6.8, 20% β-mercaptoethanol, 40% glycerol and 8% SDS). Proteins were resolved by SDS-PAGE (Laemmli, 1970) using 12% gels. Conditions for protein transfer have been described elsewhere (Jakomin et al., 2008). Primary antibody was anti-Flag M2 monoclonal antibody (1:5,000, Sigma-Aldrich, St. Louis, MO). Goat antimouse horseradish peroxidase-conjugated antibody (1:5,000; Bio-Rad, Hercules, CA) was used as secondary antibody. Proteins recognized by the antibody were visualized by chemoluminescence using the luciferin–luminol reagents of Supersignal West Pico Chemiluminiscent Substrate (Thermo Scientific, Waltham, MA, USA) and developed in a LAS3000 mini system (Fujifilm, Tokyo, Japan). Purification of StdE protein and antibody generation A DNA fragment containing stdE was amplified using oligonucleotides StdEprotUP and StdEprotDO, and cloned into pET28a (Novagen) using the NdeI and EcoRI sites. The recombinant plasmid (pIZ1991) was verified by restriction analysis and DNA sequencing. For 6×His-StdE purification, plasmid pIZ1991 was transformed into E. coli BL-21. BL- 21/pIZ1991 was grown in LB broth containing Kanamycin, and expression of 6×His-StdE was induced with 1 mM isopropyl β-D-thiogalactopyranoside (IPTG). After 3 h of induction, cells were centrifuged and resuspended in 10 ml of lysis buffer (20 mM Tris, 300 mM NaCl, 10 mM imidazole) per g of pelleted cells, and were lysed by sonication. The suspension was centrifuged at 10,000 rpm for 20 min at 4º C and the supernatant containing the soluble fraction of 6×His-StdE was transferred to a HisTrap HP nickel affinity chromatography column (GE Healthcare, Wauwatosa, WI, USA). The column was washed 3 times with 4 ml of washing buffer (20 mM NaH2PO4·H2O, 0.5 mM NaCl, 30 mM imidazole). Protein elution was performed with 3 ml of elution buffer (20 mM NaH2PO4·H2O, 0.5 mM NaCl, 300 mM Materials and Methods 69 imidazole). Aliquots of each fractios were loaded in SDS-PAGE gel and stain with Coomassie Brilliann Blue R-250 (Sigma-Aldrich) solution (0.25% (w/v) Coomassie Brilliant Blue R-250, 10% (v/v) acetic acid and 10% (v/v) ethanol). Elution fractions enriched in 6×His-StdE were selected. Imidazole was removed by dialyzing in cellulose membranes with PBS 1X. Purified 6×His-StdE protein was sent to Biomedal S.L (Sevilla, Spain) for polyclonal antisera production in rabbits. Working dilution was made based on manufacture recommendations. RNA isolation To prepare cells for RNA extraction, 3 ml of fresh LB was inoculated with a 1:100 dilution from an overnight bacterial culture, and incubated with shaking at 200 rpm in an Infors Multritron shaker at 37ºC. A 2 ml aliquot from a stationary culture (O.D.600∼2) was centrifuged at 13000 rpm, 4ºC, during 5 min. The pellet was resuspended in 100 µl of a solution of lysozyme (3 mg/ml in water; Sigma Chemical Co.). Cell lysis was facilitated by a freeze-thaw cycle. After lysis, RNA was extracted using 1 ml of TRIsure reagent following manufacture’s instructions (Bioline, Taunton, Massachusetts, USA). Total RNA was resuspended in 30 µl of RNase-free water, and subsequently clean by extraction with acidic phenol, followed by a second extraction with chloroform:isoamilic alcohol (24:1). After extraction, RNA was precipitated with ethanol and 3 M sodium acetate, and the dried pellet was resuspended in RNase-free water. The quantity and quality of the RNA was determined using a ND-1000 spectrophotometer (NanoDrop Technologies). Quantitative reverse transcriptase PCR (qRT-PCR) For qRT-PCR, Salmonella RNA was extracted from stationary phase cultures (O.D.600∼2) as described above, and the concentration was determined using a ND-1000 spectrophotometer (NanoDrop Technologies). An aliquot of 1 µg of RNA was used for cDNA synthesis using QuantiTec® Reverse Transcription Kit (Quiagen) following manufacturer’s instructions. Quantitative RT-PCR reactions were performed in a Light Cycler 480 II apparatus (Roche). Each reaction was carried out in a total volume of 10 µl on a 480-well optical reaction plate (Roche) containing 5 µl SYBR mix, 0.2 µl DYE II (Takara), 4 µl cDNA (1/10 dilution) and two gene-specific primers at a final concentration of 0.2 mM each. Real-time cycling conditions were as follows: (i) 95ºC for 10 min and (ii) 40 cycles at 95ºC for 15s, 60ºC for 1min. A non-RT control was included for each primer set. Triplicates were run for each reaction, and the Ct value is averaged from them. Absence of primer dimers was corroborated by running a dissociation curve at the end of each experiment to determine the melting temperature of the amplicon. Melting curve analysis verified that each reaction contained a single PCR product. Gene-specific primers were designed with ProbeFinder software (http://www.universalprobelibrary.com) from Roche Applied Science and are listed in Table M2. For quantification, the efficiency of each primer pair was determined to be between 90%- 110%, following the instructions for efficiency determination described in the “Guide to Performing Relative Quantification of Gene Expression Using Real-Time Quantitative PCR” (Applied Biosystems). These efficiencies indicate that the amount of DNA is doubled in each PCR cycle, and allows for direct comparison between different genes. Relative RNA levels were determined using the ∆∆Ct method as described in the above mentioned guide. Briefly, each gene Ct value is normalized to the Ct value for the internal control (rfaH), which gives the ∆Ct value. This value is then related to a given gene in the reference strain (S. enterica, in this case) giving us the ∆∆Ct value. Since the amount of DNA doubles in each PCR cycle, the relative amount of input cDNA can be determined by using the formula 2-∆∆Ct. Each ∆∆Ct determination was performed at least in three different RNA samples (three biological replicates), and the results are representative example of such determinations. Analysis of hilD mRNA decay Use of quantitative RT-PCR to monitor mRNA decay has been previously described (Baker et al., 2007). An overnight LB culture of the strain under study was diluted 100 fold, and incubated at 37ºC with shaking until O.D.600∼2). Transcription initiation was stopped by adding rifampicin (500 mg/ml). Cultures were kept at 37ºC and aliquots were extracted at 1 min intervals. Each aliquot was immediately immersed in liquid N2 and kept frozen until RNA extraction. RNA was extracted using the standard protocol above described and amounts of hilD mRNA were determined with qPCR, using oligos RT-hilD FOR and RT- hilD REV as described above. Three independent qRT-PCR reactions were performed. Materials and Methods 71 Southern blot Genomic DNA was isolated as previously described from stationary cultures in LB (O.D.600 ~2). Forty µg of each DNA sample were digested with SspI (New England Biolabs, Ipswich, MA), purified and divided into four fractions, three of which were subsequently digested with DpnI, MboI or Sau3AI (New England Biolabs). After digestion the samples were run in a denaturing 8% TBE-polyacrilamide (19:1) 8 M urea gel. Electrophoresis was carried out in a Hoefer SE400 (Hoefer Scientific Instruments, San Francisco, California) subjected to an electric field of 35 mA for 60 minutes. DNA was transferred to an Amersham Hybond-N+ membrane (GE Healthcare, Wauwatosa, WI, USA) using a semidry Electroblotting system (Thermo Scientific, Waltham, MA, USA). The DNA in the membrane was then immobilized by UV crosslinking. A radioactive probe was prepared by PCR using dCTP [α-32P] (Perkin Elmer, Waltham, MA) and oligonucleotides stdPLtetOUP and std southern DO. After the PCR reaction, non-incorporated nucleotides were removed by treatment in a Sephadex G-25 column (Illustra MicroSpin G-25 columns, GE Healthcare, Wauwatosa, WI) following manufacturer’s instructions. Prior to hybridization the double-stranded DNA probe was denatured by heating at 95ºC for 3 min, followed by incubation on ice. Hybridization with the probe was performed overnight at 52ºC in hybridization buffer (0.5 M sodium phosphate pH 7.2, 10 mM EDTA, 7% SDS). Excess probe was removed with washing buffer (40 mM sodium phosphate pH 7.2, 1% SDS) at 48ºC (three washes, 30 min each). The membrane was developed using a FLA-5100 Scanner (Fujifilm, Tokyo, Japan). Microarray procedures and data analysis Salmonella enterica comparative transcriptomic analyses were performed in a strain that constitutively expressed stdEF (SL1344 PLtetOstdEF- SV8141) and in a control strain carrying an in-frame deletion of both genes (SL1344 PLtetO∆stdEF- SV8142). RNA from two biological replicates of each strain was isolated and used for the assay. Transcriptomic analyses were performed using Salmonella enterica serovar Typhimurium SL1344 4X72K array, custom prepared for Dr. Antonio Juarez (Paytubi et al., 2014). Hybridation and microarray scanning were performed at the Functional Genomics Core of the Institute for Research in Biomedicine, Baldiri Reixac, Barcelona, Spain (http://www.dnaarrays.org/). Normalization of the expression signals was done with RMA (Irizarry et al., 2003) using Partek Genomics suite6.5 (6.11.0207). Raw transcriptomic data were deposited at the Gene Expression Omnibus, G.E.O, database (http://www.ncbi.nlm.nih.gov/geo/) under accession number GSE45488. Raw transcriptomic data was analysed building the ratio between the two conditions of interest by subtracting. Those genes with a difference higher than 4-fold in expression were further study. Chromatin Immunoprecipitation followed by sequencing (ChIP-seq) and data analysis ChIP-seq assays were used to determine chromosome-wide DNA binding profiles of StdE and StdF in Salmonella enterica serovar Typhimurium SL1344. PLtetOstdEF-3xFLAG strain (SV9287) was used to perform ChIP-seq experiments. 