Recombination within genomic regions of (A) <i>S. alvi</i> and (B) <i>G. apicola</i>.

<p>Sequence divergence at all sites is plotted for pairwise comparisons in a sliding window of 200 nucleotides with a step size of 50 nucleotides. Arrows indicate genes for which intragenic recombination between pairs has been detected with the program Geneconv. Arrow colors correspond to the different pairwise comparisons. (A) Sequence divergence over the urease gene cluster of <i>S. alvi</i>. Note the drastic decrease in sequence divergence between O02 and the other two SAGs in urease gene E. Recombination seems also to have occurred in the gene encoding a hypothetical protein (hypo) at the end of the gene cluster. (B) Sequence divergence over a genomic region of <i>G. apicola</i>. I20 and P17 show variation in sequence divergence, particularly in genes, for which intragenic recombination was detected. In contrast, no evidence for recombination could be found between B02 and wkB2.</p

Genome features of SAGs and comparisons to their reference genome.

a<p><i>S. alvi</i> SAGs were compared to the reference genome of strain wkB2. <i>G. apicola</i> SAGs were compared to the reference genome of strain wkB1.</p>b<p>Estimated genome coverage is based on the presence of 206 and 189 genes constituting the minimal, essential gene set of wkB1 and wkB2, respectively. The minimal gene sets were determined as described previously <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1004596#pgen.1004596-Engel2" target="_blank">[22]</a>, <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1004596#pgen.1004596-Gil1" target="_blank">[97]</a>.</p>c<p>Id, identities.</p>d<p><i>d</i>S, rate of synonymous substitutions per synonymous site, averaged over all shared genes.</p>e<p><i>d</i>N/<i>d</i>S, ratio of <i>d</i>N (rate of non-synonymous substitutions per non-synonymous site) to <i>d</i>S.</p><p>Genome features of SAGs and comparisons to their reference genome.</p

Taxonomic classification of 126 bacterial cells sorted from midguts and ileums of honey bees.

<p>Classification is based on best BLASTN hit of partial 16S rRNA sequences. <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1004596#pgen.1004596.s009" target="_blank">Table S1</a> provides a complete list of all best BLASTN hits.</p

Sequence divergence and phylogenetic analysis of protein-encoding genes of (A and C) <i>S. alvi</i> SAGs and (B and D) <i>G. apicola</i> SAGs.

<p>Pairwise sequence divergence was measured by estimated rates of synonymous substitutions per site (<i>d</i>S) for (A) 226 orthologs of <i>S. alvi</i> and (B) 348 orthologs of <i>G. apicola</i>. Pairwise <i>d</i>S values of SAGs and reference genomes of <i>A. mellifera</i> isolates (in (A), <i>S. alvi</i> wkB2; in (B), <i>G. apicola</i> wkB1) are plotted on the x-axes. Pairwise <i>d</i>S values of SAGs and reference genomes of bumble bee (<i>Bombus bimaculatus</i>) isolates (in (A), <i>S. alvi</i> wkB12; in (B), <i>G. apicola</i> wkB11) are plotted on the y-axes. Mean <i>d</i>S values are given in parentheses (SAG compared to <i>A. mellifera</i> isolate; SAG compared to <i>B. bimaculatus</i> isolate). For visualization purposes, genes with unrealistically high <i>d</i>S values were excluded from representation. Complete data is presented in <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1004596#pgen.1004596.s005" target="_blank">Figure S5</a>. Note that genes with <i>d</i>S value ≥3 can be considered at or near saturation due to the four possible bases in the genetic code. (C and D) Maximum likelihood trees based on the concatenated alignments of 114 and 211 conserved orthologs of <i>S. alvi</i> and <i>G. apicola</i>, respectively. Values above branches represent bootstrap values ≥80 for 100 replicates. Values below branches indicate the percentage of single-gene trees with congruent topology at this node.</p

Ternary plots of sequence divergence at synonymous sites.

<p>Plots show gene-to-gene variation for synonymous substitution frequencies, for (A) 239 orthologs of <i>S. alvi</i> SAGs and (B) 400 orthologs of <i>G. apicola</i> SAGs. Each dot represents a triplet of orthologs. The sum of all three pairwise <i>d</i>S values have been normalized to 1 and plotted onto the ternary plot. The mean relative <i>d</i>S value for each pair is shown on the axes of the ternary plot. Distributions of absolute <i>d</i>S values are shown in histograms for each pair next to the ternary plot. X-axes show <i>d</i>S value categories, y-axis show number of genes. Colors represent the maximum absolute <i>d</i>S value in each comparison, with yellow for <i>d</i>S<0.1, orange for <i>d</i>S≥0.1, and red for <i>d</i>S≥1. Spread of each ternary plot is the median distance to the average point. <i>d</i>S values have been restricted to a maximum of three, because higher values cannot be reliably estimated and suggests substitution frequencies to be at saturation. For <i>S. alvi</i>, two ternary plots are shown to present comparisons among all four SAGs.</p

