ABSTRACT
The capacity to associate symbiotic bacteria with vital aspects of plant and animal biology is outpacing our understanding of the mechanisms shaping these interactions. A major barrier to mechanistic studies is the paucity of tools for genetically manipulating wild and diverse bacterial isolates. Solving this problem is crucial to elucidating the cellular and molecular rules that govern symbiotic relationships and ultimately harnessing them for agricultural and biomedical applications. Therefore, we constructed a series of vectors that expedite genetic knock-in and knock-out procedures across a range of bacterial lineages. This was accomplished by developing strategies for domestication-free bacterial conjugation, designing plasmids with customizable features, and streamlining allelic exchange using visual markers of homologous recombination. These tools enabled a comparative study based on live imaging of diverse bacterial symbionts native to the zebrafish intestine, with which we discovered heterogeneous colonization patterns and a striking correlation between bacterial population biogeography and cellular behavior.
INTRODUCTION
High-throughput metagenomic sequencing has exposed the previously unseen diversity of symbiotic bacteria that live in close contact with plants and animals throughout the biosphere1,2. Associations are being made at breakneck speed between the membership and activity of resident bacteria and the health, development, and evolution of their hosts3–7. However, the cataloging of symbiotic relationships—whether mutualistic, commensal, or pathogenic—is vastly outpacing their cellular and molecular interrogation8,9. Elucidating the mechanisms by which symbiotic bacteria live and interact with each other and their hosts will inform how they can be harnessed for agricultural and biomedical applications2,10–12.
Characterizing the biology of symbiotic bacteria requires methods for precisely manipulating their genomes. For example, stable chromosomal insertion of genes encoding fluorescent proteins allows cellular behaviors and interactions to be directly observed within native host-associated environments13,14. Additionally, gene deletion and complementation studies are essential for rigorously dissecting the genetic pathways that control specific phenotypes15. Such knock-in and knock-out technologies have long been mainstays in microbiology labs working with entrenched model organisms like E. coli, but established genetic approaches are often inadequate for manipulating wild and novel species or strains16. This is largely because legacy protocols can involve cumbersome and outdated procedures that are difficult to use across lineages. Consequently, the in-depth study of most symbiotic bacteria remains out of reach.
A major bottleneck within the field of symbiosis research is that locating appropriate genetic tools and methods or developing them de novo are arduous and time-consuming tasks. This problem is especially burdensome for investigators aiming to manipulate multiple bacterial lineages derived from complex communities. To overcome these barriers, we have employed the zebrafish intestinal microbiota as a source of wild and diverse symbiotic bacteria17—which includes representatives of the Vibrio, Aeromonas, Pseudomonas, Acinetobacter, Enterobacter, and Plesiomonas genera—to develop and test streamlined tools and methods for bacterial genetic manipulation. We identified three main deficiencies inherent to current genetic approaches that if resolved, will immediately improve the genetic tractability of many bacteria. First, although conjugation is a robust and reliable method for delivering DNA into bacteria, strategies for selecting individual cells carrying the transferred DNA are not broadly compatible between different lineages and sometimes rely on deleterious domestication steps. Second, most vectors used for making genetic manipulations are not readily customized, which restricts their versatility and prevents further innovation. And third, techniques for generating chromosomal modifications via allelic exchange often depend on specific selection conditions that can vary between bacterial lineages and are difficult to troubleshoot when they fail. To address these shortcomings, we rationally designed a centralized set of genetic engineering vectors with new and updated functionalities. For DNA delivery, we developed alternative schemes for post-conjugation counterselection that avoid initial domestication of engineered bacteria, thereby preserving their natural physiology and behavior. For customization, we designed gene expression scaffolds with interchangeable sequence elements that can be tailored to different bacterial genomes, and with these produced a variety of ready-made vectors for fluorescently tagging bacteria. Moreover, an extensive collection of marked zebrafish intestinal symbionts was generated during this work that will accelerate research in the growing zebrafish-microbiota community. Lastly, we devised a means of visually following homologous recombination events during allelic exchange protocols for more tractable generation of markerless chromosomal alterations.
To demonstrate the potential of these modernized tools to uncover new aspects of host-microbe interactions, we examined the colonization patterns of several bacterial symbionts native to the larval zebrafish intestine by light sheet fluorescence microscopy18. The intestinal microbiota is an especially important target for exploration because of its impact on host health and disease; however, its phylogenetic diversity and concealed location make it difficult to investigate in situ by conventional techniques. Unexpectedly, live imaging of bacterial symbiont behavior within larval zebrafish revealed that genome sequences and in vitro-based phenotypes were poor predictors of whether a given bacterium exhibits free- swimming motility in vivo. Most strikingly, we also discovered a general relationship between the growth mode of individual bacteria and their overall biogeography; namely, the average location of a population along the intestinal tract is strongly correlated with the fraction of planktonic cells it contains. In addition to revealing the existence of previously undocumented interactions within a vertebrate intestine, this exploratory experiment underscores how tools for genetically manipulating diverse bacterial symbionts facilitates comparative studies involving multiple species.
In total, the tools and step by step protocols described here will empower a wide range of researchers studying different host-microbe systems as well as free-living bacteria to explore deeper into the inner workings of wild and novel bacterial isolates. Our solutions equally enhance the genetic manipulation of both established and newly emerging model bacterial lineages and will speed the research pipeline from metagenomics to mechanistic microbiology.
RESULTS
The lack of compatible post-conjugation counterselection strategies and the deleterious nature of bacterial domestication
The process of genetically manipulating a bacterium typically begins with the delivery of recombinant DNA into its cell. Several methods can be used for this purpose, but conjugation or bacterial mating—which is the transfer of DNA from a donor cell to a target cell—is highly efficient and versatile, working with a wide range of bacterial lineages. During this procedure, successfully modified target cells are made drug resistant to facilitate their selective recovery. However, a constraint of conjugation is that it also depends on a strategy for simultaneously counterselecting against the donor cells that harbor the same drug resistance. This can be problematic when dealing with novel and uncharacterized bacteria because counterselection schemes often rely on known aspects of a target strain’s physiology. For example, many donor strains (e.g., E. coli SM10) are auxotrophic for amino acids and vitamins and thus, can be counterselected on defined growth media lacking specific metabolites. But for this approach to work the target strain cannot itself be auxotrophic and the right mixture of nutrients and ions must be formulated, which is not always straightforward due to the complex metabolic needs of different bacteria. We found that this drawback represents an immediate hurdle when working with several zebrafish-derived bacterial symbionts. None of the isolates we aimed to genetically manipulate could grow on a standardized defined growth medium (i.e., M9 minimal media) that is used for E. coli SM10 counterselection. This ultimately meant that customized counterselection media would need to be painstakingly developed for each new target strain.