20 ml of fresh LB was inoculated with a 1:100 dilution from an overnight bacterial culture, and incubated with shaking at 200 rpm in an Infors Multritron shaker at 37ºC. Cells collected at O.D.600 ∼2 were crosslinked with 1% formaldehyde (37% solution) at 37ºC for 25 min, following by quenching the unused formaldehyde with 450 mM glycine for 5 min. Crosslinked cells were harvested by centrifugation at 9000 rpm at 4ºC for 10 min and washed with 10 mL of TBS pH7.6 (2.42 g/l Trizma base, 8 g/l NaCl). The washed cells were re-suspended in 1 mL of lysis buffer (10 mM Tris-HCl pH 8, 20% sucrose, 50 mM NaCl, 10 mM EDTA) and after an additional centrifugation step, the cells were re-suspended in 0.5 mL of lysis buffer with lysozyme (20 mg/ml in water; Sigma Chemical Co.). The cells were incubated for 30 min at 37°C and then treated with 4 mL of IP buffer (50 mM HEPES-KOH pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% Na deoxycholate, 0.1% SDS and 1mg/mL phenylmethylsulfonyl fluoride (PMSF). The lysate was then sonicated using a Bioruptor (Diagenode) with 5 cycles of 7 minutes (30″ON/30″OFF) at high intensity. Cell debris was removed by centrifugation at 13000 rpm at 4°C for 201min and the resulting supernatant was used as cell extract for the immunoprecipitation. The range of the DNA size resulting from the sonication procedure was 100 - 500 bp, and the average DNA size was 300 bp. To immunoprecipitate StdE-DNA and StdF-DNA complexes, 800 µL of chromatin, 20 µL of Ultralink Immobilised protein A/G beads (Pierce) and 2 µL of the corresponding antibody Materials and Methods 73 were used. Control sample (mock-IP) was performed with no antibody added. Four samples were used for each antibody and four samples for the control. They were then incubated for 90 minutes at room temperature on a rotating wheel. Beads were transferred to a Spin-X column tube (Costar) and centrifuge at 3000 rpm for 1min. Beads were gently re-suspended in 500 µL of IP buffer and incubated on wheel for additional 3 min. This step was done twice. Beads were washed with 500 µL of IP salt buffer (50 mM HEPES-KOH pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% Na deoxycholate, 0.1% SDS, 1mg/mL phenylmethylsulfonyl fluoride (PMSF) and 500 mM NaCl), IP wash buffer and TE pH 8.0 (10 mM Tris-HCl and 1 mM EDTA) by re-suspending and re-centrifuging sample. The column was transferred to a fresh tube and the beads were re-suspended in 100 µL of elution buffer (50 mM Tris-HCl at pH 7.5, 10 mM EDTA, and 1% SDS) and incubated at 65ºC for 20 min. After centrifuging at 3000 rpm for 1 min the flow-through was treated with 10 µL of 40 mg/mL of a mixture of proteases (Pronase, Roche) made up in TBS pH7.6 (2.42 g/l Trizma base, 8 g/l NaCl). The samples were heat at 42ºC for 2h and 65ºC for 6 hours. The reactions were then kept at 4ºC overnight. The samples were clean up by using a PCR clean up Kit (Promega) and re-suspended in 50 µL of H2O. Chapter I 81 Expression of the std operon It has been described that the std fimbrial operon is tightly repressed in Salmonella enterica in a Dam-dependent manner (Balbontín et al., 2006). Dam– mutants show high expression of the std operon (Balbontín et al., 2006), and this expression is dependent on the HdfR protein (Jakomin et al., 2008). Moreover, bimodal expression for this operon has been suggested (Jakomin et al., 2008), since the authors observed two subpopulations in Dam– and SeqA– backgrounds. To further understand the expression conditions of the std operon, and based on the possibility of heterogeneous expression suggested by Jakomin et al., 2008, single cell analysis of stdA expression was performed using an stdA::gfp transcriptional fusion. This fusion is defective in StdA synthesis, since only the first 129 nt of the stdA gene are present in the construction. However, the downstream genes of the operon are expressed since the construct does not contain a transcriptional terminator downstream of the gfp gene. Flow cytometry assays were performed using strains stdA::gfp, ∆dam231 stdA::gfp (non-polar deletion of dam), ∆dam231 ∆hdfR stdA::gfp, ∆hdfR stdA::gfp. A representative experiment is presented in Figure C1. 1. Figure C1. 1 Regulation of the Dam-dependent std operon and role of HdfR. GFP fluorescence intensity in a strain carrying an stdA::gfp fusion in different genetic backgrounds (∆dam, wt, ∆hdfR). The Dam– mutant shows bimodal expression of the stdA::gfp fusion and the expression is dependent on HdfR, as described by (Jakomin et al., 2008). In a wild type background, a minor subpopulation (0.3% of the cells, StdON) appears to express stdA::gfp. The subpopulation disappears in a HdfR- background, underlining the essential role of HdfR in std regulation. The experiment shows that std expression is repressed by Dam methylation in a HdfR- dependent manner as previously described. Just ∼ 0.3 to 3% of cells produce stdA::gfp in a ∆dam ∆dam ∆hdfR ∆hdfRwt GFP fluorescence intensity Forward scatter StdON StdOFF wild type background. As observed for expression in a Dam– background, this expression is also dependent on HdfR. This observation indicates that expression of the std operon occurs only in a fraction of the population under laboratory conditions, suggesting bistable expression of the operon. Also, upon labeling Std fimbriae and under the fluorescence microscope, only a small fraction of Salmonella enterica wild type cells appeared to be Std-frimbriated (Figure C1. 2). Bacteria were labelled with rabbit anti-StdA serum (obtained from A. Baümler’s lab) and goat anti-rabbit IgG FITC conjugate for the detection of Std fimbriae. This observation, along with previous studies of phase variation of fimbrial operons (Hernday et al., 2004; Humphries et al., 2001; Norris et al., 1998), supports the idea that the std fimbrial operon undergoes heterogeneous expression. Figure C1. 2 Detection of StdA fimbriae in Salmonella enterica. Immunofluorescence microscopy for the detection of StdA fimbriae in Salmonella enterica in a wild type background. Left column (panels A and D) shows bacterial cells of Salmonella enterica (Hoechst stain). Panel B shows StdA detection with anti-StdA antiserum and goat anti-rabbit antibody conjugated to FITC (green signal). In panels C, E and F both channels were merged. A CB D FE Chapter I 83 Both flow cytometry and fluorescence microscopy approaches reveal that expression of stdA (and in consequence, expression of std operon) occurs in a small fraction of the population. Based on flow cytometry results, the StdON subpopulation varies from 0.3 to 3% of the population depending on the growth phase of the cells. However, no expression was detected when using the stdA::lacZ fusion on X-gal plates, despite the fact that the stdA::lacZ fusion behaves as expected in a Dambackground (∆dam stdA::lacZ ), showing blue colonies on X- gal plates. Transcriptional regulation of the std operon The mechanisms that prevent expression of the std operon outside the animal intestine are not well understood. It is already well-known that several factors repress std expression, such as Dam methylation (Balbontín et al., 2006; Chessa et al., 2008), RosE, an homolog of E. coli ArgR, and the GATC-binding protein SeqA (Chessa et al., 2008). On the other hand, the poorly known HdfR protein (a LysR-like factor) is required for std expression in S. enterica Dam– mutants (Jakomin et al., 2008), and it is also needed for std expression in a wild type background. During this Thesis, we had the chance of working in collaboration with the group of Dr. Mónica Delgado, INSIBIO, CONICET-UNT, Argentina. This group is interested in the regulatory mechanism of the signal transduction system RcsCDB. Previous results showed that a RcsC11 mutant undergoes constitutive activation of the RcsCDB system, and this activation leads to attenuation of S. typhimurium virulence (Costa & Antón, 2001; Mouslim & Groisman, 2003; Mouslim et al., 2003; 2004), partially through the repression of hilA, invF, sipC and invG genes (Delgado et al., 2006; Mouslim et al., 2004). Unpublished data from Delgado’s group reveals that the rcsC11 strain has a defective phenotype of adhesion to Caco-2 cells. These observations led us to speculate about a potential relation between constitutive expression of rcsB and std operon expression. In order to study whether the RcsCDB system controls std operon expression, we used the chromosomal stdA::gfp transcriptional fusion mentioned above. The gfp fusion was introduced into an RcsC11 background, and the distribution of GFP activity during exponential growth (O.D ∼0.8) was determined. Figure C1. 