The author thanks Roberto Kolter (Harvard Medical School) for his invaluable advice and assistance during the development of this coculture screen. She also thanks Matthew Powers for reading the manuscript for clarity, and Chia-yi Cheng for assistance with obtaining Figure 6.

1. Select a Reporter Gene and Construct a Fluorescent Transcriptional Reporter
For B. subtilis:
<ol>
	<li>See the JoVE article in reference 19 for a protocol describing the construction of fluorescent transcriptional reporters in Bacillus subtilis.</li>
</ol>
For other bacterial species:
<ol>
	<li>Identify a gene that is upregulated during the physiological response of interest. This can be based on existing literature or transcriptional analysis of the microbe under particular conditions.</li>
	<li>Construct a fluorescent transcriptional reporter for this gene to act as a proxy for the change in phenotype. This construct should include the promoter of this upregulated gene driving the production of an appropriate fluorescent protein (see Figure 1).</li>
	<li>Integrate this construct into a neutral locus on the chromosome. This ensures that native regulation of the gene of interest is not disrupted, and avoids the need for plasmid selection mechanisms (e.g. antibiotics) that might interfere with the growth of environmental microbes.</li>
</ol>
2. Determine Coculture Conditions
For B. subtilis PtapA-yfp reporter:
<ol>
	<li>Use 0.1x LB, Lennox (1 g tryptone, 0.5 g yeast extract, 0.5 g NaCl per liter) medium for this reporter, since B. subtilis matrix-production is minimal on Luria Broth20. This medium allows B. subtilis colonies to grow to submillimeter but observable size, while allowing diverse taxa from soil to grow5.</li>
	<li>Include 100 mM MOPS buffer to minimize potential pH changes.</li>
</ol>
For other bacterial species:
<ol>
	<li>Use published transcriptional data or empirically test various culture conditions to identify one where the microbe grows but the activation of the fluorescent reporter is negligible (to allow its activation to be detected when the reporter strain is grown in coculture with inducing microbes.)</li>
	<li>Use a medium with low nutrient content (relative to traditionally rich microbiological media) when screening environmental microbes from oligotrophic environments (such as the soil), since many oligotrophic bacteria do not grow when presented with high nutrient conditions21. A low nutrient medium also reduces colony size, increasing the throughput of the screen.</li>
	<li>Select a medium with low background fluorescence and good optical clarity.</li>
	<li>Optimize growth temperature to allow both the reporter strain and environmental microbes to grow simultaneously.</li>
	<li>Consider the addition of a buffering agent. The use of buffer in the plates will reduce the possibility of detecting pH-mediated changes in physiology22, unless such interactions are of interest.</li>
</ol>
3. Prepare Reporter Aliquots
For B. subtilis PtapA-yfp reporter:
<ol>
	<li>Streak reporter strain from -80 &#176;C frozen stock onto a fresh LB plate using a sterile toothpick or applicator stick.</li>
	<li>Grow overnight at 30 &#176;C.</li>
	<li>Perform serial dilutions in liquid culture to minimize background fluorescence arising from growth on a solid medium:
	<ol>
		<li>Inoculate a 5 ml liquid culture of LB and grow with shaking at 37 &#176;C.</li>
		<li>When the culture reaches an OD600 ~0.6, dilute into 5 ml fresh LB to an OD600 of 0.02.</li>
		<li>Grow at 37 &#176;C again with shaking until cultures reaches OD600 ~0.6.</li>
		<li>Repeat serial growth dilutions a total of 3x.</li>
		<li>Let final serial dilution culture grow to OD600 ~0.4.</li>
	</ol>
	</li>
	<li>Add glycerol to 15-20%.</li>
	<li>Aliquot 50-200 &#956;l into 0.5 ml microfuge tubes and freeze at -80 &#176;C.</li>
	<li>Make equivalent aliquots for the nonfluorescent parent strain of the reporter (they will be required during secondary screening).</li>
</ol>
For other bacterial species:
<ol>
	<li>Use cells that have a low background fluorescence (i.e. are grown under conditions where there is little expression from the promoter used in the reporter).</li>
	<li>Make and freeze aliquots containing known cfu/ml (colony forming units per ml) of the reporter strain so that an appropriate number of colonies can be grown on each coculture screen plate (see section above for details).</li>
	<li>Make equivalent aliquots for the nonfluorescent parent strain (they will be required during secondary screening).</li>
</ol>
4. Obtain Soil Samples
<ol>
	<li>Collect soil into sterile conical tubes or sterile bags using a spatula, discarding the top 0.5 cm of exposed surface soil.</li>
	<li>Add sterile saline (0.85% NaCl) at a ratio of 10 ml per 1 g of soil to make a soil slurry.</li>
	<li>Select a method to dislodge bacteria from soil particles: a) immediate use of fresh sample or b) delayed use after freezing of sample.
	<ol>
		<li>For immediate use, vortex slurry for 1 min.</li>
		<li>For delayed use, blend soil slurry in blender for three 1 min cycles, placing the blender jar on ice for 1 min rests between blending cycles.</li>
	</ol>
	</li>
	<li>Let soil slurry settle for ~1 min.</li>
	<li>Move upper aqueous layer to a fresh tube.</li>
	<li>Add glycerol to a final concentration of 15-20%.</li>
	<li>Aliquot 50-200 &#956;l into 0.5 ml microfuge tubes and freeze at -80 &#176;C.</li>
</ol>
5. Determine cfu/ml of Frozen Reporter And Soil Aliquots
<ol>
	<li>Thaw a frozen soil and reporter aliquot. Soil microbes can be thawed on ice. Because B. subtilis lyses at 4 &#176;C, those aliquots should be thawed quickly at RT to minimize the time spent at low temperature.</li>
	<li>Make two replicate serial dilutions (to 10-8) in 0.1x LB or other isotonic buffer.</li>
	<li>Plate 5 &#956;l spots of each serial dilution onto agar plates of the same medium that will be used for coculture screening.</li>
	<li>Grow at RT (or the temperature that will be using for screening).</li>
	<li>The next day, count the number of colonies within each spot and calculate cfu/ml of each of the frozen aliquots.</li>
</ol>
6. Confirm Aliquot Concentrations for Spread Screen Plates
<ol>
	<li>Thaw a frozen aliquot as in step 5.1.</li>
	<li>Dilute to 1 x 105, 2.5 x 105, 5 x 105, 1 x 106, and 2.5 x 107 cfu/ml.</li>
