Transcription-driven DNA supercoiling counteracts H-NS-mediated gene silencing in bacterial chromatin.

In all living cells, genomic DNA is compacted through interactions with dedicated proteins and/or the formation of plectonemic coils. In bacteria, DNA compaction is achieved dynamically, coordinated with dense and constantly changing transcriptional activity. H-NS, a major bacterial nucleoid structuring protein, is of special interest due to its interplay with RNA polymerase. H-NS:DNA nucleoprotein filaments inhibit transcription initiation by RNA polymerase. However, the discovery that genes silenced by H-NS can be activated by transcription originating from neighboring regions has suggested that elongating RNA polymerases can disassemble H-NS:DNA filaments. In this study, we present evidence that transcription-induced counter-silencing does not require transcription to reach the silenced gene; rather, it exerts its effect at a distance. Counter-silencing is suppressed by introducing a DNA gyrase binding site within the intervening segment, suggesting that the long-range effect results from transcription-driven positive DNA supercoils diffusing toward the silenced gene. We propose a model wherein H-NS:DNA complexes form in vivo on negatively supercoiled DNA, with H-NS bridging the two arms of the plectoneme. Rotational diffusion of positive supercoils generated by neighboring transcription will cause the H-NS-bound negatively-supercoiled plectoneme to "unroll" disrupting the H-NS bridges and releasing H-NS.

Dynamics of macrophage polarization support Salmonella persistence in a whole living organism.

Numerous intracellular bacterial pathogens interfere with macrophage function, including macrophage polarization, to establish a niche and persist. However, the spatiotemporal dynamics of macrophage polarization during infection within host remain to be investigated. Here, we implement a model of persistent Salmonella Typhimurium infection in zebrafish, which allows visualization of polarized macrophages and bacteria in real time at high resolution. While macrophages polarize toward M1-like phenotype to control early infection, during later stages, Salmonella persists inside non-inflammatory clustered macrophages. Transcriptomic profiling of macrophages showed a highly dynamic signature during infection characterized by a switch from pro-inflammatory to anti-inflammatory/pro-regenerative status and revealed a shift in adhesion program. In agreement with this specific adhesion signature, macrophage trajectory tracking identifies motionless macrophages as a permissive niche for persistent Salmonella. Our results demonstrate that zebrafish model provides a unique platform to explore, in a whole organism, the versatile nature of macrophage functional programs during bacterial acute and persistent infections.

Mapping Transposon Insertion Sites by Inverse Polymerase Chain Reaction and Sanger Sequencing.

Inverse polymerase chain reaction (PCR) is a method designed to amplify a segment of DNA for which only a portion of the sequence is known. The method consists of circularizing the DNA fragment by self-ligation and performing PCR with primers annealing inside the known sequence but pointing away from each other (hence the technique is also called "inside-out PCR"). Here we describe how inverse PCR can be used to identify the site of transposon insertion in the bacterial chromosome. This protocol, implemented here with a class of transposons generating reporter gene fusions, involves (i) preparing genomic DNA from the strain harboring the unknown insertion, (ii) cleaving the genomic DNA with a restriction enzyme, (iii) performing a ligation reaction under conditions favoring circularization of the DNA fragments, and (iv) performing inverse PCRs with inside-out primers annealing near either or both termini of the transposon. This last step results in the amplification of the chromosomal sequences immediately adjacent to the transposon, which can then be identified by Sanger sequencing. The protocol can be performed in parallel on several strains providing an effective and economic way for rapidly identifying multiple transposon insertion sites.

Use of Transposable Reporters in the Analysis of Bacterial Regulatory Networks.

Transposable elements are genetic entities that have the capacity to promote their own translocation from one site to another within a genome. Initially discovered in Zea mays by Barbara McClintock at the Cold Spring Harbor Laboratory, transposable elements have been found to populate the genomes of all forms of life. In bacteria, the discovery of transposons significantly enhanced genetic analyses; they have been widely used to make insertion mutants and have inspired elegant strategies for strain construction and in vivo genome engineering. In one application, transposons have been modified to include a reporter gene engineered in such a way that the reporter can become fused to a chromosomal gene upon inserting randomly in the bacterial chromosome. Screening this type of transposon library for expression of the reporter under different conditions allows identifying fusions that respond coordinately to a specific treatment or stress condition. Characterization of these fusions provides a genome-wide snapshot of the organization of a bacterial regulatory network.

Generating Libraries of Random lacZY or GFP Gene Fusions in the Bacterial Chromosome: Screening for Differentially Expressed Fusions.