3. Distribution of stdA::gfp activity in wild type and rcsC11 backgrounds. The stdON subpopulation disappears in the strain rcsC11 stdA::gfp, indicating repression of std when rcsB is constitutively expressed. As observed in Figure C1. 3, constitutive expression of rcsB (in a RcsC11 mutant background) eliminates the StdON subpopulation, indicating that RcsB represses std expression. In order to further understand the regulatory mechanism that controls std expression through rcsB, the β-galactosidase activity assay of the stdA::lacZ translational fusion was monitored in different Dam– genetic backgrounds. Note that Dam– background was chosen in order to have enough stdA::lacZ expression. From the results summarized in Figure C1. 4, we can conclude that constitutive expression of rcsB (RcsC11 mutant) ends up in a 10-fold reduction of stdA::lacZ expression in the Dam– mutant. Figure C1. 4 Regulation of stdA::lacZ expression by RcsB A. β-galactosidase activity of stdA::lacZ translational fusion in different genetic backgrounds. B. Relative mRNA levels of stdA in different genetic backgrounds. RNA levels were determined for at least 3 biological replicates and a representative experiment is shown. GFP fluorescence intensity Forward scatter 0 500 1000 1500 dam dam rcsC11 dam rcsB56N dam tolB dam p-rcsB dam p-rcsB +IPTG stdA::lacZ Miller Units 0 0.5 1 1.5 dam dam rcsC11 dam rcsB56N dam tolB dam p-rcsB dam p-rcsB +IPTG 2 Relative RNA’s levels A B Chapter I 85 In contrast, inactivation of RcsB (the rcsBD56N strain contains a point mutation in the rcsB gene that inactivates RcsB by a single amino acid change) in a Dambackground increases stdA::lacZ expression two fold compared with the Dam– mutant (Figure C1. 4A). Previous reports from Dr. Delgado’s group demonstrated that the RcsCDB system is activated by a tolB mutation (TolB is involved in the integrity of cell envelope) and by rcsB overexpression from the prcsB plasmid (Mouslim & Groisman, 2003, Pescaretti et al., 2010). As we expected, the β-galactosidase activity levels obtained in dam tolB and dam prcsB (+ IPTG to overexpress rcsB) strains were similar to those observed in the dam rcsC11 (Figure C1. 4A). When IPTG was not added to the culture medium of the dam prcsB strain, the βgalactosidase activity levels produced were similar to those observed in the Dammutant (Figure C1. 4A). To further confirm these results, transcription of stdA was monitored by qRT-PCR. As shown in Figure C1. 4B, the results correlate well with those obtained by monitoring β-galactosidase activity, and the stdA expression levels in dam rcsC11, dam tolB and dam prcsB (+ IPTG) strains were significantly lower than those observed in the Dammutant. Moreover, the stdA transcription levels of the Dam- RcsBD56N double mutant were higher than those observed in the Dammutant. These results, together with the observations presented above, indicate that the RcsCDB system downregulates stdA gene expression, both in wild type and Dam– backgrounds. In addition, we studied whether std downregulation is due to specific binding of RcsB on the std promoter region. For this purpose, we first carried out a bioinformatic analysis using the std promoter region to identify a putative RcsB-binding site. The architecture of the std promoter including the GATC sequences, -10 and -35 modules, and the +1 transcription site was described by Jakomin et al. (2008) (Figure C1. 5A). As shown in Figure C1. 5A, a consensus RcsB-binding site, already described in other RcsB-regulated genes (Carballès et al., 1999; Mouslim & Groisman, 2003; Wehland & Bernhard, 2000), was found overlapping the -35 box of the std regulatory region. These findings suggest that RcsB binding to the -35 box may prevent binding of RNA polymerase, thereby inhibiting transcription of std. Binding of RcsB to this region was further confirmed by Dr. Delgado`s group, showing that migration of 300 pb and 735 bp PCR products (generated with EMSA-std300 FOR and EMSA-std REV; and EMSA-std735 FOR and EMSA-std REV pair of oligonucleotides respectively), both containing the std promoter and the putative RcsB-binding site, were retarded in Electrophoretic Mobility Shift Assay (EMSA) with increasing RcsB concentration (Farizano, JV, personal communication). A 234 bp DNA fragment was used as a negative control and, as expected, no retardation was observed (Figure C1. 5B). Altogether, these observations indicate that RcsB is a repressor of std expression, and that it acts by direct binding to the std promoter. Figure C1. 5 RcsB binds to std promoter A. In silico analysis showing RcsB putative binding sites in the std promoter. In blue are highlighted the three GATC sites in the std promoter, in green the start codon of the stdA gene, underlined are highlighted the -35, -10 and +1 sites and the RcsB putative binding site is distinct with a box. B. Electrophoretic Mobility Shift Assay (EMSA) of the std promoter with increasing concentrations of RcsB protein shows the slow mobility of the RcsB-stdA DNA complex compared with the control DNA. Expression and autoregulation of the transcription factor HdfR HdfR is essential for std expression in both the wild type (this study) and the Dam– background (Jakomin et al., 2008). Three GATC sites are present upstream the std promoter, in positions -243, -230 and -221 upstream of the transcription start of std. The methylation state of GATC sites can be affected by DNA-protein interactions. Therefore, Dam TTCACCCGCTTATAATTACATTAAAAAGTATTTCTTTGATGATTATTCTTAAATTACGATCATAATCATCGATCGTATCG ATCTATTTTTATTGTTTTAATTTGAATAAGAATTTTCTTACAACTGATTAGGCATTAGCTGAAATCAATATTTTTCGATG GAAAGTTCAGGTGCTTCGTTTAACACCAGGCGTTTATTATTCATACGAATCTTTTCTGAACGCTTTCATTAATACCCTCA GAGTTGTTTTCAGCCTTTGCAAAATAATTCTCATTCACCCAAAGGACATATTATCTATGCGTAATAAAATAATACTTGCC ATGGCGGCTGCCGGTATGATGTATGGTGCTTCTGTATTTGCCGCCGATACTACACCCACAGCAGGCCCGTTCGGCTCAGC -35 -10 +1 std- CCTTTGCAAAATAATTCTCATTC ams+ TATATTGAGAATAATCTTAATTT ugd + CGTAATGAAGATAATCTGAATTG RcsB/stdA DNA complex RcsB stdA DNA stdA DNA control DNA RcsB 234bp - 735bp - 300bp - A B Chapter I 87 methylation can act as a regulator of transcription by influencing binding of RNA polymerase or transcriptional factors (Wion & Casadesús, 2006). Previously in the lab, purified HdfR- His6 was used to study the interaction of HdfR with the std promoter region (from -295 to +11) in vitro, and to test the effect of Dam methylation on HdfR binding. Unpublished data show that HdfR binds the std regulatory region when the three GATCs of this region are nonmethylated, while it is unable to bind when the three GATCs are methylated or hemimethylated. Chromatin Immunoprecipitation (ChIP) coupled with quantitative PCR was used to investigate HdfR-std promoter interaction in vivo using the strain ∆dam hdfR-3xFLAG. A chromatin fragment containing the std promoter was 13 fold enriched in the inmunoprecipitated (IP) sample compared with the mock IP sample. Therefore, confirming in vitro data, HdfR binds to the non-methylated std promoter (Figure C1. 6). Figure C1. 6 Chromatin Immunoprecipitation (ChIP) coupled with quantitative PCR analysis of HdfR binding to the std promoter. However, and as shown above, std expression in wild type cells is only detected in a small subpopulation, whereas Dam– cells show high level of expression. We wondered if these differences in expression were due to a difference in the level of the expression of hdfR. To address this question, the β-galactosidase activity of an hdfR::lacZ fusion was measured in the wild type and in a Dam– strain in exponential and stationary cultures (Figure C1. 7). No significant differences were observed in hdfR expression, indicating that the differences in std expression observed between Dam– and wild type Salmonella cannot be explained by different hdfR expression levels. 0 mocK IP IP sample 10 15 5 hdfR3xflag Enrichment of DNA Figure C1. 