	<li>Add 50 &#956;l spot of each dilution to center of individual plates. These should yield plates with the calculated optimum of 25,000 colonies per plate, as well as 2- and 5- fold more and less, allowing the best actual dilution to be determined.</li>
	<li>Add approximately 20 sterile glass (3 mm) beads by gently tapping them onto the plate. Beads provide a more even distribution of colonies across the plate than a bent glass spreader does.</li>
	<li>Spread cells by keeping plates on the benchtop and shaking them back and forth, rotating them as you work, until the liquid has been absorbed. Do not continue shake the beads once the plate is dry; otherwise it will begin to kill the bacteria.</li>
	<li>Flip plates over and discard beads into waste beaker containing ethanol.</li>
	<li>Let plates grow for the time of your assay (e.g. 24 hr) at correct temperature for your assay (e.g. 24 &#176;C/RT).</li>
	<li>Using a dissecting stereoscope, count the number of colonies in two or more fields of view.</li>
	<li>Calculate the number of colonies per area, and determine how many colonies are on each plate.</li>
	<li>Adjust future dilutions as necessary. The actual number of colonies is not as important as having the same number on each comparable screen plate.</li>
</ol>
7. Prepare Coculture Plates
<ol>
	<li>Thaw reporter aliquot (and soil aliquot if frozen) as in step 5.1.</li>
	<li>Dilute reporter (to concentrations optimized in section 6) in 0.1x LB or other low nutrient, isotonic solution.
	<ol>
		<li>For frozen soil: Dilute to concentration optimized in section 6 in 0.1x LB or other low nutrient, isotonic solution.</li>
		<li>For fresh soil: Make dilutions of fresh soil slurry, based on prediction that the cfu/ml of the soil slurry could range from 10-4 to 10-9 cfu/ml.</li>
	</ol>
	</li>
	<li>Spot 50 &#956;l of soil and reporter dilutions onto center of coculture screen plate. Also, plate soil alone and reporter alone as controls.</li>
	<li>Spread using glass beads as described in step 6.4-6.6.</li>
	<li>If there are known growth conditions that activate your fluorescent reporter, streak or plate your reporter under these conditions and grow to use as a positive control during screening.</li>
	<li>Incubate at 24 &#176;C for 24 - 28 hr (or as appropriate for your reporter/assay)</li>
</ol>
8. Screen CoCulture Plates for Fluorescence
<ol>
	<li>Using brightfield illumination, focus your stereoscope so that the colonies are sharp.</li>
	<li>Look at the soil-only plates to determine whether your soil sample has autofluorescent colonies. If so, this soil may result in a high rate of false positives (with a concomitant increase in secondary screening).
	<ol>
		<li>Use a high ratio of reporter:soil in your coculture plates to increase the chance that an inducer will be surrounded by multiple reporter colonies, reducing the chance they will be detected as false-positives (Figure 2).</li>
		<li>Use a different fluorescent protein (one that emits in a different channel).</li>
	</ol>
	</li>
	<li>Determine the assay timing. For most new reporters, the timing of potential induction is unknown, and thus must be determined empirically.
	<ol>
		<li>Begin screening for fluorescence as soon as colonies become clearly visible with the dissecting stereoscope (magnification ~30X) and continue examining the plates periodically until growth has ceased and/or background fluorescence becomes too high.</li>
		<li>Once the time window of potential induction is determined for a particular reporter, it should be similar for all coculture plates containing that reporter, simplifying the monitoring of the screen plates. For the B. subtilis PtapA-yfp reporter, the appropriate time to examine the plates is between 24-28 hr after plating the cells; for the B. subtilis PsspB-yfp reporter, it is between 26-32 hr of growth.</li>
	</ol>
	</li>
	<li>Maximize your fluorescence sensitivity:
	<ol>
		<li>Ensure your fluorescence dissecting scope is in a dark room or surrounded by blackout curtains. Induction will likely be less intense than a constitutively produced FP and requires greater sensitivity to detect.</li>
		<li>Allow time for the fluorescence lamp to stabilize and your eyes to adjust to the darkness before attempting to detect fluorescence from your coculture plates (at least 1-2 min).</li>
	</ol>
	</li>
	<li>Using a positive control (if you have one), ensure that the magnification you are using allows you to detect fluorescence. The magnification is typically best if your field of view is approximately 30-50x your typical colony diameter (200-400X).</li>
	<li>Turn off the bright light and open the shutter for your fluorescence.</li>
	<li>After your eyes have adjusted to the darkness, slowly move the plate back and forth across your field of view, looking for bright spots.
	<ol>
		<li>Start from the top of the plate and use a zigzag pattern to move the plate side-to-side as you move towards the bottom of the plate.</li>
		<li>Practice moving the plate in brightfield to get a sense for how slowly to move to plate, and to be sure that you are covering the entire surface area.</li>
		<li>Move the plate slowly enough that the colonies do not become blurry. It is better to oversample the surface rather than miss areas.</li>
		<li>After one full sweep, turn the plate 90&#176; and repeat. Human eyes are remarkably good at detecting even faint fluorescence through this method.</li>
	</ol>
	</li>
	<li>If you detect fluorescence, stop moving the plate and go back and find the fluorescent area.</li>
	<li>Turning the brightfield on slowly, determine whether the fluorescence is associated with a bacterial colony (and not autofluorescent soil detritus or media components). If so, the nonfluorescent colonies proximal to the fluorescent colony are putative inducing organisms.</li>
</ol>
9. Isolate Putative Inducing Organisms
<ol>
	<li>Once induced (fluorescent) colonies have been identified, isolate those colonies secreting the inducing compounds.
	<ol>
		<li>If a high enough concentration of reporter colonies are growing on the plate (&#62;0.5:1 reporter:soil cfu), the putative inducing colonies will be surrounded by multiple fluorescent colonies (again, see Figure 2).</li>