Faced with this scenario, we turned to domestication, which is the process of modifying target strains with a selective trait that is not expressed by the donor. This is commonly done by isolating target strain variants with spontaneous antibiotic resistance. However, despite the wide use of this technique, antibiotic resistances associated with domestication (e.g., to rifampicin or streptomycin) typically arise because of mutations within critical cellular machinery, such as RNA polymerase or components of the ribosome, which can severely impair the natural physiology of bacteria and render them unsuitable for study19–25. Consistent with the deleterious consequences of domestication, we found that a rifampicin-resistant variant of the zebrafish intestinal symbiont Vibrio cholerae ZWU0020, denoted ZWU0020R, exhibits highly perturbed growth kinetics in vitro (Figure 1A). In addition, although domestication of Vibrio ZWU0020 did allow us to successfully insert a gene encoding green fluorescent protein (GFP) within its genome, this modification further aggravated its poor growth phenotype (Figure 1A). As another assessment of ZWU0020R’s altered physiology, we characterized its motility phenotype and observed that it displays attenuated swimming in soft agar compared to wild-type (Figure 1B). To determine if altered swimming is a phenotype of all rifampicin-resistant Vibrio ZWU0020 variants, we inspected the motility phenotype of four independently derived clones. Interestingly, two of the clones are attenuated like the original ZWU0020R strain, whereas the other two perform similar to wild-type, suggesting that they may carry alternative and/or compensatory mutations (Figure 1C). Altogether, our experience attempting to genetically manipulate a variety of novel zebrafish bacterial symbionts using conventional approaches highlights the limitations of current counterselection strategies and the deleterious nature of domestication.
Temperature and kill switch-based systems for domestication-free counterselection of donor cells
To address the lack of adequate post-conjugation counterselection methods, we set out to develop strategies that are technically straightforward and not reliant on inherent or domesticated traits of target strains. We devised two plasmid-based counterselection systems that control donor cell growth by a mechanism similar to that of common suicide vectors. The first system is temperature-based and works through a temperature-sensitive origin of replication that restricts donor cell growth in the presence of antibiotic selection at or above 37°C. Temperature-based control of plasmid replication is well established, but has not been widely implemented as a method of post-conjugation counterselection despite its amenability and previous indications that it can be used in this way26. The second system restricts donor cell growth through a genetic kill switch that, when induced, leads to the expression of three toxic peptides. These two approaches differ in their mode of action and offer slightly different procedural advantages. Notably, we chose to develop plasmid-based counterselection systems because their portability allows the use of alternative donor strains. To initially test the utility of each counterselection system, we incorporated them into existing vectors that are commonly used for making targeted Tn7 transposon-based chromosomal insertions27.
Temperature-based counterselection was achieved by replacing the R6K origin of replication of the Tn7-tagging vector pUC18R6KT-mini-Tn7T-GM (pTW56) with the temperature-sensitive origin of replication ori101/repA101ts 28 (Figure 2A). The resulting vector, pTn7xTS (Temperature-Sensitive), mediates temperature-dependent growth of E. coli SM10 (Figure 2-Figure Supplement 1). At the permissive temperature of 30°C, SM10/pTn7xTS grows normally on rich media in the presence of antibiotic selection. At the restrictive temperature of 37°C, the vector is unable to be maintained, leading to loss of antibiotic resistance and a drop in viability by up to three orders of magnitude (Figure 2-Figure Supplement 1). In the context of an example Tn7-tagging protocol, conjugation is performed at 30°C without antibiotic selection between two SM10 donor strains and a Vibrio target strain (Figure 2B, left). The SM10 donors carry either pTn7xTS (donorTn) or the transposase-encoding helper plasmid pTNS2 (donorhelper). At this point in the procedure, only the donorTn strain is resistant to the selective antibiotic being used, which in this scenario is gentamicin. Successfully modified Vibrio cells harboring a chromosomal copy of the Tn7 transposon, along with the gentamicin resistance gene it encodes, are then selected for by plating the mating mixture in the presence of gentamicin at 37°C (Figure 2B, right). The donorTn strain is counterselected because it is unable to maintain plasmid-based resistance at 37°C, whereas the donorhelper strain remains sensitive to gentamicin throughout the procedure.
A strength of temperature-based counterselection is that it is technically simple, requiring only a shift in growth temperature, but it is limited to target strains that can grow at 37°C. This constraint is problematic for several bacterial lineages native to zebrafish as well as other ectotherms, such as stickleback or fruit flies, which cannot survive at temperatures above the growth temperature of their host (in these cases, ≤ 30°C). Therefore, we developed a second strategy based on an inducible kill switch that functions independently of growth temperature.
The kill switch we designed consists of two elements: a constitutively expressed lacl gene, which encodes the lac repressor, and a synthetic operon containing three E. coli-derived genes encoding toxic peptides—HokB, GhoT, and TisB—placed under the control of the Lacl-repressible promoter Ptac (Figure 2-Figure Supplement 2A)29–33. Upon induction by the allolactose analogue isopropyl-β-D-thiogalactoside (IPTG) these toxic peptides act to disrupt the proton-motive force within donor cells, leading to impaired ATP synthesis and death. We built this kill switch counterselection system into the backbone of the Tn7-tagging vector pUC18T-mini-Tn7T-GM (pTW54), producing pTn7xKS (Kill Switch) (Figure 2C). In the presence of antibiotic selection and IPTG, pTn7xKS is capable of inhibiting SM10 growth by up to four orders of magnitude (Figure 2-Figure Supplement 2B). Of note, initial kill switch prototypes carrying only a single toxin gene were less potent, which may be an important consideration in future kill switch designs (Figure 2-Figure Supplement 2B). In the context of a Tn7-tagging scenario, kill switch-based counterselection is carried out in much the same way as temperature-based counterselection, except that selection of modified target cells is done on media containing IPTG at a growth temperature suitable for the target strain being used (Figure 2D).
Chromosomal insertion of rationally designed gene expression scaffolds into wild and uncharacterized bacterial lineages using domestication-free counterselection systems
To test the effectiveness of our domestication-free counterselection systems, we employed them to integrate genetically encoded fluorescent proteins into the chromosome of various uncharacterized zebrafish bacterial symbionts. However, while exploring available gene expression constructs, we found that many are inflexible and inadequately designed. Specifically, vectors often contain extraneous DNA sequences left over from previous imprecise subcloning procedures and have little to no options for customizing important sequence motifs. The ability to customize expression constructs is critical when working with lineages that differ in, for instance, optimal promoter sequences or ribosome binding sites. Therefore, we first addressed the need for standardized expression constructs by rationally designing a modular gene expression scaffold.