7 β-galactosidase activity of hdfR::lacZ in exponential and stationary phase in a wild type and Dam – backgrounds. HdfR is a LysR-type transcriptional regulator, and these transcription factors commonly activate the expression of target genes but repress their own expression. To investigate whether HdfR represses its own expression, the intergenic region between hdfR and the upstream gene in the chromosome (yifE), which most likely contains the hdfR promoter, was cloned on the promoter-probe vector pIC552 (Macián et al., 1994), generating a transcriptional fusion with the lacZ gene (pIZ2320) (Figure C1. 8A). The activity of the phdfR::lacZ fusion was monitored in HdfR+ and HdfR– backgrounds to determine whether HdfR undergoes autogenous transcriptional regulation. Figure C1. 8 Autoregulation of hdfR. A. Diagram of the hdfR promoter region. The DNA fragment cloned on pIC552 is represented. The fragment contains the hdfR promoter region. B. β-Galactosidase activities of pIC552 and pIC552 phdfR in the presence and in the absence of HdfR. 0 100 200 300 wt O.D ~ 0.8 dam 400 500 O.D ~ 2 lacZ hdfRyifE hdfR::lacZ 0 1000 2000 3000 pIC552 hdfR + pIC552 phdfR 4000 hdfR - phdfR::lacZ Miller Units Miller Units A hdfRyifE phdfR lacZ phdfR pIZ2320 0 1000 2000 3000 pIC552 HdfR + pIC552 phdfR 4000 HdfR - phdfR::lacZpIC552 Miller Units B Chapter I 89 As shown in Figure C1. 8B, HdfR undergoes autogenous repression of the hdfR promoter, in a fashion analogous to other LysR-type transcription factors. Autogenous repression might ensure a constant intracellular level of the protein. Moreover, as it will be shown in Chapter 2, we have identified additional factors that affect hdfR expression. In order to determine whether high HdfR levels might affect std expression, we placed hdfR under the control of the heterologous promoter PLtetO. Despite the fact that PLtetO is a moderate promoter, higher hdfR expression levels were obtained (Figure C1. 9A), and the autorepression was avoided. The expression of the stdA::gfp and stdA::lacZ fusions was monitored by fluorescence analysis and β-galactosidase assays, respectively. Figure C1. 9 Constitutive expression of hdfR increase stdON subpopulation. A. Constitutive expression from the heterologous promoter PLtetO increases 100 fold the expression of hdfR compared with the wild type strain. Data are normalized to the RNA level obtained in wild type Salmonella cells (Value for wild type=1; not shown). B. Constitutive expression from the heterologous promoter PLtetO increases 10 fold the expression of the stdA::lacZ fusion compared with the wild type strain. C. Fluorescence intensity of the stdA::gfp fusion in a PLtetOhdfR shows an increase of the mean fluorescence and an increment of the stdON subpopulation. C. stdA::lacZ PLtetOhdfR on X-gal plates shows colonies with different stdA::lacZ expression (blue colonies pointed with arrows). 010 100 P LtetO hdfR 0 20 40 60 wt P LtetO hdfR stdA::lacZ BD Relative RNA’s levels A wt C GFP fluorescence intensity Forward scatter P LtetOhdfR As shown in Figure C1. 15B, the three GATC sites (GATC1, GATC2, and GATC3) are fully methylated in a wild type background. Considering that the majority of the population is in an StdOFF state, in the Southern the wild type strain largely corresponds to the StdOFF subpopulation. These results correlate well with previous assays of HdfR binding to the std promoter, in which binding occurred exclusively to non-methylated DNA. Identification of the DNA methylation pattern of the StdON subpopulation was difficult due to its small size (only 0,3 to 3%) of the population. StdE and StdF are located in the cytoplasm of Salmonella enterica StdE and StdF are homologues of known transcriptional regulators. Specifically, StdE shares 40-50% identity with the transcriptional activators GrlA from E.coli and CaiF from Enterobacter cloacae. In turn, StdF is related to the SPI-1 Salmonella protein SprB, a transcriptional regulator that represses the hilD promoter and activates the siiA promoter (López-Garrido & Casadesús, 2012). This homology with transcriptional regulators correlates well with the data presented in this chapter indicating that these proteins exert autogenous regulation of std transcription and also control hdfR expression (see above). The subcellular location of StdE and StdF was studied using 3xFLAG-tagged versions of the proteins in a Dambackground. Electrophoretic separation of cell fractions (cytoplasm, inner membrane and outer membrane) was performed, followed by Western blot analysis of the separated protein samples using a commercial anti-FLAG antibody. Based on the results obtained, we can conclude that both StdE and StdF are cytoplasmic proteins in Salmonella enterica (Figure C1. 16). Chapter I 97 Figure C1. 16 Subcellular localization of StdE and StdF proteins in S. enterica subcellular fractions. 3XFLAG versions of the proteins were localized with anti-FLAG Western hybridization for different subcellular fractions (cytoplasm, inner membrane and outer membrane). Search for an internal promoter in the std operon Because a fraction of the bacterial population expresses the std operon under conditions in which the main promoter appears to be inactive, we wondered whether expression of the StdE and StdF activators might occur independently of the std main promoter. Previous results in the lab had shown that the stdE and stdF genes were transcribed from the std promoter (upstream of stdA) in a Dambackground (López-Garrido & Casadesús, 2012). However, the same study raised the possibility that an internal promoter might exist that allow transcription of these two genes independently. To test this possibility we constructed a stdE::gfp fusion, in which the upstream genes (up to stdD) were deleted (together with the main promoter) (Figure C1. 17A). As observed in Figure C1. 17 B, low expression is observed when cells are run in a flow cytometer. We are confident that this expression, no matter if very low, is real, since it was repeatedly observed for 10 biological replicates run in different days, and previous studies had reported an expression of stdE that was not detected for the remaining std genes (López-Garrido & Casadesús, 2012). This result suggests the presence of a promoter that may drive the expression of stdE, and probably also of stdF. Several attempts have been done to identify this putative internal promoter. Cloning of different gene fragments onto the promoter-probe plasmid pIC552 did not reveal promoter activity. 5´RACE experiments performed in wild type and Dam– genetic backgrounds failed also in the identification of such promoter. We are currently working an alternative solution for this identification since we do think that an std internal promoter exists. StdE-3xFLAG StdF-3xFLAG Cytoplasm Inner Membrane Outer Membrane Figure C1. 17 Evidence of the existence of an internal promoter upstream stdE. A. Diagram of the ∆pStdA-stdC stdE::gfp construction. B. stdE::gfp expression in different genetic backgrounds. Dam- and HdfR- backgrounds are used to define stdE::gfp expression. ! ! ! ! ! ! ! ! ! ! ! ! ! ! GFP fluorescence intensity % cell count wt strain (no GFP) stdE::gfp ∆dam stdE::gfp ∆hdfR stdE::gfp ∆pStdA-stdC stdE::gfp stdE::gfp ∆pStdA-stdC stdE::gfp A B Chapter II: Impact of stdE and stdF overexpression in Salmonella enterica Chapter II 101 Transcriptomic analysis of S. enterica cells constitutively expressing stdEF In order to identify genes regulated by stdE and stdF, comparative transcriptomic analyses were performed in a strain that constitutively expressed stdEF and in a control strain carrying an in-frame deletion of both genes. To achieve moderate, constitutive expression of stdE and stdF, strain SV8141 was used (PLtetOstdEF). In this strain, transcription of both genes is driven by the PLtetO promoter (Lutz & Bujard, 1997; McQuiston et al., 2008). To avoid potential artefacts, the native stdA promoter and all the genes upstream of stdE in the std operon (Figure C2. 1A) were deleted, and the PLtetO promoter was placed upstream of stdE in the chromosome. The control strain contained the same insertion of the PLtetO promoter in the chromosome, lacking the upper part of the std operon but carrying also a deletion of stdE and stdF (SV8142; PLtetO∆stdEF, (Figure C2. 1B). Figure C2. 