Transposable elements engineered to generate random gene fusions in the bacterial chromosome are valuable tools in the study of gene expression. In this protocol, we describe the use of a new series of transposons designed to obtain random fusions to either the lacZY operon or the gene for superfolder green fluorescent protein (sfGFP). Transposition is achieved through the activity of the hyperactive variant of Tn5 transposase (Tnp) whose gene is positioned in cis with respect to the transposable module and under the control of the anyhydrotetracycline (AHTc)-inducible Ptet promoter. The transposable module comprises a promoter-less lacZY operon or the sfGFP gene, with or without the lacZ or sfGFP ribosome-binding site, plus a kan gene for selection. The transposon-transposase unit is harbored on an R6K-based suicide plasmid. The plasmid is introduced into recipient cells by electro-transformation and the synthesis of Tn5 Tnp is induced transiently by including AHTc in the recovery medium. Cells are then plated on kanamycin-supplemented medium (without AHTc) where the plasmid DNA is lost and only cells in which transposition has occurred can form colonies. Fusions are detected by screening for colony color on lactose indicator plates (lacZ transposition) or monitoring green fluorescence (sfGFP transposition). Depending on whether the reporter gene carries or lacks the ribosome binding sequence, the fusions obtained will either be transcriptional or translational. Parallel screening of colonies grown in the absence and presence of a drug (or condition) eliciting a global regulatory response allows identification of fusions specifically activated or repressed as part of such response.

Scarless DNA Recombineering.

The method described here allows editing of the bacterial genome without leaving any secondary changes (scars) behind. This method uses a tripartite selectable and counterselectable cassette comprising an antibiotic-resistance gene (cat or kan) and the tetR repressor gene linked to a Ptet promoter-ccdB toxin gene fusion. In the absence of induction, the tetR gene product represses the Ptet promoter, preventing ccdB expression. The cassette is first inserted at the target site by selecting for chloramphenicol or kanamycin resistance. It is subsequently replaced by the sequence of interest by selecting for growth in the presence of anhydrotetracycline (AHTc), which inactivates the TetR repressor thereby causing CcdB-induced lethality. Unlike other CcdB-based counterselection schemes, which require specifically designed λ-Red delivery plasmids, the system described here uses the popular plasmid pKD46 as the source of λ-Red functions. This protocol allows a wide variety of modifications, including the intragenic insertion of fluorescent or epitope tags, gene replacements, deletions, and single base-pair substitutions, to be made. In addition, the procedure can be used to place the inducible Ptet promoter at a chosen position in the bacterial chromosome.

Recombineering 101: Making an in-Frame Deletion Mutant.

DNA recombineering uses phage λ Red recombination functions to promote integration of DNA fragments generated by polymerase chain reaction (PCR) into the bacterial chromosome. The PCR primers are designed to have the last 18-22 nt anneal on either side of the donor DNA and to carry 40- to 50-nt 5' extensions homologous to the sequences flanking the chosen insertion site. The simplest application of the method results in knockout mutants of nonessential genes. Deletions can be constructed by replacing a portion or the entirety of a target gene with an antibiotic-resistance cassette. In some commonly used template plasmids, the antibiotic-resistance gene can be coamplified with a pair of flanking FRT (Flp recombinase recognition target) sites that, following insertion of the fragment into the chromosome, allow excision of the antibiotic-resistance cassette via the activity of the site-specific Flp recombinase. The excision step leaves behind a "scar" sequence comprising an FRT site and flanking primer annealing sequences. Removal of the cassette minimizes undesired perturbations on the expression of neighboring genes. Even so, polarity effects can result from the occurrence of stop codons within, or downstream of, the scar sequence. These problems can be avoided by the appropriate choice of the template and by designing primers so that the reading frame of the target gene is maintained past the deletion end point. This protocol is optimized for use with Salmonella enterica and Escherichia coli.

DNA Recombineering Applications.

The ability to manipulate the bacterial genome is an obligatory premise for the study of gene function and regulation in bacterial cells. The λ red recombineering technique allows modification of chromosomal sequences with base-pair precision without the need of intermediate molecular cloning steps. Initially conceived to construct insertion mutants, the technique lends itself to a wide variety of applications including the creation of point mutants, seamless deletions, reporter, and epitope tag fusions and chromosomal rearrangements. Here, we introduce some of the most common implementations of the method.

Converting an FRT-Tagged Gene into a Fluorescent Protein Gene Fusion by Flp-Mediated Site-Specific Recombination.