An expression scaffold containing four interchangeable elements—a promoter, 5’ and 3’ untranslated regions (UTR), and an open reading frame (ORF)—was built into the multiple cloning site (mcs) of pGEN-mcs34, producing pXS (eXpression Scaffold) (Figure 3). pGEN-mcs was chosen to house the expression scaffold because it enables fast and easy prototyping of scaffold parts in E. coli, which like many zebrafish bacterial symbionts, is a member of the Gammaproteobacteria and shares basic genetic control elements. Restriction sites underlie the modular architecture of the scaffold and allow each part to be customized (Figure 3-Figure Supplement 1A). Sequence motifs can be replaced individually or the entire scaffold can be subcloned. As initially built, a minimal Ptac promoter without the lac operator sequence, which avoids interference from an endogenously encoded lac repressor if present, is used to achieve constitutive transcription. A synthetic 5’ UTR containing both an epsilon enhancer sequence and ribosome binding site controls translation35,36. The 3’ UTR, which was originally present within pGEN-mcs, contains a trpL attenuator sequence for transcriptional termination37. Lastly, three different ORFs encoding the fluorescent proteins sfGFP38, dTomato39, and mPlum40 were each used to produce three separate expression scaffold variants, pXS-sfGFP, pXS-dTomato, and pXS-mPlum (Figure 3-Figure Supplement 1A). After assembly, each expression scaffold was subcloned into Tn7-tagging vectors with either temperature (i.e., pTn7xTS) or kill switch (i.e., pTn7xKS) counterselection systems (Figure 3-Figure Supplement 1B and C).
Tn7-tagging vectors equipped with rationally designed expression scaffolds were next used to carry out chromosomal tagging of Vibrio ZWU0020 as outlined in Figure 2B and 2D. Unlike previous attempts (Figure 1A), domestication-free manipulation of Vibrio ZWU0020—using either temperature or kill switch counterselection systems—both worked and preserved this strain’s normal physiology (Figure 3-Figure Supplement 2). To demonstrate the compatibility of these tools across different zebrafish symbiont lineages, we tagged 10 strains representing 6 different genera—including Vibrio, Aeromonas, Pseudomonas, Acinetobacter, Enterobacter, and Plesiomonas (Supplementary File 1). Multiple variants that express either sfGFP, dTomato, or mPlum were generated for many of the lineages (Supplementary File 1). This collection of marked zebrafish symbionts serves as a resource of ready-made strains for the zebrafish microbiota community. Notably, several of the lineages manipulated are novel and have yet to be assigned species designations, which is typical of symbionts associated with complex host-associated communities. We also verified that domestication-free counterselection systems can be used to manipulate other species not associated with the zebrafish intestinal microbiota, including a human-derived strain of E. coli (HS)41, the leech symbiont Aeromonas veronii HM2142, and an environmental isolate of Shewanella oneidensis (MR-1)43 (Supplementary File 1).
Streamlining allelic exchange by visualizing homologous recombination events using a fluorescent tracker
Allelic exchange is a robust and versatile homologous recombination technique for making targeted genetic knock-ins and knock-outs in bacteria44–46. Therefore, to extend the utility of our domestication-free counterselection systems, we incorporated them into currently available vectors that are used for mediating allelic exchange. As expected, these updates facilitated the domestication-free engineering of gene deletions in several uncharacterized symbiotic bacteria. However, not all bacteria tested could be successfully manipulated using current allelic exchange protocols, highlighting another breakdown in the compatibility.
Allelic exchange involves two successive homologous recombination events between an allelic exchange vector and the bacterial chromosome. The crux of allelic exchange is isolating rare unmarked mutant cells from large populations of heterozygous intermediates known as merodiploids that arise after the vector integrates into the chromosome during the first recombination step. A longstanding strategy for recovering variants that have undergone the second recombination, which results in vector loss, works by restricting merodiploid growth. This is typically done by expressing a gene called sacB located within the allelic exchange vector backbone that confers growth inhibition in the presence of sucrose47. Although widely used, sacB counterselection of merodiploids does not always work and can be difficult to troubleshoot when it fails. We experienced these shortcomings while attempting to delete a gene associated with chemotactic behavior in a zebrafish symbiont, Vibrio furnissii ZOR0035, using the common sacB-based allelic exchange vector pDMS19748. Vibrio ZOR0035 merodiploids are refractory to sacB counterselection, which made it impossible to isolate cells with the desired mutation. We surmise that the counterselection fails in some bacterial lineages because the expression or activity of the levansucrase enzyme encoded by sacB, which synthesizes high-molecular-weight fructose polymers, is inadequate. To overcome lineage-specific limitations of sacB counterselection, we developed a more tractable strategy based on visual markers.
Our solution uses GFP to track the merodiploid status of target cells (Figure 4A). In this way, the initial recombination step generates GFP-positive merodiploid populations that can be readily screened for cells where the second recombination step has occurred, producing GFP- negative mutants (i.e., instances of “successful” allelic exchange), which typically occur with equal frequency as wild-type revertants (i.e., instances of “aborted” allelic exchange) (Figure 4A). To test the feasibility of this approach, we revisited the engineering of a gene deletion in Vibrio ZOR0035. A constitutively expressed GFP gene was inserted into the backbone of a prototype pDMS197 vector containing a kill switch counterselection system and an allelic exchange cassette targeting the chemotaxis gene cheA. At the time of this work, cheA was the focus of an unrelated project, and it is used here merely to demonstrate proof of concept. As outlined in Figure 4B, the GFP marked allelic exchange vector was delivered into Vibrio ZOR0035 via conjugation. GFP-positive merodiploids, harboring an integrated copy of the allelic exchange vector, were readily isolated and purified. Of note, over the course of this work we empirically determined that the kill switch toxins do not interfere with merodiploid growth in several different bacteria, indicating that either they have restricted activity and are only lethal to E. coli donor cells or they fail to reach toxic levels when expressed from a single chromosomal locus. Next, populations of merodiploids were expanded in liquid culture and plated on nonselective media at a density that allowed discrete colonies to form. Colonies exhibiting sectored regions of GFP loss were then purified to obtain isogenic clones, and putative mutants were genotyped by polymerase chain reaction (PCR). Genotyping was done using PCR primers that flank the cheA locus, yielding a single large amplification product if the cheA gene is present and a smaller sized product if the mutant allele is present. Because they are heterozygous, merodiploids produce both products. Ultimately, our visual merodiploid tracking strategy proved extremely efficient and straightforward to perform, allowing us to successfully engineer a targeted gene deletion in a bacterial strain that was otherwise genetically intractable using previous methodologies.
Gene deletion and complementation with modernized engineering vectors
To complete the genetic toolkit for manipulating wild and diverse bacterial isolates, we combined the tools and approaches described thus far to construct a set of adaptable allelic exchange vectors that further improve the tractability of making markerless genetic alterations. These modernized vectors incorporate fluorescent merodiploid tracking and domestication-free counterselection systems within a highly customizable plasmid architecture (Figure 5, “vector design”). Molecular scaffolds for holding antibiotic selection markers, fluorescent trackers, and a counterselection kill switch were built into a pUC-derived vector backbone that has a temperature-sensitive origin of replication and a single blunt restriction site for straightforward insertion of allelic exchange cassettes. This modular design allows virtually every functional element to be customized for different bacterial lineages (Figure 5-Figure Supplement 1).