1. Diagrams of the std operon and of the strains SV8141 and SV8142. A. The std operon and surrounding genes in a Salmonella enterica wild type strain. Transcription of the operon is driven by the stdA promoter (PstdA). B. Diagrams of the std operon and surrounding genes in the strains used for transcriptomic analysis (PLtetOstdEF; SV8141 and PLtetO∆stdEF; SV8142) is shown. Transcriptomic analysis was performed using an array custom prepared by Dr. A. Juárez and co-workers (Paytubi et al., 2014), with RNA isolated from cells grown to stationary phase (OD600~2). Raw data from transcriptomic analysis have been deposited at the Gene Expression Omnibus (G.E.O) database (Edgar et al., 2002), with accession number GSE45488. The results show that, under the conditions used in the study, a large number of Salmonella loci showed differences in their RNA levels in an StdEF-dependent manner. Relevant data are summarized in Figure C2. 2 in which genes showing a higher than 4-fold difference in A PstdA std operon stdE stdFstdDstdCstdB stdA SL1344_3009 yohM B p LtetO stdE stdF SL1344_3009 yohM SL1344_3009 yohM SV8141 SV8142 p LtetO their RNA amount are depicted. Complete data from the transcriptomic analysis, along with gene descriptions and fold changes, are shown in Supplementary Table SM.1. Figure C2. 2 Pie chart summarizing the microarray data obtained. Numbers of genes up- and down-regulated, and physiological processes in which they are involved. As shown in Figure C2. 2, almost 200 genes showed a difference in expression higher than 4 fold upon constitutive expression of stdEF. About 85% of these genes appear to be downregulated by StdEF, suggesting that these proteins may be often repressors of gene expression in Salmonella. Among the down-regulated loci, genes involved in virulence and motility are highly represented. Regarding virulence, genes belonging to pathogenicity islands SPI-1 and SPI-4 are repressed by StdEF. As mentioned before, motility related genes are also found among down-regulated genes. The list includes not only flhDC, which encodes the master regulator of flagellar synthesis (Kutsukake et al., 1990), but also the flg, flh, fli, and motAB operons (Supplementary Table SM.1) are also repressed. FlhDC has been described as an activator of SPI-1 genes (Chubiz et al., 2010; Saini et al., 2008), which might explain the presence of strongly down-regulated SPI-1 genes in our transcriptomic analysis and in a previous study focused on the involvement of StdEF in Salmonella virulence (López-Garrido & Casadesús, 2012). Among down-regulated genes, we also find ygiD, annotated as a hypothetical protein, which seems to be involved in biofilm formation (Supplementary Table SM.1). The che operon and the trg and aer genes involved in chemotaxis were also found to be down-regulated (Figure C2.2). Among the up-regulated loci, we observe a high representation of genes involved in general metabolism, along with genes of the tra operon encoded on the pSLT plasmid (Supplementary Table SM.1). The only up-regulated gene that may be involved in Chapter II 103 virulence is the SL0982 locus, annotated as a “Gifsy-2 prophage attachment and invasion protein homolog”, which shows a 64-fold increase compared with the control strain. Interestingly, we also found yifA among the up-regulated genes, with an 8.4-fold change of expression. yifA (henceforth, hdfR) encodes the transcriptional activator HdfR, which is responsible for the expression of the std operon not only in a Dammutant (Jakomin et al., 2008) but also in a wild type background, as it was shown in Chapter 1. Regulation of hdfR by StdEF defines a positive feedback loop for autogenous control of the std operon. Additional results regarding mechanisms of self-regulation of the std operon were presented in Chapter 1. Validation of transcriptomic data To validate the transcriptomic data described above, we performed quantitative RT-PCR analysis of some of the genes differentially expressed in the presence of StdEF. We analysed expression of genes involved in motility (motA, flgE), conjugation (traA), chemotaxis (trg), virulence (hilA, sipB) and the transcriptional regulator of the std operon, hdfR. Data presented in Figure C2. 3 show that all the genes analysed undergo a change in expression in cells that constitutively express stdEF (PLtetOstdEF), compared with the control (PLtetO∆stdEF) and with wild type S. enterica. Changes in gene expression correlate well with those observed in the array, with fold-changes higher than 10-fold in all cases. Figure C2. 3 Validation of microarray data. RNA levels for some of the genes under stdEF control identified by microarray data in strains PLtetOstdEF and PLtetO∆stdEF. Data are normalized to the RNA level obtained in wild type Salmonella cells (Value for wild type=1; not shown). RNA levels were determined for at least 3 biological replicates and a representative experiment is shown. n=29n=164 A B p LtetO ΔstdEF p LtetO stdEF 1.000.01 0.10 0.001 10.00 C hilA motA sipB trA flgE trg hdfR Relative RNA’s levels Up regulated Down regulated Chemotaxis Motility Virulence Metabolism Conjugation Others pLtetO stdE stdF SL1344_3009 yohM pLtetO SL1344_3009 yohM pLtetO ΔstdEF pLtetO stdEF 1.00 0.01 0.100.001 10.00 hilA motA sipB trA flgE trg Relative RNA’s levels pLtetO ΔstdEF ∆hdfR pLtetO stdEF ∆hdfR A Although, as shown in chapter 1, the std operon is tightly repressed under the conditions employed in our microarray analysis (lab conditions), the expression levels of the analysed genes in the PLtetO∆stdEF strain vary slightly from the normalized Salmonella wild type strain. The existence of a small fraction of the population that expresses std (StdON cells) may explain this tiny difference, reinforcing the view that std is expressed under laboratory conditions in a subpopulation of bacterial cells. Functional validation of the transcriptomic data was also performed. In the case of Salmonella pathogenicity island-1 (SPI-1), regulation by StdEF had been previously proven in a study from our laboratory (López-Garrido & Casadesús, 2012). Constitutive expression of stdE and stdF had been shown to down-regulate SPI-1 genes, to decrease invasion of HeLa cells in vitro, and to attenuate Salmonella virulence upon infection of BALB/c mice (López- Garrido & Casadesús, 2012). The results presented in this Thesis correlate well with such data since a large number of SPI-1 genes appear to be down-regulated in the transcriptomic analysis. As a further functional validation of the transcriptomic analysis, we carried out tests of motility, biofilm formation, and conjugation in Salmonella strains that constitutively expressed the stdE and stdF genes (Figure C2. 4). The three phenotypes under examination respond to stdEF expression. Regarding motility, Salmonella cells expressing stdEF show no motility on a soft agar plate, indicating that constitutive expression of stdEF impairs motility of S. enterica cells. This phenotype correlates well with the transcriptomic data that show down-regulation of flagellar genes. Interestingly, the control strain (PLtetO∆stdEF) is more motile than Salmonella wild type cells (Figure C2. 4A). Control cells are wild-type cells in which the std operon has been deleted, as described in Materials and Methods and shown in Figure C2. 1B. This result confirms a role of the std operon in repression of motility, and reinforce the view that the std operon is expressed in a small fraction of the population (0.3-3%), under laboratory conditions. Chapter II 105 Figure C2. 4 Functional validation of the transcriptomic analysis. A. Motility agar plates (see Materials and Methods) were used to test the motility of the strains used in this study. The experiments were performed in triplicate, and a representative plate is shown. B. The biofilm formed by S. enterica SL1344 (wt) and the strains PLtetO-stdEF and PLtetO- ∆stdEF was stained with crystal violet as described in Materials and Methods. The experiments were performed in triplicate, and a representative assay is shown. C. Effect of stdEF on conjugal transfer of pSLT in microaerobiosis. The recipient was SV4938 in all matings. Donors were wild type Salmonella SL1344, and strains PLtetO-stdEF and PLtetO-∆stdEF. Data are averages and standard deviations from 3 independent matings. Balbontín et al. described that Salmonella Dam– mutants have a defect in motility (Balbontín et al., 2006). Since std overexpression impairs motility in S. enterica cells and std expression occurs in a Dam– mutants, we cannot rule out the possibility that Dam-dependent regulation of Salmonella motility is mediated by std expression. To corroborate this possibility we determined the motility of a Dam– mutant lacking stdEF. A B C wt pLtetOstdEF pLtetO∆stdEF wt p LtetO ΔstdEF p LtetO stdEF p LtetO ΔstdEF p LtetO stdEF wt wt pLtetO stdEF pLtetO ΔstdEF transconjugants/donors 10 -6 10 -9 10 -8 10 -7 ChIP-seq analysis of Salmonella enterica cells constitutively expressing StdEF-3xFLAG. The strain used for the ChIP-seq experiment expressed stdEF under the control of PLtetO (like the strain used in transcriptomic analysis) and contained an StdF variant labelled at its C- terminal end by a 3xFLAG epitope (PLtetOstdEF-3xFLAG; SV7850). This construction, depicted in Figure C2. 