		<li>In cases where the complexity of the coculture growth makes it ambiguous which colony is the inducer, isolate multiple potential inducing colonies for subsequent testing in the secondary screen.</li>
	</ol>
	</li>
	<li>Localize the colonies you want to pick in the center of the field of view.
	<ol>
		<li>If they are close to the edge of the plate, rotate the plate so that the lip of the plate is away from your dominant hand (i.e. if you are right-handed, put the lip of the plate on the left). This allows you to approach the colonies at a low angle, which improves the accuracy of your picking.</li>
	</ol>
	</li>
	<li>Place fresh plates to streak the putative inducing organisms on to nearby, as well as a waste beaker to discard used tips into.</li>
	<li>Leaving the fluorescence lamp on, turn the bright light slowly on, so that you will be able to identify (by shape and position), the fluorescent colony and the surrounding putative inducing organisms. You may need to go back and forth with the light a few times to be able to identify the colonies you want to pick when there is no fluorescence and only bright light.</li>
	<li>Use a glass rod (200 mm long x 5 mm diameter) to pick up a sterile, round 200 &#956;l gel-loading tip and hold it like a pencil. This is the picking tool you will use to isolate individual colonies.</li>
	<li>Rest your outer palm against the stage to stabilize it on the work surface. Place your other hand on the inner, thumb side of your hand to stabilize your picking tool.</li>
	<li>If necessary, flip again between fluorescence and brightfield views to identify the colonies you want to pick.</li>
	<li>Keeping the pipette tip above the plate surface, move it into your field of view and center it above the colony you would like to pick. The tip will be out of focus.</li>
	<li>Using the outer edge of your hand (which is resting on the microscope stage or work surface), pivot the pipette tip slowly down to the colony to be picked. Touch it very lightly, trying to minimize how many other colonies the tip contacts.</li>
	<li>Without rotating the picking tool, streak onto a section of a fresh plate. Spread with a soft touch to avoid gouging the agar. Because the number of cells transferred by this method is quite small (the colonies are much smaller than typically manipulated), a single continuous streak will result in isolated colonies.</li>
	<li>Repeat this process with other putative inducing organisms.</li>
	<li>Incubate plates at 24 &#176;C (or the temperature of your assay).</li>
</ol>
10. Streak Putative Inducing Organisms to Obtain Isolated Single Colonies
<ol>
	<li>Using a stereoscope, determine - based on colony structure or morphology - whether there are different colony types contained within each of your putative picked inducing organisms.
	<ol>
		<li>Look at the colonies using a stage where you can illuminate the colonies from both above as well as from below to detect potential colony differences.</li>
	</ol>
	</li>
	<li>Restreak each different morphotype onto a fresh plate and incubate until grown.</li>
	<li>Restreak one more time and let grow. If different morphotypes persist, continue to restreak to purity.</li>
</ol>
11. Retest Putative Inducing Organisms in Secondary Screen
<ol>
	<li>Retest all of the putative inducing organisms in a secondary screen to determine which are activating the fluorescent transcriptional reporter. The secondary screen consists of a lawn of microcolonies of the fluorescent transcriptional reporter strain, along with control plates, onto which putative inducing organisms are patched or spotted.</li>
	<li>Set up three identical plates: one containing a microcolony lawn of the fluorescent transcriptional reporter strain (at the same concentration as was used during coculture screening); one containing a microcolony lawn of the wild-type parent strain without a fluorescence reporter; and one containing no lawn.
	<ol>
		<li>Mark the top of the back of each plate to determine the plate orientation.</li>
		<li>Add positional markers for the patches/spots; up to ten putative inducing organisms can be tested on each set of plates (Figure 3).</li>
		<li>Thaw lawn aliquots (reporter and nonfluorescent parent strain) as in step 5.1.</li>
		<li>Spread 50 &#956;l of a 5 x 105 cfu/ml dilution onto lawn plates (or other optimized dilutions) using sterile beads as in step 6.4.</li>
		<li>Let plates dry.</li>
	</ol>
	</li>
	<li>Patch or spot putative inducing organisms:
	<ol>
		<li>Select patching if an easier and faster approach is desired, and it is acceptable to have a less precise number of cells deposited. To patch:
		<ol>
			<li>Touch a sterile toothpick to the colony to test - do not pick up all of the cells.</li>
			<li>Patch (make a small streak) onto the blank plate.</li>
			<li>Repeat patch with fresh toothpick onto reporter plate.</li>
			<li>Repeat patch with fresh toothpick onto control plate.</li>
		</ol>
		</li>
		<li>Select spotting if a quantitative and reproducible approach is desired. Spotting allows the number of deposited cells to be normalized (see reference 5 for complete details), and permits the relative potency of the different inducing organisms to be compared. To spot:
		<ol>
			<li>Resuspend putative inducing organisms in 1 ml of liquid media in a sterile plastic cuvette.</li>
			<li>Take the OD600 of the resuspensions.</li>
			<li>Using the formula X = 250 &#247; (OD600 - 0.5), add the volume X for each resuspension to 500 &#956;l of liquid medium to get a solution with an OD600 of 0.5. This method simplifies the required pipetting steps when performing multiple dilutions to normalize OD&#39;s because you can use the same volume (500 &#956;l) for all of your dilutants.</li>
			<li>Spot 1 &#956;l of each OD-normalized resuspension to each of the three plates.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Let grow at 24 &#176;C for 24-28 hr (or as appropriate for your reporter/assay).</li>
	<li>Use the fluorescent dissecting microscope to identify putative inducing organisms that activate your fluorescent reporter strain but not your parental control strain. These isolates are your positive hits - the environmental microbes that secrete compounds that induce your phenotype of interest.</li>
</ol>