In total, two allelic exchange vectors were generated, pAX1 and pAX2 (Allelic eXchange), which differ in their domestication-free counterselection systems (Figure 5). Both vectors mediate temperature-based counterselection of SM10 donor cells, but pAX2 also contains a TetR-regulated kill switch that can be induced by anhydrotetracycline (Figure 5-Figure Supplement 2). Notably, the dual temperature/kill switch counterselection activity of pAX2 is quite potent, reducing SM10 viability by over five orders of magnitude (Figure 5-Figure Supplement 2B). Two resistance markers for gentamicin and chloramphenicol were included to give pAX1 and pAX2 greater “off the shelf” compatibility with different target strains.
These new allelic exchange vectors were next used to engineer markerless gene deletions. For this proof of concept, we designed an allelic exchange cassette to delete two neighboring genes in Vibrio ZWU0020, pomA and pomB, which encode components of the polar flagellar motor. After inserting the cassette into an early, but functionally equivalent, version of pAX1 (see Materials and Methods), GFP-positive merodiploids were generated and isolated as before and screened for loss of GFP expression and thus, the integrated vector (Figure 6-Figure Supplement 1). Mutants harboring the desired mutation, which fused the start codon of pomA with the stop codon of pomB, were confirmed by PCR (Figure 6A and 6B). As anticipated, ZWU0020 ΔpomAB exhibited complete loss of swimming motility in soft agar (Figure 6C, bottom left) without overt growth defects in liquid media (Figure 6-Figure Supplement 1). To demonstrate the cross-lineage compatibility of these tools, we successfully employed pAX2 to create a similar markerless deletion of a homologous pomAB locus in another zebrafish symbiont, Aeromonas veronii ZOR0001 (Figure 6-Figure Supplement 2). Notably, while screening Vibrio ZWU0020 and Aeromonas ZOR0001 merodiploid colonies for putative mutants, we observed that many different patterns of GFP loss can arise (Figure 6- Figure Supplement 3). Remarkably, even in situations where mutant cells reside within miniscule GFP-negative patches, a single colony purification step can often be used to isolate them. The ability to readily identify and recover mutant cells with such sensitivity highlights the robustness of our visual screening approach.
Constructing deletion mutants is just the first step in dissecting the genetic pathways that underlie a given activity or behavior. Complementation must also be performed to rigorously confirm that a mutant phenotype is the result of a specific genetic disruption and not polar effects or other unintended consequences of chromosomal manipulation15. Therefore, we wanted to demonstrate how our domestication-free Tn7-tagging vectors could be employed to complement the ZWU0020 ΔpomAB mutant. The pomAB locus of ZWU0020, including the native pomA promoter, was PCR-amplified and inserted within the Tn7 transposon of the tagging vector pTn7xTS-sfGFP, which also contains a constitutively expressed sfGFP (Figure 6D). Chromosomal insertion of this construct at the attTn7 site of ZWU0020 ΔpomAB fully restores wild-type motility, thus confirming that sole disruption of these genes caused the loss of motility in the mutant (Figure 6C, bottom right).
Live imaging of diverse bacterial symbionts yields insights into host-microbe interactions within the vertebrate intestine
The power to genetically manipulate a diverse range of bacterial symbionts provides an opportunity for comparative studies focused on identifying unique or broadly conserved features of host–microbe interactions. To illustrate how the tools developed in this work facilitate such investigations, we examined the intestinal colonization patterns and cellular behaviors of seven fluorescently tagged zebrafish symbionts by light sheet fluorescence microscopy13,49. The strains chosen included: Aeromonas ZOR0001, Aeromonas ZOR0002, Enterobacter ZOR0014, Plesiomonas ZOR0011, Pseudomonas ZWU0006, Vibrio ZOR0036, and Vibrio ZWU0020. A tagged version of the ZWU0020 ΔpomAB motility mutant was also analyzed. Prior to imaging, bacteria were associated individually with 4-day old, germ-free larval zebrafish for 24 hours. Mono-association provides unrestricted access to the intestinal environment free of competition with resident microbiota, and therefore allows interactions between a single symbiont and its host to be studied in isolation. For each strain, real-time two-dimensional movies and three-dimensional images spanning the entire volume of the larval intestine were acquired from three separate hosts (Figure 7, Figure 7-Figure Supplement 1, and Supplementary Movies 1-8). From these data, distinct population structures are readily identified, and can be summarized by three properties: cell motility, growth mode (i.e., planktonic vs aggregated), and biogeography. Additional features of each strain, including estimated abundance, are provided in Figure 7-Figure Supplement 2.
Four of the seven wild strains examined display active motility within the intestine. Surprisingly, we found several discrepancies between the motility phenotype of strains in vivo and their predicted capacity for motility based on phylogenetic relatedness, genome sequence, and performance during in vitro assays (Figure 7-Figure Supplement 3 and Supplementary Movies 1-7). Each strain carries genes for flagellum biogenesis and displays free-swimming motility in liquid media. In addition, all strains were found to swim in soft agar, with the exception of Vibrio ZOR0036 (Figure 7-Figure Supplement 4). Yet despite these attributes, motile individuals were not observed within intestinal populations of Enterobacter ZOR0014, Aeromonas ZOR0001, and Aeromonas ZOR0002 (Supplementary Movies 2-4). We note that in a previous live imaging study involving Aeromonas ZOR0001 motile cells were detected, but that their occurrence was incredibly rare13. In addition, although Vibrio ZOR0036 exhibits little motility in soft agar, it gives rise to a considerable number of highly motile cells in vivo (Supplementary Movie 5); however, its overall motility phenotype is muted compared to closely related Vibrio ZWU0020 (Figure 7-Figure Supplement 3 and 4, and Supplementary Movie 1).
The growth mode of cells is categorized as either planktonic, which includes both motile and non-motile individuals, or aggregated. Both modes are typically represented across different populations, but the ratio of cells in each mode tends to be strain-specific. For example, at one extreme, populations of Vibrio ZWU0020 are almost entirely comprised of highly motile planktonic cells; this is more apparent in movies than three-dimensional image scans because of this strain’s high population density and fast movement (Supplementary Movie 1). At the opposite extreme, Enterobacter ZOR0014 forms large multicellular aggregates without motile individuals (Figure 7C and Supplementary Movie 2). The remaining strains produce populations with intermediate mixtures of planktonic and aggregated cells (Figure 7C and Figure 7-Figure Supplement 1).
Regarding biogeography, we observed a range of strain-specific spatial distributions along the length of the intestine. We can coarsely classify the location of bacterial populations as primarily residing in one of two regions, the proximal gut (referred to as the “bulb”) or the mid-gut, which we approximate in Figure 7B based on previous studies of larval zebrafish intestinal development50,51. For most strains, the bulk of their population is distributed throughout the mid-gut (e.g., Aeromonas ZOR0001, Aeromonas ZOR0002, Enterobacter ZOR0014, Plesiomonas ZOR0011, Pseudomonas ZWU0006, and Vibrio ZOR0036) (Figure 7C). By contrast, populations of Vibrio ZWU0020 are located within the proximal portion of the bulb, consistent with previous findings (Figure 7C)13. Notably, compared to wild-type, the ZWU0020 ΔpomAB motility mutant exhibits a reduction in overall population size and a slight shift in distribution to an area between the bulb and mid-gut (Figure 7C), indicating that for this strain, motility controls abundance and biogeography.