10, permitted detection of StdE with a cognate anti-StdE antibody, and of StdF using an anti-FLAG antibody. Because StdE is unaltered and StdFx3FLAG remains functional, transcription of the stdEF regulon was unaffected. Figure C2. 10 Diagram of the strain used for ChIP-seq analysis (PLtetOstdEF-3xFLAG). Chromatin immunoprecipitation was performed with cultures grown to stationary phase until OD600~2 (the same conditions used for microarray analysis). Bound proteins were crosslinked to DNA by formaldehyde treatment to form covalent unions. Cells were lysed and chromatin was sheared. Fragmentation of chromatin is a key point of ChIP sequencing since 300 bp fragments are required for the preparation of an optimal sequencing library, and for sequencing of the immunoprecipitated fragments. After immunoprecipitation of the specific proteins (and their bound DNA), de-crosslinking of the sample is performed to discard the proteins obtaining naked DNA fragments for the sequencing step. It is worth noting that the crosslinking step anchors the proteins to the bound DNA but also locks protein-protein interactions. Therefore our assays cannot discriminate between direct binding of the protein to DNA or indirect binding (through another protein). The controls used for the assay were: (i) total sheared and decrosslinked chromatin and (ii) mock IP sample. The mock IP is a sample that follows the same protocol as the IP sample but without adding the antibody. Despite the fact that the results from both controls were similar, we decided to use A PstdA std operon stdE stdFstdDstdCstdB stdA SL1344_3009 yohM B p LtetO stdE stdF SL1344_3009 3xFLAG SV9287 Chapter II 113 the mock IP control for analysis of results because we considered it more reliable than the total sheared chromatin. From the results shown in Supplementary Table SM.2 we concluded that StdE and StdF are able to bind to multiple sites in the Salmonella genome. Figure C2. 11 indicates the numbers of binding sites for StdE, StdF and for both StdE and StdF. StdE binds to 316 sites and StdF binds to 166 sites. StdE and StdF share the 20% of the sites. A higher number of DNA sequence reads were found upon inmunoprecipitation with the specific StdE antibody, suggesting that StdE binds to the Salmonella genome more efficiently than StdF, and that they bind in an independent manner. StdE StdF Figure C2. 11 Number of StdE, StdF and both StdE and StdF binding sites along Salmonella enterica genome. Looking at the binding sites for StdE and StdF, we observed that some of such sites may permit the interpretation of the data provided by the transcriptomic analysis. In other words, the StdEF binding patterns might help to explain StdEF-mediated regulation of a variety of physiological processes (e. g., by direct binding of the proteins to promoters or regulatory regions of genes). However in Supplementary Table SM.2 we can observe that there are also binding sites that do not seem to be involved in transcriptional regulation. A tentative explanation may be that the amount of StdE and StdF proteins present in our system is relatively high, and this might produce non-specific or non/physiological bindings to the DNA. An alternative possibility is that regulation might occur under conditions different from those used in this Thesis for transcriptomic analysis. Repression of the flagellar master operon by StdE and StdF binding As shown in Figure C2. 12, a significantly higher number of DNA sequencing reads were found in the promoter region of the flhDC genes for both StdE and StdF immunoprecipitations. As mentioned above, flhDC is the master operon for flagellar regulation (Kutsukake et al., 1990), and we have observed that StdEF–mediated regulation of flagellar genes is fully dependent on FlhC (Figure C2. 9). These results suggest that transcription of the flhDC operon may be regulated by direct StdE and StdF binding to its promoter. Figure C2. 12 StdE and StdF regulation of the flagellar system through FlhDC. Chip-seq data show StdE and StdF binding peaks at the flhDC promoter. StdE-C and StdF-C denote control reactions (mock IP) for StdE and StdF inmunoprecipitations, respectively, Regulation of chemotaxis by StdE and StdF is mediated by FlhDC Transcriptomic data described above showed that StdE and StdF repress the che operon, as well as the trg and aer genes involved in chemotaxis and aerotaxis. Analysis of the ChIP-seq results did not show a binding peak close to those operons. However, it is well known that chemotaxis regulation is dependent on the flhDC master regulator (Chilcott & Hughes, 2000). Based on this fact, we wondered whether regulation by StdEF might be mediated by FlhDC. To investigate this possibility, quantitative RT-PCR was performed to evaluate the mRNA levels of genes differentially expressed in the presence of StdE and StdF according to the transcriptomic data (cheA, cheM, che, and trg) in FlhC+ and FlhC– backgrounds. Chapter II 115 Figure C2. 13 Regulation of chemotaxis by StdE and StdF is mediated by flhDC. A. ChIP-seq data shows no relevant peaks surrounding the che operon. B. Ratios of the mRNA levels of genes involved in chemotaxis between the strains PLtetOstdEF and PLtetO∆stdEF. RNA levels were determined for at least 3 biological replicates and a representative experiment is shown. Data presented in Figure C2. 13B confirm that regulation of chemotaxis by StdEF is actually mediated by the flagellar master regulator FlhDC. This observation reminds of results described in both Salmonella and E. coli, where FlhDC is the transcriptional activator of σ28 (fliA) (among other second class operons). FliA drives transcription of class 3 flagellar operons, such as tar-tap-cheRBYZ and motAB-cheAW (Chilcott & Hughes, 2000). Similar observations have been made in other motility or chemotaxis related genes such as trg and aer (Ghigo, 2001; May & Okabe, 2008) whose transcription is dependent on RpoF (fliA-σ28) (Fitzgerald et al., 2015). StdE binds the hilD-coding region StdEF downregulate SPI-1 expression, and downregulation takes place through hilD at the post-transcriptional level (López-Garrido & Casadesús, 2012). We examined the data obtained from the ChiP-seq analysis in the SPI-1 region, and we found that only StdE appears to bind within the region, and that binding occurs at the end of the hilD coding region (Figure C2. 14). Based on the data from the ChIP-seq analysis (Supplementary Table SM.2) other minor binding peaks in SPI-1 region were found in sipB and sipC genes, which encode effector proteins. Outside SPI-1, no binding was detected in genes known to code for SPI-1 regulators (e. g., hilE). Figure C2. 14 StdE binding to the hilD coding region. Previous work on StdEF-mediated regulation of SPI-1 (López-Garrido & Casadesús, 2012) concluded that regulation involved post-transcriptional repression of HilD, and that the main player in this regulation was StdE. These data correlate well with our ChIP-seq data: only StdE seems to bind the region. Hence, we considered the possibility that binding of StdE to the hilD coding region might somehow affect the stability of hilD mRNA. In these experiments, expression of hilD was driven by the PLtetO promoter. This choice was based upon the fact that the levels of hilD mRNA upon stdEF constitutive expression were very low, thereby making difficult the calculation of hilD mRNA half-life. We first verified that the strain PLtetOstdEF PLtetOhilD was able to maintain the regulation pattern of SPI-1 genes in the same way as when hilD was transcribed from its own promoter. Data in Figure C2. 15A show that regulation occurs as expected, and that StdEF-mediated regulation of SPI-1 genes does not require hilD to be transcribed from its native promoter. Although the hilD mRNA level was lower in the strain that constitutively expressed StdEF, the stability of hilD mRNA was similar in both strains (t ½ for PLtetOstdEF PLtetOhilD and PLtetO∆stdEF PLtetOhilD is 2,98 and 2,82 min respectively), ruling out a mechanism of regulation involving hilD mRNA decay (Figure C2. 15B). Chapter II 117 Figure C2. 15 Stability of hilD mRNA. A. β-Galactosidase activity of the invH::lacZ fusion. B. Stability of hilD mRNA in PLtetO∆stdEF PLtetOhilD and PLtetOstdEF PLtetOhilD strains. Values are averages from 3 independent qRT-PCR reactions. Error bars are not shown because the estándar deviations were extremely small. The mechanism of post-transcriptional regulation of hilD upon StdE binding to its coding sequence cannot be easily explained. One speculation is that StdE binding at the downstream portion of the coding region might interfere with the DNA-RNA hybrid formed during transcription, reducing the transcription rate. Another possibility is that binding of StdE to the hilD coding region might regulate the expression of a putative antisense RNA, resulting in a decrease of hilD mRNA Control of pSLT conjugal transfer by StdEF Transcriptomic data (Supplementary Table SM.1) and phenotypic assays (Figure C2. 