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D., Kolter, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pseudomonas-Saccharomyces+interactions:+influence+of+fungal+metabolism+on+bacterial+physiology+and+survival.">Pseudomonas-Saccharomyces interactions: influence of fungal metabolism on bacterial physiology and survival.</a> J. Bacteriol. 187, 940-948 (2005).</li><li>Branda, S. S., Chu, F., Kearns, D. B., Losick, R., Kolter, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+major+protein+component+of+the+Bacillus+subtilis+biofilm+matrix.">A major protein component of the Bacillus subtilis biofilm matrix.</a> Mol. Microbiol. 59, 1229-1238 (2006).</li><li>Zengler, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cultivating+the+uncultured.">Cultivating the uncultured.</a> Proc. Natl. Acad. Sci. U.S.A. 99, 15681-15686 (2002).</li></ol>

The authors declare that they have no competing financial interests.

One of the inherent limitations of this protocol is that it relies on the cultivability of microbial organisms. As has been well documented24, most microbial life on the planet cannot (yet) be grown under the culturing conditions explored to date. Thus, a huge number of interactions between microbial species that are occurring in natural settings will go undetected using this approach. However, since our desire is to not only identify the existence of such interactions, but then also study the mechanisms and m...

We are interested in understanding how the metabolites that bacteria secrete affect the physiology and development of neighboring microbes. Many metabolites have been characterized for their bioactive effects on other microbes. Two well-described examples include antibiotics, which inhibit the growth of other microbes, and quorum sensing molecules, which alter the global gene expression of other microbes. However, bacteria produce many other small molecule natural products that have no known bioactivities1. We hypothesize that bacteria have evolved and preserved the ability to produce some of these metabolites because they allow them to modulate the cellular physiology of their microbial neighbors in the complex microbial communities within which most bacteria exist.
Bacillus subtilis cell types
We have focused our studies on chemically mediated microbial interactions that involve Bacillus subtilis. This is not only because of its status as the Gram-positive model bacterium and the resultant genetic tools available for its manipulation, but also because of its ability to differentiate into characterized cell types. Examples include cells that are: swimming; producing the extracellular matrix that is required for robust biofilm formation; competent to take up DNA from the environment; and sporulating, among others2. Each of these cell types expresses a characteristic transcriptional regulon that makes them physiologically and/or physically distinct from their genetically identical siblings. Under many growth conditions, multiple cell types coexist as various subpopulations within a single colony of B. subtilis cells3. Although many species of bacteria may exhibit analogous cell type heterogeneity, this phenomenon has been particularly well studied in B. subtilis.
In particular, genes that are upregulated within each of these specific B. subtilis cell types have been identified. Identifying such upregulated genes is essential for the work described here because many of these microbial phenotypes of interest are difficult or impossible to observe directly. For instance, we cannot visually detect a trait such as swimming on solid (1.5%) agar plates, even though a subpopulation of B. subtilis cells produce flagella under those conditions3. Another example is biofilm matrix-production. Matrix production can be visualized by colony morphology (as it results in macroscopically wrinkly colonies), but only on certain growth medium, and only after multiple days of growth4. However, by knowing which genes are upregulated during differentiation, we can construct transcriptional reporters that act as markers for cellular differentiation into these cell types.
Reporter constructs
These fluorescent transcriptional reporters consist of the promoters for cell-type specific genes driving the production of a reporter gene, for instance a fluorescent protein. Examples include Phag-yfp (for swimming cells), PtapA-yfp (for biofilm matrix-producing cells), and PsspB-yfp (for sporulating cells), where Px indicates the promoter region for gene x. These reporter constructs are integrated into a neutral locus on the chromosome (Figure 1 and see below) so that the native regulation of the phenotype is left intact. However, now when a cell expresses this phenotype, it also expresses a fluorescent protein. This provides an easily visualized read-out of the activation of particular phenotypic behavior, allowing us to screen for microbes that activate this physiological response. Although such reporters are commonly used in microbiology, they have not been broadly applied in screens to identify metabolic interactions between microbes before this method was described5.
There are a number of important considerations in the design and construction of cell-type-specific reporter strains. We have utilized exclusively transcriptional fluorescent reporters, although other types of constructs are certainly possible. We discourage the use of translational fusions as markers for cell type differentiation in our screen, however, for two reasons: 1) the desire to leave the native cell-type-specific protein unperturbed, and 2) the recognition that a diffuse, cell-wide fluorescence will be easier to detect than localized puncta within cells (common with translational fusions).
Reporter gene selection
After deciding to use transcription as a read-out, the reporter gene must be selected (e.g. LacZ, fluorescence, or luciferase). LacZ has the advantage of needing the least specialized equipment to detect, but there is a much higher likelihood of false positives among environmental microbes. In our hands, the background level of Lac+ organisms among soil microbes was prohibitively high (&#62;&#62;10% of soil microbes were blue (Lac+) on X-gal plates; data not shown). It is possible that by titrating the concentration of X-gal in the medium, this could be optimized to allow the use of an X-gal reporter, although we did not attempt this. Luciferase provides high sensitivity of detection and is the most orthogonal reporter: there is almost no chance of environmental microbes being inherently luminescent. However, we found it difficult to identify instrumentation at our institution that allowed luminescence detection across entire Petri plates, as most were designed to scan only localized regions in multi-well plates. There might also be complications in visualizing luminescent colonies in a manner that also allowed the simultaneous physical isolation of inducing organisms. While using fiduciaries may have made this possible, we instead elected to use fluorescent transcriptional reporters, which were proven to work in B. subtilis, provided adequate sensitivity of detection and low false positive rates among soil organisms, and allowed to use of easily available instrumentation for both visualization and isolation procedures.
Fluorophore selection
The specific fluorophore selected will depend on your bacterial species, the agar growth medium you are using, and the particular fluorescence filter sets you have available. With our instrumentation, we found that both the B. subtilis colonies themselves and the agar they were grown on exhibited less background fluorescence when YFP (yellow fluorescent protein) filters were used, making that reporter superior to GFP (green fluorescent protein) in our hands. The codon usage of fluorescent proteins are frequently optimized for eukaryotes, making it important to select a fluorophore either known from the literature to work in your bacterial species, or to test it explicitly using a constitutive promoter. A large number of ever-evolving fluorescent protein variants are currently available6, which have been reviewed in a number of sources7,8, some of which explicitly provide guidance on choosing an appropriate fluorescent protein for your experiment9.
Promoter selection
The selection of a promoter will largely depend on your cell type or phenotype of interest. For organisms such as B. subtilis, some cell-type specific reporter genes have been established in the literature. For other bacterial strains, examining microarray or transcriptional data will be necessary to provide information about which genes are highly upregulated under the conditions where your cell type of interest is manifested. A recent study cataloged the transcription of B. subtilis under 104 different growth conditions using tiling microarrays10. This paper provides comprehensive information about which genes are highly upregulated under different conditions, which is invaluable for less-well-characterized phenotypes.
Rather than mapping precise promoter regions for every gene of interest, we typically simply use the sequence 200-500 bp upstream of the gene as the promoter. The exact sequence length depends on the genomic context: shorter regions are used when necessary to avoid including upstream coding regions from neighboring open reading frames.
Neutral loci and integration
How to maintain the reporter construct in your bacterial strain becomes the final question in designing a fluorescent transcriptional reporter strain. In bacteria, genes of interest are frequently maintained on plasmids using antibiotic selection. However, it may not be possible to use antibiotics during coculture without killing the environmental microbes. If plasmids are stably maintained in your bacterial species, it may be possible grow your bacteria containing a plasmid-borne reporter in the presence of antibiotics to prepare your reporter for screening, and then eliminate antibiotics during the coculture itself in the hope that the plasmid will be sufficiently maintained to allow for fluorescence. However, if plasmids are easily lost in your bacterium, or are lost under stress conditions, this will not be a viable option. In many cases, the best solution will be to integrate the reporter construct onto the bacterial chromosome, which allows stable maintenance of the reporter even in the absence of selection. In order for the integration to not disrupt the normal expression or regulation of your gene of interest, we recommend integrating into an ectopic site on the chromosome that can act as a &#34;neutral locus.&#34; In B. subtilis these integration sites are genes that - when mutated - convey a phenotype in certain minimal media (allowing integrants to be identified without antibiotic selection), yet do not alter growth or sporulation rates in rich media, and include such genes as amyE , lacA, thrC , pyrD , gltA, and sacA (conveying the ability to utilize starch, &#946;-galactosides, threonine, uracil, glutamate, and sucrose, respectively)11-13 .
While integration in these genes have been used reliably for many years in B. subtilis (particularly at amyE and lacA), similar knowledge may not be available for genes in many other bacterial species. The use of phage attachment sites are great alternatives for neutral chromosomal integration sites: many species-specific14-16, as well as general integration sites such as the Tn7 attachment site (attTn7) have been identified and utilized for gene insertions in many bacterial species17,18.
Environmental microbes
We use soil as a direct source of environmental microbes for our coculture screen. The soil contains a high diversity of microbes, and many of these organisms are rich source of natural products. By using liquid suspensions of soil placed directly onto plates with our fluorescent transcriptional reporter strain (without prior isolation of bacteria from the soil), we greatly simplify the experimental approach. The soil can either be used immediately after harvesting, or be frozen at -80 &#176;C for future use. Immediate use has the advantage that a greater diversity of microbes can potentially be grown, including those that will not survive freezing well. It has the disadvantage that the concentration of cultivable soil organisms from these samples is unknown, increasing the number of screen plates that must be used. Delayed use has the advantage that the cfu/ml for each soil source can be determined in advance, allowing an optimized number of colonies to be grown on each screen plate. However, it requires that the soil organisms be capable of surviving freezing.
Note that diversifying the inducer pool being examined (i.e. the soil sources) appears to be more effective at identifying new interspecies interactions than in-depth screening on the same soil: greater phylogenetic diversity was observed in the hits identified in our matrix-induction screen as additional soil sources were examined rather than screening the same soil sources more thoroughly (E.A. Shank and R. Kolter, Harvard Medical School, unpublished results).
Overview
The approach we describe here is straightforward in terms of its technical requirements. It involves: 1) constructing a fluorescent transcriptional reporter in B. subtilis or another bacterial species of interest, 2) identifying conditions under which this reporter is not activated, 3) preparing aliquots of this reporter strain and organisms to be screened (in our case soil, but other sources could be utilized instead), 4) mixing these two sets of microbes on solid media, 5) identifying and isolating putative inducing organisms, and 6) confirming that these organisms do indeed activate this phenotype in a secondary screen. Once identified, these organisms and their metabolites provide us with chemical tools to modulate bacterial behavior, to study bacterial physiology and microbial interactions, and to potentially act as novel scaffolds for future therapeutic compounds.