To distill and quantify our observations, we devised scalar metrics of biogeography and growth mode using computational image analysis (Materials and Methods). For each population, we computed the center of mass along the anterior-posterior axis of the intestine to represent biogeography and enumerated the fraction of cells that exist as planktonic individuals to represent growth mode. Plotting these data against each other shows a striking and unanticipated relationship: the fraction of planktonic cells within a strain’s population is strongly correlated with its biogeography (Figure 8). We quantified this relationship by linear regression of log-transformed planktonic fraction and population center medians, excluding the ZWU0020 ΔpomAB motility mutant (Figure 8, dashed trend line; r2 = 0.6762). Remarkably, the ZWU0020 ΔpomAB motility mutant conforms to this trend (Figure 8). As expected, populations of ZWU0020 ΔpomAB contain no motile individuals, but unexpectedly they adopt a more aggregated state compared to wild-type ZWU0020 (Supplementary Movie 8). This change in growth mode and the concomitant change in biogeography moves the ZWU0020 motility mutant along the trend line.
DISCUSSION
Impact on Bacterial Symbioses Research
Over the last decade, the landscape of bacterial symbiosis research has shifted due to an explosion in omics technologies and large-scale initiatives like the Human and Earth Microbiome Projects52,53. The traditionally static “one host, one microbe” view has given way to one that is more dynamic and complex, taking into account the highly contextual nature of symbiotic relationships and involvement of diverse multi-member microbial communities. This paradigm shift has generated several new challenges; chiefly, the demand for more efficient genetic manipulation of wild and novel symbiotic bacterial lineages. Addressing this problem is critical to experimentally unravelling the cellular and molecular determinants of host-microbe systems.
The impetus behind the genetic tools and approaches described in this work emerged from setbacks encountered while attempting to manipulate members of the zebrafish intestinal microbiota. The diversity of species and strains exposed several key inadequacies and weaknesses in conventional techniques, including the lack of domestication-free strategies for donor cell counterselection, poor modularity of available tools, and intractability of allelic exchange. Therefore, we designed tools and methods to circumvent these points of failure and quickly adapt to unforeseen idiosyncrasies of species or strains. In this way, an individual researcher or laboratory can focus on a single operating procedure using a centralized set of tools while being empowered to innovate when needed.
Utility of the Tools Produced by this Work
An extensive collection of molecular tools is available for genetically manipulating bacteria; however, many can only be used with a small number of species or strains. The siloed nature of genetic tools puts a significant burden on researchers looking to manipulate diverse bacterial lineages because it forces them to sift through and become familiar with numerous different vectors and protocols. Furthermore, for those working with novel and uncharacterized bacteria or new to performing genetic manipulations altogether, developing a molecular starter kit is overwhelming. We addressed these problems by constructing a set of standardized engineering vectors that streamline the process of making genetic knock-ins and knock-outs across different lineages. These tools are briefly summarized in Figure 9 and their features are discussed below.
Minimizing laboratory-based domestication of wild bacterial isolates
The deleterious nature of domestication is well-documented, yet it is often overlooked and domestication steps are unfortunately routinely performed because of their convenience23,24. Domestication is commonly used to improve genetic tractability or to help discern specific bacterial strains from other lineages within complex environments (e.g., the vertebrate intestine, water, or soil). Compensatory mutations can rescue or mask physiological defects associated with domestication in vitro25; however, there is no guarantee that critical aspects of symbiont biology, such as those involved in host engagement, are left unperturbed. Therefore, the accurate modeling of symbiotic interactions requires careful attention to preserving natural symbiont behaviors. The incorporation of temperature and kill switch-based domestication-free counterselection systems into a previously described Tn7 tagging vector27, which produced pTn7xTS and pTn7xKS (Figure 2), and the novel allelic exchange vectors pAX1 and pAX2 developed in this work (Figure 5), offers ready-made tools for manipulating various symbiotic bacterial lineages in a way that preserves their natural physiology.
Achieving broad utility through modularity
The incredible genetic and phenotypic diversity of bacteria challenges cross-lineage compatibility of genetic tools. A major contributing factor to this problem is that many available tools are irreversibly constructed, which impedes the customization of important sequence motifs for different bacteria. We addressed this by building tools with highly modular architectures so that they can be easily reconfigured and are thus molecularly nimble. This feature makes it possible to continuously innovate and build off original designs. For example, expanding on the expression scaffolds within the pXS series of vectors (Figure 3 and Figure 3-Figure Supplement 1), we have engineered more elaborate genetic devices, including reporters of gene expression and genetic switches. Additionally, the modularity of these vectors can be advantageous in situations where rational design is not possible due to the unavailability of suitable sequence elements; libraries in which constructs containing a single variable motif (e.g., a promoter or ribosome binding site) can be readily assembled and screened for optimal activity. While the flexibility of our expression scaffolds is conducive to further engineering, as built they have immediate utility for stably tagging bacteria and thus, facilitating the direct observation of symbionts within their natural host-associated environments. Illustrating this point, the mere fluorescent tagging of the zebrafish symbionts Vibrio ZWU0020 and Aeromonas ZOR0001 using these tools revealed that each bacterium interacts with the physically dynamic confines of the intestine in distinct ways and that this differential interplay unexpectedly shapes their apparent competition13. We also incorporated modularity into the design of the allelic exchange vectors pAX1 and pAX2 (Figure 5 and Figure 5-Figure Supplement 1). Several elements within these vectors can be customized, including the antibiotic resistance genes, the fluorescent merodiploid tracker, and the kill switch. With this modularity, we are exploring alternative utilities of these vectors. For example, swapping the GFP merodiploid tracker for one that encodes a red fluorescent protein has allowed us to engineer GFP fusions at endogenous chromosomal loci. In total, tailorable tools such as those described here offer a way of tuning or customizing functionality to increase the experimental potential of bacterial lineages.
Streamlining genetic manipulations with visual screening
To improve the tractability of allelic exchange we used GFP to visually track recombination events (Figure 4). This simple update proved extremely powerful. It allowed merodiploids to be confidently identified and isolated while final mutant derivatives could be screened for with incredible sensitivity, sometimes being found as small subpopulations within merodiploid colonies growing on an agar plate (Figure 4 and Figure 6-Figure Supplements 1-3). The successful manipulation of a previously intractable bacterium (i.e., Vibrio ZOR0035) highlights the utility of this approach. Although conventional selection schemes (e.q., based on sacB) are adept at recovering mutants that arise at low frequencies due to rarely occurring recombination events, their use is contingent on specific conditions that can be difficult to translate between species or strains. By contrast, our visual screening approach operates freely across lineages and differs from selections in that it allows for the progression of recombination events to be more precisely monitored. As a result, the engineering and isolation of bacterial mutants is more efficient and attainable.