4C), show increased conjugal transfer of the Salmonella virulence plasmid (pSLT) if the donor strain constitutively expresses stdEF (SV8141). This increase in plasmid transfer may be correlated, at least in part, with up-regulation of tra operon genes upon StdEF constitutive synthesis. 0 0,01 0,1 PLtetO∆stdEF PLtetOhilD PLtetOstdEF PLtetOhilD Relative hilD mRNA level 612 10 8 4 2 0 Time (min) A B 0 2000 4000 6000 P LtetO stdEF hilD wt P LtetO hilD P LtetO ∆stdEF 8000 invH::lacZ Miller Units Figure C2. 16 ChIP-seq data showing pSLT plasmid region. In the upper part, peaks along the tra operon are shown. StdE and StdF binding peaks are detected along tra operon. In the lower part of the figure, the tra operon region is zoomedin. A closer look to the ChIP-seq data in the tra operon shows an increase of StdE and StdF binding peaks, mainly upstream of traA and upstream of traK (Figure C2. 16). In the microarray (Figure C2. 17), the most up-regulated gene in the tra operon is traA with a 24- fold increase. Gene Fold change PLtetOstdEF/PLtetO∆stdEF traA 24.3 traY 16.0 traL 11.3 traE 10.6 traP 6.5 traB 3.0 Figure C2. 17 Expression of tra genes in PLtetOstdEF vs PLtetO∆stdEF strains. Chapter II 119 Based on the strength of regulation, the binding peak closer to traA gene might be functional, and binding to the traA promoter might activate transcription. Moreover, traB appears as the less up-regulated gene in the tra operon, which suggests that the peak upstream traK and traB is not functional. StdEF-mediated regulation might thus involved the traA promoter. StdE and StdF binding sites identified in hdfR and std promoters Transcriptomic data and functional validation congruently indicate that increased StdEF synthesis in the strains that carry the PLtetOstdEF construct increase the level of hdfR expression, the transcriptional regulator of the std operon. Therefore, std expression increases the amount of the transcription factor needed for keeping the transcription of std operon, generating a self-regulatory positive loop (see Chapter 1). ChIP-seq upon StdE immunoprecipiation identified a peak upstream of the hdfR gene (Figure C2. 18), suggesting a direct activation of transcription by the StdE binding to the hdfR promoter. Figure C2. 18 ChIP- seq data obtained in the hdfR locus. Regarding std autoregulation, a binding peak was found upstream of the std promoter (Figure C2. 19), as expected according to the results presented in Chapter 1. However, we were lucky to identify this binding site, since the PLtetOstdEF-3xFLAG strain used for the ChIP-seq assay was constructed placing the PLtetO promoter upstream of the stdE gene and deleting the remaining sequences of operon upstream of stdE (which included the std promoter). The binding site identified by ChIP-seq is located 40 nt upstream of the deletion fragment, and 80 nt upstream of the first GATC site, suggesting a mechanism of autoregulation in which binding of StdF in this region favours std transcription. This result confirms previous data from the genetic screen (see Chapter 1) that outlined the importance of StdF for autoregulation of the std operon. It is interesting to point out that the binding site upstream of the std operon is the only one case in which StdF binds independently of StdE. The fact that StdF binding sites are mostly shared by StdE suggests a mechanism in which StdF binding to DNA might be accessory for regulation. An alternative possibility is that StdF does not bind directly DNA and its presence at the site is due to an interaction with other DNA-binding proteins (i.e., HdfR). For instance, StdF might act as a chaperone to facilitate binding of the regulator to cognate DNA. Figure C2. 19 ChIP-seq data showing StdF binding to std promoter region. Based on the ChiP-seq data, together with the results presented in Chapter 1, we propose a mechanism in which StdE binding to the hdfR promoter activates its expression, therefore maintaining HdfR levels high enough to allow expression from the std promoter, and to keep the autoregulatory loop active. StdF binding to the std promoter favours its transcription, and the accessory role of StdF might be either assisting HdfR binding, favouring a positive regulation, or by obstructing Dam methylation, which represses expression, therefore resulting in the positive regulation observed. Based in the results presented in Figure C2. 20 it might be suggested that StdF is involved in HdfR binding. In a wild type strain, both HdfR and StdF are essential for expression of the std operon. In a Dam– mutant, were there is no methylation, HdfR is essential but also StdF. Hence, when no blocking action is needed, StdF seems to be still essential for std expression. Chapter II 121 Figure C2. 20 Role of StdF in std activation. Heat maps represent GFP fluorescence intensity in S. enterica strains carrying an stdA::gfp fusion in different backgrounds. Based on these results and others described in Chapter 1, we confirm that StdE and StdF have a crucial role in autoregulation the std operon (Figure C2. 21). The involvement of these regulators takes place at two different levels. StdE binds the hdfR promoter region and increases hdfR transcription. It would be interesting to figure out if the increase is due to an active StdE action over the hdfR promoter or to blocking HdfR negative loop. On the other hand, StdF binds to the std promoter and may act as an auxiliary protein that cooperates with HdfR to activate std transcription. FS Log: FS Log 10 0 10 1 10 2 10 3 FL1 Log: FL1 Log 10 0 10 1 10 2 10 3 10 4 FS Log: FS Log 10 0 10 1 10 2 10 3 FL1 Log: FL1 Log 10 0 10 1 10 2 10 3 10 4 FS Log: FS Log ∆dam ∆stdEF ∆stdEF FS Log: FS Log 10 0 10 1 10 2 10 3 FL1 Log: FL1 Log 10 0 10 1 10 2 10 3 10 4 FS Log: FS Log 10 0 10 1 10 2 10 FL1 Log: FL1 Log 10 0 10 1 10 2 10 3 10 4 FS Log: FS Log dam pLtetostdEF ∆dam ∆stdEF ∆stdEF GFP fluorescence intensity ∆dam ∆dam ∆hdfR ∆hdfR StdON StdOFF wt Forward scatter PLtetOhdfR PLtetOhdfR ∆stdEF PLtetOstdEF ∆hdfR PLtetOstdEF signal transduction system. RcsB represses std expression. This regulation takes places not only in a Dam– background but also in the wild type strain, decreasing the fraction of StdON cells in the population. Based on the results presented here, this regulation is not dependent on Dam methylation, since repression is observed both in the presence and in the absence of Dam methylation. RcsB may exert transcriptional regulation by binding directly to the std promoter region, and in silico analysis has identified a putative binding site for RcsB near the -35 module of the std promoter. A footprint assay will be necessary to confirm the existence of RcsB binding to the putative binding site identified. The significance of RcsBCD-mediated repression of the std operon is difficult to grasp, even in a speculative manner, as the environmental signals that activate the RcsBCD signal transduction system remain largely unknown (Majdalani & Gottesman, 2005). Fractionation assays have identified StdE and StdF as cytoplasmic proteins, and in silico information reveals that StdE and StdF are homologues of known transcriptional regulators. StdE shares 40-50% identity with the transcriptional activators GrlA from E. coli and CaiF from Enterobacter cloacae. In turn, StdF is related to the SPI-1 Salmonella protein SprB, a transcriptional regulator that represses the hilD promoter and activates the siiA promoter (López-Garrido & Casadesús, 2012). Homology with transcriptional regulators correlates well with our results, although no DNA binding motifs were identified in silico. In 2012, López-Garrido & Casadesús proposed that the std operon may be the link between Dam methylation and SPI-1 regulation: the downregulation of SPI-1 expression observed in Dam– mutants (Balbontín et al., 2006) is suppressed upon deletion of two std genes, stdE and stdF (López-Garrido & Casadesús, 2012). This antecedent, combined with the fact that this Thesis has uncovered an active role of StdE and StdF as transcriptional activators of their own operon, led us to wonder whether these products of the std operon might have other roles in Salmonella enterica. To our surprise, microarray analysis revealed that StdE and StdF are global regulators of transcription, mainly as repressors. Constitutive expression of stdE and stdF downregulates not only SPI-1 expression