This protocol uses conditional plasmids carrying the open reading frame (orf) of either superfolder green fluorescent protein (sfGFP) or monomeric Cherry (mCherry) fused to a flippase (Flp) recognition target (FRT) site. In cells expressing the Flp enzyme, site-specific recombination between the plasmid-borne FRT and an FRT "scar" in a target gene in the bacterial chromosome results in chromosomal integration of the plasmid with the concomitant in-frame fusion of the target gene to the fluorescent protein orf. This event can be positively selected using an antibiotic-resistance marker (kan or cat) present on the plasmid. This method is slightly more laborious than generating the fusion directly by recombineering and has the limitation that the selectable marker is no longer removable. However, it has the advantage that it can be more readily integrated in mutational studies, allowing conversion of in-frame deletions resulting from Flp-mediated excision of a drug-resistance cassette (e.g., all those of the "Keio collection") into fluorescent protein fusions. Furthermore, in studies that require that the amino-terminal moiety of the hybrid protein keeps its biological activity, presence of the FRT "linker" sequence at the fusion junction makes it less likely for the fluorescent domain to sterically interfere with the folding of the amino-terminal domain.

Construction of Single-Copy Fluorescent Protein Fusions by One-Step Recombineering.

We describe a simple recombineering-based procedure for generating single-copy gene fusions to superfolder GFP (sfGFP) and monomeric Cherry (mCherry). The open reading frame (orf) for either protein is inserted at the targeted chromosomal location by λ Red recombination using an adjacent drug-resistance cassette (kan or cat) for selection. The drug-resistance gene is flanked by flippase (Flp) recognition target (FRT) sites in direct orientation, which allows removal of the cassette by Flp-mediated site-specific recombination once the construct is obtained, if desired. The method is specifically designed for the construction of translational fusions producing hybrid proteins with a fluorescent carboxyl-terminal domain. The fluorescent protein-encoding sequence can be placed at any codon position of the target gene's mRNA where the fusion produces a reliable reporter for gene expression. Internal and carboxyl-terminal fusions to sfGFP are suitable for studying protein localization in bacterial subcellular compartments.

Working with Bacteria, Phage, and Plasmids.

Methods for the in vivo manipulation of bacterial genomes have improved greatly in recent years because of the discovery of new mechanisms and the gigantic leap forward in DNA-sequencing technology. Many cutting-edge approaches still rely on a variety of technical routines, the correct implementation of which is critical for the success of an experiment. Here, we introduce some of these procedures as used for Escherichia coli and Salmonella enterica We begin by reviewing the aspects of the biology of these two species that are most relevant for their manipulation in the laboratory.

Quick Transformation with Plasmid DNA.

Genomic engineering of Escherichia coli and Salmonella often requires introducing plasmids into strains obtained during the intermediate stages of the process. Such strains are typically transformed only once, making the preparation of large batches of competent cells for storage purposes unnecessary. Here, we describe a simple scaled-down procedure for transforming E. coli or Salmonella with plasmid DNA that uses as little as 2 mL of culture.

Basic Bacteriological Routines.

In experimental bacteriology, bacteria are generally manipulated, stored, and shipped in the form of cultures. Depending on various factors, including strain genotype, storage and shipping methods, and manipulator skills, the culture may contain genetic variants or simply contaminants. It is therefore important to begin an experiment by streaking the culture on an agar plate. Streaking, a technique to disperse bacterial cells on the surface of the agar, serves the purpose of isolating individual colonies. A colony originates from a single cell and is a nearly pure culture. On rich LB medium after 24 h of incubation at 37°C, a colony of Salmonella contains ∼5 × 108 cells (about 29 generations). Streaking is also required in experiments that themselves generate single colonies as a result of selection (e.g., when constructing strains or introducing plasmids). Except in the few instances in which the selection efficiently kills all counter-selected bacteria, colonies growing on the selective plates are contaminated, sometimes heavily, with cells from the bacterial lawn. "Purifying" the colonies arising in such experiments by streaking on selective plates is therefore a mandatory step. Here, we show how this can be conveniently done using simple toothpicks. We also briefly describe the steps involved in inoculating liquid cultures, spreading plates, and replica plating.

Preparing Plasmid DNA from Bacteria.

The most common method for isolating plasmid DNA is derived from an alkaline lysis procedure. The procedure exploits the differential partitioning of plasmid and chromosomal DNA when denatured by alkali and subsequently renatured by neutralization of the medium. The circular covalently closed nature of plasmid DNA allows the denatured DNA strands to quickly find each other and reanneal during the renaturation step. This is not the case for chromosomal DNA, which, upon neutralization, aggregates with denatured proteins through hydrophobic interactions. As a result, plasmid DNA remains in solution and can be easily separated from most of the other macromolecules that coprecipitate. For the subsequent purification step, one can use the silica membrane technology integrated in many commercial kits. This technology exploits the ability of DNA to bind to silica in the presence of chaotropic salts. DNA is retained by a silica-based column, whereas most of the polysaccharides and proteins flow through. After wash steps to eliminate residual contaminants and salts, DNA is selectively eluted under low-salt conditions. A kit-free but relatively more cumbersome alternative to this procedure is the traditional phenol-chloroform extraction method followed by ethanol precipitation. Both methods are detailed here.