Demonstrating New Avenues for Research: Comparative Study of Bacterial Biogeography in the Zebrafish Gut
The technical flexibility of the tools and methods developed in this work makes it easier to genetically manipulate many different bacterial lineages in parallel. This functionality greatly facilitates comparative studies aiming to disentangle unique and widely conserved aspects of host-microbe systems. Such investigations are important because many properties of complex host-associated microbial communities, like those comprising human microbiota54,55, are extremely variable and remain unexplained. Illustrating the potential for comparative approaches, we exposed several uncharacterized phenomena by probing the colonization patterns and behaviors of multiple bacterial lineages native to the zebrafish intestine.
Upon initial examination, we found discordance between predicted and observed motility phenotypes in vivo. All strains examined carry flagellar genes and tend to be highly motile in vitro, yet several of them (e.g., Enterobacter ZOR0014, Aeromonas ZOR000l, and Aeromonas ZOR0002) display no obvious motility during intestinal colonization (Supplementary Movies 2-4). By contrast, some strains robustly sustain motility in vivo, most notably Vibrio ZWU0020, which appears to produce populations almost entirely made up of swimming cells (Supplementary Movie 1). There are several possible explanations for these behaviors that are not necessarily mutually exclusive. For example, they may be the result of bacteria dynamically responding to the intestinal environment and executing strain or lineage-specific colonization strategies. On the other hand, bacteria could be differentially susceptible to some form of host-mediated motility interference, which has recently been documented in mouse models56,57. Work focused on distinguishing these interactions is ongoing. Ultimately, this observation highlights the disconnect that can occur among in silico, in vitro, and in vivo approaches for studying bacterial symbioses. Considering how to best capture and interpret mechanistic insights across model systems will be critical to progress.
Our data additionally indicate that some strains display a relatively large amount of variation in growth mode and biogeography between hosts (e.g., Aeromonas ZOR000l, Aeromonas ZOR0002, and Enterobacter ZOR0014), while others do not (e.g., Plesiomonas ZOR0011, Pseudomonas ZWU0006, and Vibrio ZWU0020) (Figure 8). This variance is consistent with earlier reports showing that the structure of bacterial populations within the larval zebrafish gut can be highly dynamic, which is attributable in part to the physical forces of intestinal peristalsis13,49. Yet some bacteria—for example, Vibrio ZWU0020—remain stable in the face of this perturbation through still undefined mechanisms13. Our observations with the Vibrio ZWU0020 ΔpomAB mutant suggest cell motility is involved.
Most strikingly, we discovered a strong correlation between the dominant growth mode of bacterial populations and their biogeography (Figure 8). While cell behavior is recognized to influence local population structure58,59, linking cell aggregation with a global pattern of spatial organization throughout the intestine is unanticipated and profound. A possible explanation for this pattern is that physical properties of the intestinal environment (e.g., its shape and/or peristaltic movement) act to spatially segregate planktonic and aggregated cells. Alternatively, bacteria may toggle between different growth modes in response to spatial cues generated by physiologically distinct regions along the length of the intestine. Going forward, a major objective will be to dissect the potential mechanisms of this relationship and importantly, understand how it generalizes both within and across host–microbe systems.
Outlook
Elucidating the rules that govern the assembly and function of bacterial symbioses requires studying a wide range of bacterial symbiont lineages1,2. Whether abundant, rare, divergent, or closely related, all potentially hold clues to how host-microbe systems work and how they can be exploited for biotechnology applications—from boosting food production to treating human disease. As more symbiotic relationships are uncovered and new model systems come online, the continued design and modernization of genetic approaches for streamlined manipulation of diverse bacterial lineages will be paramount. Although symbiotic bacteria were the primary subject for tool development in this work, the approaches we have described are equally applicable to the study of free-living environmental bacteria. Importantly, the growing appreciation for the ubiquity of bacterial symbioses and their far-reaching influence on the lives of plants and animals is inspiring a highly cross-disciplinary generation of microbiologists with mixed and varied backgrounds. Therefore, it will be beneficial to work toward standardized tools and methods that foster the rigorous and accurate investigation of symbiont biology.
MATERIALS AND METHODS
Animal care
All experiments with zebrafish were done in accordance with protocols approved by the University of Oregon Institutional Animal Care and Use Committee and following standard protocols60.
Gnotobiology
Wild-type (AB × TU strain) zebrafish were derived germ-free (GF) and colonized with bacterial strains as previously described with slight modification61. Briefly, fertilized eggs from adult mating pairs were harvested and incubated in sterile embryo media (EM) containing ampicillin (100μg/ml), gentamicin (10μg/ml), amphotericin B (250ng/ml), tetracycline (1μg/ml), and chloramphenicol (1μg/ml) for ~6h. Embryos were then washed in EM containing 0.1% polyvinylpyrrolidone–iodine followed by EM containing 0.003% sodium hypochlorite. Sterilized embryos were distributed into T25 tissue culture flasks containing 15ml sterile EM at a density of one embryo per ml and incubated at 28-30°C prior to bacterial colonization. Embryos were sustained on yolk-derived nutrients and not fed during experiments. For bacterial mono-association, bacterial strains were grown overnight in LB liquid media with shaking at 30°C, and prepared for inoculation by pelleting 1ml of culture for 2 min at 7,000 × g, and washing once in sterile EM. Bacterial strains were individually added to the water column of single flasks containing 4-day old larval zebrafish at a final density of ~106 bacteria/ml. Bacterial colonization patterns were assessed 24h later by live imaging of three separate 5-day old zebrafish hosts per bacterial strain. Three animals were determined to be adequate for capturing general colonization features of each bacterial strain based on at least two previous independent qualitative assessments of colonization patterns.
Bacterial strains and culture
All wild and recombinant bacterial strains used or created in this study are listed in Supplementary File 1. Archived stocks of bacteria are maintained in 25% glycerol at −80°C. Prior to manipulations or experiments, bacteria were directly inoculated into 5ml Luria-Bertani (LB) media (10g/L NaCl, 5g/L yeast extract, 12g/L tryptone, 1g/L glucose) and grown for ~16h (overnight) shaking at 30°C, except for E. coli HS, which was grown at 37°C. For growth on solid media, tryptic soy agar was used. 10μg/ml gentamicin was used to select recombinant strains tagged with the Tn7 transposon, which was modified to carry a gentamicin resistance gene. When selecting merodiploid intermediates made using pAX1 or pAX2, which carry resistance to both gentamicin and chloramphenicol, either 10μg/ml gentamicin or 5μg/ml chloramphenicol was used. Selection of rifampicin-domesticated variants was done using 100μg/ml rifampicin.