as previously reported (López-Garrido & Casadesús, 2012) but also represses loci involved in motility and chemotaxis. On the other hand, StdEF activate transcription of the pSLT tra operon and, interestingly, of hdfR. Since StdEF activate hdfR expression and LysR-type transcription factors are often global regulators, we could not role out the possibility that some or all the global regulatory activities of StdE and StdF were actually performed by HdfR. However, we obtained Discussion 129 evidence that HdfR is not a global regulator but merely one of the many targets within the StdEF regulon. With hdfR transcriptional activation, the std operon may ensure its own transcription as HdfR is needed for std transcription. This regulatory pattern generates a positive feedback loop for std expression. A tempting speculation is that StdEF may increase the level of HdfR through direct binding of StdE to the regulatory region of the hdfR gene, thereby impairing autogenous repression by HdfR. Another possible intermediary in StdEF-mediated global regulation is the FlhDC master regulator, known to control invasion, motility and fimbriation (Saini et al., 2010). Furthermore FliZ, a product of FlhDC regulatory cascade, has been described as a postranscriptional regulator of SPI-1 through hilD (Chubiz et al., 2010). However, our results discard FlhDC as the connector between StdEF and their targets. FlhDC does regulate the flagellar system as well as chemotaxis (as expected), but none of the other processes under StdEF control. Because the potential intermediary factors considered above were not involved in global regulation by StdEF and we had previously observed that StdF acts as a transcription factor promoting std expression, we considered the option of direct regulation by StdEF. ChIP-seq analysis was thus performed to ascertain whether StdE and StdF are DNA-binding proteins. This analysis confirmed that StdE and StdF have DNA-binding ability, thus supporting the possibility that they may directly regulate gene expression. StdE binds twice more frequently than StdF but both proteins share 20% of the binding sites. This kind of assay does not distinguish direct binding from indirect binding. Hence, in certain cases we cannot rule out the involvement of both proteins in regulation. Although we cannot propose a mechanism regarding SPI-1 regulation, we have confirmed the occurrence of postranscriptional regulation of hilD as previously described (López-Garrido & Casadesús, 2012). Moreover, the relevant role that we assigned to StdE in SPI-1 regulation correlates well with the data presented by Lopez-Garrido et al. (2012) showing that the decrease of SP1-1 regulation is stronger when StdE is present than when only StdF is present. As discussed above, the ChIP-seq assay does not allow to distinguish between direct binding of a protein to the DNA and indirect binding through another protein. However, it would be interesting to study in depth how is it possible that StdE controls hilD expression by interacting with the downstream portion of its coding region. An attractive idea involves the hybrid DNA-RNA formed during transcription. StdE might impair RNA processing, thereby affecting translation of the HilD protein. As a consequence, disturbance of SPI-1 regulation might occur, and also disruption of the hilD self-regulatory loop. Regulation of the expression of a putative antisense RNA can be also considered, but in silico analysis based on data provided by Jay Hinton's lab seem to discard the possibility of an additional transcription start site aside from those already known in the region. Concerning the regulation of motility and chemotaxis, the ChIP-seq data raise the possibility that both StdE and StdF bind directly to the flhDC promoter. Regulation at the highest level in the hierarchy of the flagellar network (FlhDC) seems to explain why all the operons involved in regulation of motility appear to be under StdEF control. Regulation of chemotaxis is also controlled through FlhDC, probably involving the Class III transcription factor, FliA. A priori, it seems to make sense that the motility system is not active if the ability to sense the environment is not working and vice versa. Interestingly, we have observed that the regulation of the flagellar system mediated by Dam methylation is, at least partially, due to StdEF, in a way reminiscent of the crosstalk previously described for SPI-1 regulation. Even though StdE and StdF appear to be mainly repressors of transcription, it is noteworthy that the conjugal transfer operon tra and the hdfR gene are activated by StdEF. We cannot propose a mechanism for StdEF-mediated regulation of conjugation, but it is clear that StdE and StdF binds massively along the tra operon. We cannot ascertain if one or both proteins are involved in tra operon control nor define the specific target(s). However, based on the expression level of the target genes, the traA regulatory region might be the most appropriate for regulation by StdE, StdF, or both. At this point, and considering that the most widespread role of StdE and StdF appears to be to act as repressors of transcription, one may consider the possibility that up-regulation mediated by stdEF might be actually performed by repressing a repressor. We have already mentioned that StdEF-dependent regulation of hdfR may involve impairment of HdfR binding to its own promoter. Something similar may happen with StdEF-dependent regulation of tra operon. Unfortunately, among the transcriptomic data we have not found an appropriate candidate. However, we have observed that Dam methylation-dependent regulation of the tra operon is, as described previously for the flagellar regulation, also due to StdEF. So StdEF are involved in Dam dependent regulation of SPI-1 (López-Garrido & Casadesús, 2012), flagellar synthesis and conjugation. In the case of conjugation, StdEF- Discussion 131 mediated activation of tra might boost Lrp-mediated activation of traJ (Camacho & Casadesús, 2002) and counter or balance FinP-mediated repression (Camacho & Casadesús, 2005). The latter two mechanisms of conjugation control are also Dam-dependent, albeit in opposite ways: Dam methylation activates transcription of finP and represses transcription of traJ. The existence of overlapping controls acting in opposite manners may illustrate, like many other examples, the complexity of bacterial regulatory networks. A major outcome of this Thesis work is that the study provides a novel example of phenotypic variation within a clonal population. Two bacterial subpopulations, made of fimbriated and non-fimbriated Salmonella cells, appear to exist. Formation of two subpopulations may be viewed as a preadaptation of Salmonella enterica to perform two types of infection, acute and chronic. Because Std fimbriae permit adhesion to the epithelium of the caecum (Weening et al., 2005), StdON cells will be able to colonize the large intestine, and this colonization may result in persistent or chronic infection. In turn, StdOFF cells will be able to undergo invasion of the small intestine epithelium, causing salmonellosis or systemic infection. The fact that the StdON and StdOFF subpopulations differ in additional phenotypic traits may contribute to adaptation by fine-tuning the interaction of each subpopulation with a specific host environment. CONCLUSIONS Conclusions 135 1. Populations of Salmonella enterica cells undergo bistable expression of the std fimbrial operon. The StdON subpopulation is small in the wild type (0.3-3% cells) under laboratory conditions. 2. Formation of the StdON subpopulation requires the StdE and StdF proteins, which are products of the promoter-distal genes of the std operon. StdE is necessary to activate transcription of the hdfR gene, which encodes the main transcriptional activator of the std operon. In turn, StdF may be necessary to facilitate binding of HdfR to the std promoter. 3. StdE and StdF are global regulators in Salmonella enterica, and regulate SPI-1 expression, flagellar synthesis, chemotaxis, biofilm formation, and conjugal transfer of the virulence plasmid. 4. StdE and StdF are DNA-binding proteins that show homology with other transcriptional regulators, and may control transcription by direct binding to DNA. An exception is the hilD gene, which undergoes postranscriptional regulation under StdE control. The mechanism underlying hilD regulation is unknown. 5. The StdOFF subpopulation is able to invade epithelial cells, while the StdON subpopulation is deficient in epithelial cell invasion. This difference may endowe the StdOFF subpopulation with the ability to cause salmonellosis or systemic infection. 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