<table border="1">
	<tbody>
		<tr>
			<td>Name of Reagent/ Equipment</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments/Description</td>
		</tr>
		<tr>
			<td>Spectrophotometer</td>
			<td></td>
			<td></td>
			<td>Any spectrophotomer capable of measuing OD600 absorbance values.</td>
		</tr>
		<tr>
			<td>Luria broth, Lennox</td>
			<td>VWR</td>
			<td>80017-484</td>
			<td>Alternative media sources may be necessary.</td>
		</tr>
		<tr>
			<td>Glass beads, 3 mm</td>
			<td>VWR</td>
			<td>26396-508</td>
			<td></td>
		</tr>
		<tr>
			<td>Gel loading tips, round</td>
			<td>VWR</td>
			<td>29442-666</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass rods</td>
			<td>VWR</td>
			<td>59060-069</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence dissecting stereoscope</td>
			<td>Zeiss</td>
			<td>N/A</td>
			<td>The author used a Zeiss Stemi SV6 dissection stereoscope with an EXFO X-cite 120 fluorescent light source, a long-pass YFP filter cube, an achromat 0.63X objective, 10X eyepieces, and an Axio HRC HR digital camera. Most screening was done with the focusing mount at 2.0-3.2X. Any dissecting stereoscope with fluorescence capabilities is fine, provided you have the correct filters for the FP you are using. It is best if there is a shutter that allows you to easily switch between brightfield and fluorescense, as well as a stage that allows illumination from above and below. If you want to capture images, an attached camera is also necessary.</td>
		</tr>
	</tbody>
</table>