Molecular techniques and reagents
E. coli strains used for molecular cloning and conjugation, and plasmids used or created during this work are listed in Supplementary File 2. E. coli were typically grown in 5ml LB liquid media at 30°C or 37°C with shaking in the presence of appropriate antibiotic selection to maintain plasmids. For propagation on solid media, LB agar was used. Antibiotic concentrations used were as follows: 100μg/ml ampicillin, 20μg/ml chloramphenicol, 10μg/ml gentamicin, and 10μg/ml tetracycline. Supplementary File 3 lists all DNA primers used for polymerase chain reactions (PCR), which are organized based on their “Wiles Primer” (WP) number. Unless specifically noted, standard molecular techniques were applied and reagents were used according to manufacturer instructions. Restriction enzymes and other molecular biology reagents for PCR and nucleic acid modifications were obtained from New England BioLabs (Ipswich, MA). Various kits for plasmid and PCR amplicon purification were obtained from Zymo Research (Irvine, CA). The Wizard Genomic DNA Purification Kit (Promega, Madison, WI) was used for isolating bacterial genomic DNA. DNA oligonucleotides for PCR were synthesized by Integrated DNA Technologies (Coralville, IA). Sanger sequencing was done by Sequetech (Mountain View, CA). Custom gene synthesis was done by GenScript (Piscataway, NJ). A Leica MZ10 F fluorescence stereomicroscope with 1.0x, 1.6x, and 2.0x objectives and Leica DFC365 FX camera were used for screening and imaging fluorescent bacterial colonies (Leica, Wetzlar, Germany). Images were captured and processed using standard Leica Application Suite software and ImageJ62. Nucleotide sequences of 16S rRNA genes used for phylogenetic analysis are provided in Supplementary File 4, and were obtained via “The Integrated Microbial Genomes & Microbiome Samples” (IMG/M) website (https://img.jgi.doe.gov/m/)63 or the RNAmmer web tool64. 16S rRNA sequences were aligned using Clustal Omega65 and an unrooted phylogenetic tree was drawn using the Phylodendron web tool (http://iubio.bio.indiana.edu:7131/treeapp/treeprint-form.html).
Plasmid construction
The plasmid-based tools developed in this work have been deposited at Addgene (Cambridge, MA), along with their sequences (https://www.addgene.org/). Supplementary File 5 contains annotated nucleotide sequences of select genetic parts that were used to build plasmids and gene expression scaffolds. Details on how plasmids were specifically constructed are provided in Supplementary File 6.
Domestication-free Tn7 tagging using pTn7xTS and pTn7xKS
A detailed Tn7 tagging protocol based on pTn7xTS and pTn7xKS—which includes optimization and troubleshooting steps, and notes on strain-specific procedures—is provided in Supplementary File 7. Generally, and as outlined in Figure 2, triparental conjugation was performed between a single target bacterial strain, an E. coli SM10 donor strain carrying the transposase-containing pTNS2 helper plasmid, and an E. coli SM10 donor strain carrying either a pTn7xTS or pTn7xKS domestication-free tagging vector. Prior to mating, bacteria were prepared by subculturing them to an approximate optical density of 0.4-0.6 at 600nm in LB media with required antibiotics and at the appropriate growth temperature. Cells were then combined 1:1:1 (500μl each), washed once by centrifugation and aspiration in 1ml LB media or 0. 7% NaCl, and suspended in a final 25μl volume of the same media used for washing. Next, the concentrated mating mixture was transferred to a 25mm-wide 0.45μm filter disc (EMD Millipore, Billerica MA; product #HAWP02500) that had been placed on top of a TSA plate. Once the mating mixture dried, the plate was incubated at 30°C for 3-5h. After incubation, the filter disc was placed in 1ml 0.7% NaCl within a 50ml conical tube and bacteria were dislodged by vortexing and pipetting. In cases where a pTn7xTS-based vector was used, 100μl of the bacterial suspension was spread onto a TSA plate containing gentamicin and incubated overnight at 37°C to select for recombinants. To ensure the recovery of low frequency recombinants, the remaining 900μl of the suspension was pelleted by centrifugation, suspended in 100μl 0.7% NaCl, and plated in the same way. In cases where a pTn7xKS-based vector was used, 100μl of the bacterial suspension was spread onto a TSA plate containing gentamicin and 1mM isopropyl-β-D-thiogalactoside (IPTG), and incubated overnight at 30°C. The remaining 900μl was prepared as above, plated on TSA with gentamicin and IPTG, and incubated at 30°C.
The following day, putative recombinant target bacteria were colony-purified by streaking on TSA without antibiotic selection at 30°C. Of note, when recombinant bacteria are tagged with a gene encoding a fluorescent protein, performing colony-purification in the absence of antibiotic selection followed by visual screening of fluorescence is a convenient way to verify that the Tn7 transposon has chromosomally integrated and the tagging vector has been lost. Purified clones were picked, cultured in LB media containing gentamicin, and genotyped by PCR to verify correct insertion of the Tn7 transposon at the attTn7 site. The universal primer WP11, which anneals within the Tn7R region of the Tn7 transposon, was used with a species-specific primer that anneals to an adjacent chromosomal sequence within the 3’ end of the glmS gene to generate a small (~250bp) amplicon if the transposon is present. Species-specific primers used were as follows: Vibrio Z0R0018, WP50; Vibrio Z0R0035, WP51; Vibrio Z0R0036, WP12; Vibrio ZWU0020, WP12; Aeromonas Z0R0001, WP49; Aeromonas Z0R0002, WP52; Pseudomonas ZWU0006, WP256; Acinetobacter Z0R0008, WP259; Enterobacter Z0R0014, WP257; Plesiomonas Z0R0011, WP260; E. coli HS, WP150; Aeromonas HM21, WP49; Shewanella MR-1, WP48.
Generation of markerless deletions via allelic exchange using pAX1 and pAX2
A detailed protocol for carrying out allelic exchange using pAX1 and pAX2—which includes optimization and troubleshooting steps, and notes on strain-specific procedures—is provided in Supplementary File 8. Briefly, and as summarized in Figure 4, allelic exchange cassettes for mediating markerless deletion of target genetic loci (i.e., the pomAB locus of Vibrio ZWU0020 and Aeromonas ZOR0001) were generated through splice by overlap extension and inserted into a pAX-based vector. Next, diparental conjugation was performed between a single target bacterial strain (i.e., Vibrio ZWU0020 or Aeromonas ZOR0001) and an E. coli SM10 donor strain carrying the assembled allelic exchange vector. Prior to mating, bacteria were prepared by subculturing them to an approximate optical density of 0.4-0.6 at 600nm in LB media with required antibiotics and at the appropriate growth temperature. Cells were then combined 1:1 (750μl each), washed once by centrifugation and aspiration in 1ml LB media or 0.7% NaCl, and suspended in a final 25μl volume of the same media used for washing. Next, the concentrated mating mixture was transferred to a 25mm-wide 0.45μm filter disc that had been placed on top of a TSA plate. Once the mating mixture dried, the plate was incubated at 30°C for 3-5h. After incubation, the filter disc was placed in 1ml 0.7% NaCl within a 50ml conical tube and bacteria were dislodged by vortexing and pipetting. For the generation of Vibrio ZWU0020 ΔpomAB, which employed a pAX1-related vector, 100μl of the bacterial suspension was spread onto a TSA plate containing gentamicin and incubated overnight at 37°C to select for merodiploids. The remaining 900μl of the suspension was pelleted by centrifugation, suspended in 100μl 0.7% NaCl, and plated in the same way to ensure recovery of rare recombinants. For the generation of Aeromonas ZOR0001 ΔpomAB, which employed a pAX2-based vector, 100μl of the bacterial suspension was spread onto a TSA plate containing gentamicin and 10ng/ml anhydrotetracycline (aTc), and incubated overnight at 30°C. The remaining 900μl was prepared as above, plated on TSA with gentamicin and aTc, and incubated at 30°C.