Working with Phage P22.

Transduction experiments in Escherichia coli and Salmonella are usually performed with virulent phage variants. A widely used P1 mutant, called P1 vir, carries one or more uncharacterized mutations that prevent formation of lysogens. In the case of P22, by far the most frequently used variant is named P22 HT105/1 int-201 This phage has a high transducing (HT) frequency due to a mutant nuclease with lower specificity for the pac sequence. As a result, ∼50% of the P22 HT phage heads carry random transducing fragments of chromosomal DNA. The int mutation reduces the formation of stable lysogens. The basic steps in handling the P22 HT105/1 int-201 phage and in performing transduction experiments in Salmonella are described here.

Preparing Bacterial Genomic DNA.

We describe two alternative procedures for purifying bacterial chromosomal DNA. The first procedure incorporates the use of a commercial kit based on silica membrane technology. This approach relies on the selective binding of DNA to a silica-based column in the presence of chaotropic salts (guanidine salts). Polysaccharides and proteins do not bind well to the column and flow through. Their residual traces, along with the guanidine salts, are removed during alcohol-based wash steps. The DNA is then selectively eluted under low-salt conditions. This method is quick and easy and yields genomic DNA suitable for most downstream applications. The second procedure, implemented for many years in our laboratory before the appearance of commercial kits, is based on the ability of the cationic detergent cetyl trimethyl ammonium bromide (CTAB) to complex with polysaccharides and proteins, producing an emulsion that can be removed by chloroform-isoamyl alcohol extraction. This procedure is therefore especially suited to working with Gram-negative bacteria, which typically produce large amounts of polysaccharides. Its main advantage, besides cost-effectiveness, is the high yield of the DNA obtained; its main disadvantage is that the workflow is relatively cumbersome.

Pervasive transcription enhances the accessibility of H-NS-silenced promoters and generates bistability in Salmonella virulence gene expression.

In Escherichia coli and Salmonella, many genes silenced by the nucleoid structuring protein H-NS are activated upon inhibiting Rho-dependent transcription termination. This response is poorly understood and difficult to reconcile with the view that H-NS acts mainly by blocking transcription initiation. Here we have analyzed the basis for the up-regulation of H-NS-silenced Salmonella pathogenicity island 1 (SPI-1) in cells depleted of Rho-cofactor NusG. Evidence from genetic experiments, semiquantitative 5' rapid amplification of complementary DNA ends sequencing (5' RACE-Seq), and chromatin immunoprecipitation sequencing (ChIP-Seq) shows that transcription originating from spurious antisense promoters, when not stopped by Rho, elongates into a H-NS-bound regulatory region of SPI-1, displacing H-NS and rendering the DNA accessible to the master regulator HilD. In turn, HilD's ability to activate its own transcription triggers a positive feedback loop that results in transcriptional activation of the entire SPI-1. Significantly, single-cell analyses revealed that this mechanism is largely responsible for the coexistence of two subpopulations of cells that either express or do not express SPI-1 genes. We propose that cell-to-cell differences produced by stochastic spurious transcription, combined with feedback loops that perpetuate the activated state, can generate bimodal gene expression patterns in bacterial populations.

6S RNA-Dependent Susceptibility to RNA Polymerase Inhibitors.

Bacterial small RNAs (sRNAs) contribute to a variety of regulatory mechanisms that modulate a wide range of pathways, including metabolism, virulence, and antibiotic resistance. We investigated the involvement of sRNAs in rifampicin resistance in the opportunistic pathogen Staphylococcus aureus. Using a competition assay with an sRNA mutant library, we identified 6S RNA as being required for protection against low concentrations of rifampicin, an RNA polymerase (RNAP) inhibitor. This effect applied to rifabutin and fidaxomicin, two other RNAP-targeting antibiotics. 6S RNA is highly conserved in bacteria, and its absence in two other major pathogens, Salmonella enterica and Clostridioides difficile, also impaired susceptibility to RNAP inhibitors. In S. aureus, 6S RNA is produced from an autonomous gene and accumulates in stationary phase. In contrast to what was reported for Escherichia coli, S. aureus 6S RNA does not appear to play a critical role in the transition from exponential to stationary phase but affects σB-regulated expression in prolonged stationary phase. Nevertheless, its protective effect against rifampicin is independent of alternative sigma factor σB activity. Our results suggest that 6S RNA helps maintain RNAP-σA integrity in S. aureus, which could in turn help bacteria withstand low concentrations of RNAP inhibitors.