using coculture to detect chemically mediated interspecies interactions

In nature, bacteria rarely exist in isolation; they are instead surrounded by a diverse array of other microorganisms that alter the local environment by secreting metabolites. These metabolites have the potential to modulate the physiology and differentiation of their microbial neighbors and are likely important factors in the establishment and maintenance of complex microbial communities. We have developed a fluorescence-based coculture screen to identify such chemically mediated microbial interactions. The screen involves combining a fluorescent transcriptional reporter strain with environmental microbes on solid media and allowing the colonies to grow in coculture. The fluorescent transcriptional reporter is designed so that the chosen bacterial strain fluoresces when it is expressing a particular phenotype of interest (i.e. biofilm formation, sporulation, virulence factor production, etc.) Screening is performed under growth conditions where this phenotype is not expressed (and therefore the reporter strain is typically nonfluorescent). When an environmental microbe secretes a metabolite that activates this phenotype, it diffuses through the agar and activates the fluorescent reporter construct. This allows the inducing-metabolite-producing microbe to be detected: they are the nonfluorescent colonies most proximal to the fluorescent colonies. Thus, this screen allows the identification of environmental microbes that produce diffusible metabolites that activate a particular physiological response in a reporter strain. This publication discusses how to: a) select appropriate coculture screening conditions, b) prepare the reporter and environmental microbes for screening, c) perform the coculture screen, d) isolate putative inducing organisms, and e) confirm their activity in a secondary screen. We developed this method to screen for soil organisms that activate biofilm matrix-production in Bacillus subtilis; however, we also discuss considerations for applying this approach to other genetically tractable bacteria.

This screen was used to identify soil organisms secreting compounds that alter the physiology of B. subtilis. The results described here focus on the matrix-producing cell type of B. subtilis, which produces the protein and exopolysaccharide that are required for robust biofilm formation in this bacterium. We selected the promoter of the tapA-sipW-tasA operon for our fluorescent reporter construct (PtapA-yfp). This operon encodes the protein structural component of the matri...

Watch this Scientific Journal Video about Using Coculture to Detect Chemically Mediated Interspecies Interactions at JoVE.com

Using Coculture to Detect Chemically Mediated Interspecies Interactions

Bacteria produce secreted compounds that have the potential to affect the physiology of their microbial neighbors. Here we describe a coculture screen that allows detection of such chemically mediated interspecies interactions by mixing soil microbes with fluorescent transcriptional reporter strains of Bacillus subtilis on solid media.

In nature, bacteria rarely exist in isolation; they are instead surrounded by a diverse array of other microorganisms that alter the local environment by ...

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13.5K Views.  University of North Carolina at Chapel Hill. Bacteria produce secreted compounds that have the potential to affect the physiology of their microbial neighbors. Here we describe a coculture screen that allows detection of such chemically mediated interspecies interactions by mixing soil microbes with fluorescent transcriptional reporter strains of Bacillus subtilis on solid media.

Video: Using Coculture to Detect Chemically Mediated Interspecies Interactions

Microfluidic Co-culture of Epithelial Cells and Bacteria for Investigating Soluble Signal-mediated Interactions

The human gastrointestinal (GI) tract is a unique environment in which intestinal epithelial cells and non-pathogenic (commensal) bacteria coexist. It has been proposed that the microenvironment that the pathogen encounters in the commensal layer is important in determining the extent of colonization. Current culture methods for investigating pathogen colonization are not well suited for investigating this hypothesis as they do not enable co-culture of bacteria and epithelial cells in a manner that mimics the GI tract microenvironment. Here we describe a microfluidic co-culture model that enables independent culture of eukaryotic cells and bacteria, and testing the effect of the commensal microenvironment on pathogen colonization. The co-culture model is demonstrated by developing a commensal Escherichia coli biofilm among HeLa cells, followed by introduction of enterohemorrhagic E. coli (EHEC) into the commensal island, in a sequence that mimics the sequence of events in GI tract infection.

This protocol describes a microfluidic co-culture model for simultaneous and localized culture of epithelial cells and bacteria. This model can be used for investigating the role of different soluble molecular signals on pathogenesis as well as screen the effectiveness of putative probiotic bacterial strains.

The human gastrointestinal (GI) tract is a unique environment in which intestinal epithelial cells and non-pathogenic (commensal) bacteria coexist. It has ...

Analysis of Schwann-astrocyte Interactions Using In Vitro Assays

Schwann cells are one of the commonly used cells in repair strategies following spinal cord injuries. Schwann cells are capable of supporting axonal regeneration and sprouting by secreting growth factors 1,2 and providing growth promoting adhesion molecules 3 and extracellular matrix molecules 4. In addition they myelinate the demyelinated axons at the site of injury 5.
However following transplantation, Schwann cells do not migrate from the site of implant and do not intermingle with the host astrocytes 6,7. This results in formation of a sharp boundary between the Schwann cells and astrocytes, creating an obstacle for growing axons trying to exit the graft back into the host tissue proximally and distally. Astrocytes in contact with Schwann cells also undergo hypertrophy and up-regulate the inhibitory molecules 8-13.
In vitro assays have been used to model Schwann cell-astrocyte interactions and have been important in understanding the mechanism underlying the cellular behaviour.
These in vitro assays include boundary assay, where a co-culture is made using two different cells with each cell type occupying different territories with only a small gap separating the two cell fronts. As the cells divide and migrate, the two cellular fronts get closer to each other and finally collide. This allows the behaviour of the two cellular populations to be analyzed at the boundary. Another variation of the same technique is to mix the two cellular populations in culture and over time the two cell types segregate with Schwann cells clumped together as islands in between astrocytes together creating multiple Schwann-astrocyte boundaries.
The second assay used in studying the interaction of two cell types is the migration assay where cellular movement can be tracked on the surface of the other cell type monolayer 14,15. This assay is commonly known as inverted coverslip assay. Schwann cells are cultured on small glass fragments and they are inverted face down onto the surface of astrocyte monolayers and migration is assessed from the edge of coverslip.
Both assays have been instrumental in studying the underlying mechanisms involved in the cellular exclusion and boundary formation. Some of the molecules identified using these techniques include N-Cadherins 15, Chondroitin Sulphate proteoglycans(CSPGs) 16,17, FGF/Heparin 18, Eph/Ephrins19.
This article intends to describe boundary assay and migration assay in stepwise fashion and elucidate the possible technical problems that might occur.