The following day, colonies of putative merodiploid target bacteria that were expressing the GFP tracker were purified by streaking on TSA without antibiotic selection at 30°C. This purification step also served to verify that the allelic exchange vector had integrated into the chromosome. Purified clones were picked, cultured in LB media containing gentamicin to maintain their merodiploid state, and archived as a frozen stock. To screen for second recombination events, merodiploids were cultured overnight in LB media without antibiotic selection and spread onto several TSA plates, again without antibiotic selection, at a density that allowed ~100-200 discrete colonies to form. Colonies exhibiting partial or complete loss of GFP expression were purified by streaking on TSA at 30°C. Putative mutants were screened and genotyped by PCR using primers that flanked the modified locus, which produced two differently sized amplicons that represented the wild-type and mutant alleles. Primers WP163 and WP164 were used to genotype Vibrio ZWU0020 ΔpomAB mutants and primers WP192 and WP195 were used to genotype Aeromonas Z0R0001 ΔpomAB mutants.
In vitro growth measurements
In vitro growth of bacterial strains was assessed using the FLUOstar Omega microplate reader (BMG LABTECH, Offenburg, Germany). Prior to growth measurements, bacteria were grown overnight in 5ml LB media at 30°C with shaking. Cultures were diluted 1:100 into fresh LB media and dispensed in quadruplicate (i.e., four technical replicates) (200μl/ well) into a sterile 96 well clear flat bottom tissue culture-treated microplate (Corning, Corning, NY; product #3585). Absorbance measurements at 600nm were then recorded every 30 minutes for 16 hours (or until stationary phase) at 30°C with shaking. Growth measurements were repeated at least two independent times for each strain (i.e., two biological replicates) with consistent results. Data were exported and graphed using GraphPad Prism 6 software.
Swim motility assays
Prior to the assessment of swimming motility, bacteria were grown overnight in 5ml LB media at 30°C with shaking. 1ml of bacterial culture was then washed by centrifuging cells at 7,000xg for 2 minutes, aspirating media, and suspending in 1ml 0.7% NaCl. This centrifugation/ aspiration wash step was repeated once more and bacteria were suspended in a final volume of 1ml 0.7% NaCl. 1μl of washed bacterial culture was then inoculated into a TSA plate containing 0.2% agar (30g/L tryptic soy broth and 2g/L bacto agar). Swim plates were incubated at 30°C for 5-7h and imaged on a Gel Doc XR+ Imaging System (Bio-Rad, Hercules, CA). Motility assays were repeated at least two independent times (i.e., two biological replicates) with consistent results.
Spot tests
E. coli SM10 donor cells carrying vectors that contain temperature and/or kill switch-based post-conjugation counterselection systems were grown overnight in LB media with required antibiotics and at the appropriate growth temperatures. For assessing temperature-based counterselection, ten-fold serial dilutions were made on TSA plates containing gentamicin and incubated overnight at 30°C or 37°C. For assessing kill switch-based counterselection, ten-fold serial dilutions were made on TSA plates containing gentamicin +/− 1mM IPTG (in the case of pTn7xKS) or 10ng/ml aTc (in the case of pAX2) and incubated overnight at 30°C. Plates were imaged on a Bio-Rad Gel Doc XR+ Imaging System. All spot tests were performed at least two independent times (i.e., two biological replicates), each including at least two technical replicates, with consistent results.
Live Imaging
Live larval zebrafish were imaged using a home-built light sheet fluorescence microscope described in detail elsewhere49,66. In brief, a thin sheet of laser light is obtained by rapidly scanning the excitation beam with a galvanometer mirror. Fluorescence emission is captured by an objective lens mounted perpendicular to the sheet. 3D images are obtained by translating the sample along the detection axis. The entire volume of the intestine (approximately 1200x300x150 microns) is imaged in four sub-regions that are computationally registered after acquisition. Total acquisition time of a single intestine is less than 1 min with 1-micron steps between planes. For all images, the exposure time was 30ms and the excitation laser power was 5mW prior to entering the imaging chamber.
Image Analysis
Bacterial abundances and locations were estimated using the analysis pipeline described in49. In brief, we identify individual cells in 3D using a wavelet filtering-based algorithm67 and identify multicellular aggregates using a graph-cut segmentation algorithm68. The number of cells in an aggregate is estimated by dividing the total aggregate intensity by the average intensity of individual bacteria. Individuals detected within an aggregate are discarded. One-dimensional population distributions are obtained by dividing the intestine into 5-micron bins constructed down the length of the intestine along a manually drawn line and assigning the centroid of each detected object to a bin. Global population centers are computed as the center of mass of this 1D distribution. Of note, this analysis pipeline was originally developed and optimized for a different strain not imaged here49 and its performance on the 8 present strains has not been rigorously assessed. Based on manual inspection and analysis, we estimate an uncertainty of at most 10% for the planktonic fraction and for the population center of mass, which is certainly more than adequate to detect the global trends we report.
COMPETING INTERESTS
The authors declare that the tools and methods described in this work are the subject of an ongoing patent application.
ACKNOWLEDGEMENTS
We would like to thank numerous members of the Guillemin lab for their willingness to test and provide feedback on the genetic tools developed in this work, particularly Dr. Annah Rolig and Dr. Cathy Robinson. We would also like to thank Dr. Andrew Camilli for his generosity in sharing overlap extension and allelic exchange protocols. This project was supported through the M.J. Murdock Charitable Trust and an award from the Kavli Microbiome Ideas Challenge, a project led by the American Society for Microbiology in partnership with the American Chemical Society and the American Physical Society and supported by The Kavli Foundation. Work was also supported by the National Science Foundation under Awards 0922951 and 1507115. Authors received funding from the National Institutes of Health (NIH, http://www.nih.gov/), P50GM09891 to KG, F32AI112094 to TJW, and T32GM007759 to BHS. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
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