Comparative Metabolomics and Structural Characterizations Illuminate Colibactin Pathway-Dependent Small Molecules

The gene cluster responsible for synthesis of the unknown molecule “colibactin” has been identified in mutualistic and pathogenic Escherichia coli. The pathway endows its producer with a long-term persistence phenotype in the human bowel, a probiotic activity used in the treatment of ulcerative colitis, and a carcinogenic activity under host inflammatory conditions. To date, functional small molecules from this pathway have not been reported. Here we implemented a comparative metabolomics and targeted structural network analyses approach to identify a catalog of small molecules dependent on the colibactin pathway from the meningitis isolate E. coli IHE3034 and the probiotic E. coli Nissle 1917. The structures of 10 pathway-dependent small molecules are proposed based on structural characterizations and network relationships. The network will provide a roadmap for the structural and functional elucidation of a variety of other small molecules encoded by the pathway. From the characterized small molecule set, <i>in vitro</i> bacterial growth inhibitory and mammalian CNS receptor antagonist activities are presented

Fiji macro.

The honey bee is a powerful model system to probe host–gut microbiota interactions, and an important pollinator species for natural ecosystems and for agriculture. While bacterial biosensors can provide critical insight into the complex interplay occurring between a host and its associated microbiota, the lack of methods to noninvasively sample the gut content, and the limited genetic tools to engineer symbionts, have so far hindered their development in honey bees. Here, we built a versatile molecular tool kit to genetically modify symbionts and reported for the first time in the honey bee a technique to sample their feces. We reprogrammed the native bee gut bacterium Snodgrassella alvi as a biosensor for IPTG, with engineered cells that stably colonize the gut of honey bees and report exposure to the molecules in a dose-dependent manner through the expression of a fluorescent protein. We showed that fluorescence readout can be measured in the gut tissues or noninvasively in the feces. These tools and techniques will enable rapid building of engineered bacteria to answer fundamental questions in host–gut microbiota research.</div

Testing of different IPTG-inducible constructs in <i>S</i>. <i>alvi</i>.

(a) Maps of the IPTG-inducible plasmids built in this study. (b) S. alvi cells engineered with our inducible plasmids respond to IPTG exposure in vitro. Graph shows box plots representing median value of GFP fluorescence of 5 biological replicates for each construct tested. Each replicate value is based on the average fluorescence of at least 9,000 S. alvi cells measured by flow cytometry, which were grown in liquid with (+) or without (−) IPTG. As a reference, wild-type S. alvi, S. alvi bearing the pAC08 plasmid constitutively expressing GFP, and S. alvi carrying the previously built pBTK552 vector [28] were also analyzed. Fold-changes of average fluorescence between uninduced and induced cells are indicated. The data underlying this Figure can be found in the S1 Data file, sheet “Supplementary Fig 6B.” (PDF)</p

Validation of qPCR primers and standard curves.

Primers specificity was confirmed by visualization of amplicons on an agarose gel (left panel) and generation of melting curves (middle panel). Standard curves were generated using serially diluted genomic DNA of (a) S. alvi, (b) B. apis, or (c) miniprep of pBTK570. The data underlying this Figure can be found in the S1 Data file, sheets “Supplementary Fig 3A,” “Supplementary Fig 3B,” and “Supplementary Fig 3C.” (PDF)</p

Characterization of functional broad-host range replicons in the honey bee gut symbiont <i>B</i>. <i>apis</i>.

(a) Broad-host range plasmids have different copy numbers in B. apis. Box plots show median values of plasmid copy numbers obtained by qPCR from 3 independent experiments with 5 biological replicate each (total n = 15). Median copy number are indicated with the corresponding box plots. (b) The difference in plasmid copy number results in different protein expression levels in B. apis. Graph shows mean of E2-crimson fluorescence ± standard deviations of 5 biological replicates. Each replicate represents the average fluorescence of at least 9,000 cells measured by flow cytometry. Plasmids used for panels a and b in B. apis were pBTK570, pAC06, pAC11, and pAC04, carrying the RSF1010, RK2, pTF-FC2, and pBBR1 origins of replication, respectively. (c) Some replicons are compatible and can be cotransformed in B. apis. Matrix table indicates compatible (green boxes with check mark) and incompatible (red boxes with cross mark) replicons. Vectors were found compatible upon their successful cotransformation by conjugation in B. apis cells. The data underlying this Figure can be found in the S1 Data file, sheets “Supplementary Fig 4A” and “Supplementary Fig 4B.” (PDF)</p