Analysis of Schwann-astrocyte Interactions Using In Vitro Assays

This article intends to describe in stepwise fashion the commonly used in vitro assays used in studying Schwann cell-asrtocyte interactions.

Schwann cells are one of the commonly used cells in repair strategies following spinal cord injuries. Schwann cells are capable of supporting axonal ...

The MultiBac Protein Complex Production Platform at the EMBL

Proteomics research revealed the impressive complexity of eukaryotic proteomes in unprecedented detail. It is now a commonly accepted notion that proteins in cells mostly exist not as isolated entities but exert their biological activity in association with many other proteins, in humans ten or more, forming assembly lines in the cell for most if not all vital functions.1,2 Knowledge of the function and architecture of these multiprotein assemblies requires their provision in superior quality and sufficient quantity for detailed analysis. The paucity of many protein complexes in cells, in particular in eukaryotes, prohibits their extraction from native sources, and necessitates recombinant production. The baculovirus expression vector system (BEVS) has proven to be particularly useful for producing eukaryotic proteins, the activity of which often relies on post-translational processing that other commonly used expression systems often cannot support.3 BEVS use a recombinant baculovirus into which the gene of interest was inserted to infect insect cell cultures which in turn produce the protein of choice. MultiBac is a BEVS that has been particularly tailored for the production of eukaryotic protein complexes that contain many subunits.4 A vital prerequisite for efficient production of proteins and their complexes are robust protocols for all steps involved in an expression experiment that ideally can be implemented as standard operating procedures (SOPs) and followed also by non-specialist users with comparative ease. The MultiBac platform at the European Molecular Biology Laboratory (EMBL) uses SOPs for all steps involved in a multiprotein complex expression experiment, starting from insertion of the genes into an engineered baculoviral genome optimized for heterologous protein production properties to small-scale analysis of the protein specimens produced.5-8 The platform is installed in an open-access mode at EMBL Grenoble and has supported many scientists from academia and industry to accelerate protein complex research projects.

Protein complexes catalyze key cellular functions. Detailed functional and structural characterization of many essential complexes requires recombinant production. MultiBac is a baculovirus/insect cell system particularly tailored for expressing eukaryotic proteins and their complexes. MultiBac was implemented as an open-access platform, and standard operating procedures developed to maximize its utility.

Proteomics  research revealed the impressive complexity of eukaryotic proteomes in  unprecedented detail. It is now a commonly accepted notion that ...

Rapid Synthesis and Screening of Chemically Activated Transcription Factors with GFP-based Reporters

Synthetic biology aims to rationally design and build synthetic circuits with desired quantitative properties, as well as provide tools to interrogate the structure of native control circuits. In both cases, the ability to program gene expression in a rapid and tunable fashion, with no off-target effects, can be useful. We have constructed yeast strains containing the ACT1 promoter upstream of a URA3 cassette followed by the ligand-binding domain of the human estrogen receptor and VP16. By transforming this strain with a linear PCR product containing a DNA binding domain and selecting against the presence of URA3, a constitutively expressed artificial transcription factor (ATF) can be generated by homologous recombination. ATFs engineered in this fashion can activate a unique target gene in the presence of inducer, thereby eliminating both the off-target activation and nonphysiological growth conditions found with commonly used conditional gene expression systems. A simple method for the rapid construction of GFP reporter plasmids that respond specifically to a native or artificial transcription factor of interest is also provided.

This protocol describes an experimental procedure for the rapid construction of artificial transcription factors (ATFs) with cognate GFP reporters&#160;and quantification of the ATFs ability to stimulate GFP expression via flow cytometry.

Synthetic biology aims to rationally design and build synthetic circuits with desired quantitative properties, as well as provide tools to interrogate the ...

Determination of Protein-ligand Interactions Using Differential Scanning Fluorimetry

A wide range of methods are currently available for determining the dissociation constant between a protein and interacting small molecules. However, most of these require access to specialist equipment, and often require a degree of expertise to effectively establish reliable experiments and analyze data. Differential scanning fluorimetry (DSF) is being increasingly used as a robust method for initial screening of proteins for interacting small molecules, either for identifying physiological partners or for hit discovery. This technique has the advantage that it requires only a PCR machine suitable for quantitative PCR, and so suitable instrumentation is available in most institutions; an excellent range of protocols are already available; and there are strong precedents in the literature for multiple uses of the method. Past work has proposed several means of calculating dissociation constants from DSF data, but these are mathematically demanding. Here, we demonstrate a method for estimating dissociation constants from a moderate amount of DSF experimental data. These data can typically be collected and analyzed within a single day. We demonstrate how different models can be used to fit data collected from simple binding events, and where cooperative binding or independent binding sites are present. Finally, we present an example of data analysis in a case where standard models do not apply. These methods are illustrated with data collected on commercially available control proteins, and two proteins from our research program. Overall, our method provides a straightforward way for researchers to rapidly gain further insight into protein-ligand interactions using DSF.

Differential scanning fluorimetry is a widely used method for screening libraries of small molecules for interactions with proteins. Here, we present a straightforward method to extend these analyses to provide an estimate of the dissociation constant between a small molecule and its protein partner.

A wide range of methods are currently available for determining the dissociation constant between a protein and interacting small molecules. However, most ...

Probing High-density Functional Protein Microarrays to Detect Protein-protein Interactions

High-density functional protein microarrays containing ~4,200 recombinant yeast proteins are examined for kinase protein-protein interactions using an affinity purified yeast kinase fusion protein containing a V5-epitope tag for read-out. Purified kinase is obtained through culture of a yeast strain optimized for high copy protein production harboring a plasmid containing a Kinase-V5 fusion construct under a GAL inducible promoter. The yeast is grown in restrictive media with a neutral carbon source for 6 hr followed by induction with 2% galactose. Next, the culture is harvested and kinase is purified using standard affinity chromatographic techniques to obtain a highly purified protein kinase for use in the assay. The purified kinase is diluted with kinase buffer to an appropriate range for the assay and the protein microarrays are blocked prior to hybridization with the protein microarray. After the hybridization, the arrays are probed with monoclonal V5 antibody to identify proteins bound by the kinase-V5 protein. Finally, the arrays are scanned using a standard microarray scanner, and data is extracted for downstream informatics analysis1,2 to determine a high confidence set of protein interactions for downstream validation in vivo.

Using protein microarrays containing nearly the entire S. cerevisiae proteome is probed for rapid unbiased interrogation of thousands of protein-protein interactions in parallel. This method can be utilized for protein-small molecule, posttranslational modification, and other assays in high-throughput.

High-density functional protein microarrays containing ~4,200 recombinant yeast proteins are examined for kinase protein-protein interactions using an ...

Using Caenorhabditis elegans to Screen for Tissue-Specific Chaperone Interactions

Correct folding and assembly of proteins and protein complexes are essential for cellular function. Cells employ quality control pathways that correct, sequester or eliminate damaged proteins to maintain a healthy proteome, thus ensuring cellular proteostasis and preventing further protein damage. Because of redundant functions within the proteostasis network, screening for detectable phenotypes using knockdown or mutations in chaperone-encoding genes in the multicellular organism Caenorhabditis elegans results in the detection of minor or no phenotypes in most cases. We have developed a targeted screening strategy to identify chaperones required for a specific function and thus bridge the gap between phenotype and function. Specifically, we monitor novel chaperone interactions using RNAi synthetic interaction screens, knocking-down chaperone expression, one chaperone at a time, in animals carrying a mutation in a chaperone-encoding gene or over-expressing a chaperone of interest. By disrupting two chaperones that individually present no gross phenotype, we can identify chaperones that aggravate or expose a specific phenotype when both perturbed. We demonstrate that this approach can identify specific sets of chaperones that function together to modulate the folding of a protein or protein complexes associated with a given phenotype.

Using Caenorhabditis elegans to Screen for Tissue-Specific Chaperone Interactions

To study chaperone-chaperone and chaperone-substrate interactions, we perform synthetic interaction screens in Caenorhabditis elegans using RNA interference in combination with mild mutations or over-expression of chaperones and monitor tissue-specific protein dysfunction at the organismal level.

Correct folding and assembly of proteins and protein complexes are essential for cellular function. Cells employ quality control pathways that correct, ...

Graphene Enclosure of Chemically Fixed Mammalian Cells for Liquid-Phase Electron Microscopy

A protocol is described for investigating the human epidermal growth factor receptor 2 (HER2) in the intact plasma membrane of breast cancer cells using scanning transmission electron microscopy (STEM). Cells of the mammalian breast cancer cell line SKBR3 were grown on silicon microchips with silicon nitride (SiN) windows. Cells were chemically fixed, and HER2 proteins were labeled with quantum dot nanoparticles (QDs), using a two-step biotin-streptavidin binding protocol. The cells were coated with multilayer graphene to maintain a hydrated state, and to protect them from electron beam damage during STEM. To examine the stability of the samples under electron beam irradiation, a dose series experiment was performed. Graphene-coated and non-coated samples were compared. Beam induced damage, in the form of bright artifacts, appeared for some non-coated samples at increased electron dose D, while no artifacts appeared on coated samples.

Presented here is a protocol for labeling membrane proteins in mammalian cells and coating the sample with graphene for liquid-phase scanning transmission electron microscopy. The stability of the samples against the damage caused by radiation can also studied with this protocol.

A protocol is described for investigating the human epidermal growth factor receptor 2 (HER2) in the intact plasma membrane of breast cancer cells using ...

Culture Methods to Study Apical-Specific Interactions using Intestinal Organoid Models

The lining of the gut epithelium is made up of a simple layer of specialized epithelial cells that expose their apical side to the lumen and respond to external cues. Recent optimization of in vitro culture conditions allows for the re-creation of the intestinal stem cell niche and the development of advanced 3-dimensional (3D) culture systems that recapitulate the cell composition and the organization of the epithelium. Intestinal organoids embedded in an extracellular matrix (ECM) can be maintained for long-term and self-organize to generate a well-defined, polarized epithelium that encompasses an internal lumen and an external exposed basal side. This restrictive nature of the intestinal organoids presents challenges in accessing the apical surface of the epithelium in vitro and limits the investigation of biological mechanisms such as nutrient uptake and host-microbiota/host-pathogen interactions. Here, we describe two methods that facilitate access to the apical side of the organoid epithelium and support the differentiation of specific intestinal cell types. First, we show how ECM removal induces an inversion of the epithelial cell polarity and allows for the generation of apical-out 3D organoids. Second, we describe how to generate 2-dimensional (2D) monolayers from single cell suspensions derived from intestinal organoids, comprised of&#160;mature and differentiated cell types. These techniques provide novel tools to study apical-specific interactions of the epithelium with external cues in vitro and promote the use of organoids as a platform to facilitate precision medicine.

Here we present two protocols that allow the modeling of intestinal apical-specific interactions. Organoid-derived intestinal monolayers and Air-Liquid Interface (ALI) cultures facilitate the generation of well-differentiated epithelia accessible from both luminal and basolateral sides, whereas polarity-inverted intestinal organoids expose their apical side and are amenable to high throughput assays.

The lining of the gut epithelium is made up of a simple layer of specialized epithelial cells that expose their apical side to the lumen and respond to ...

Bioinformatics Resources for the Study of Glycan-Mediated Protein Interactions

The Glyco@Expasy initiative was launched as a collection of interdependent databases and tools spanning several aspects of knowledge in glycobiology. In particular, it aims at highlighting interactions between glycoproteins (such as cell surface receptors) and carbohydrate-binding proteins mediated by glycans. Here, major resources of the collection are introduced through two illustrative examples centered on the N-glycome of the human Prostate Specific Antigen (PSA) and the O-glycome of human serum proteins. Through different database queries and with the help of visualization tools, this article shows how to explore and compare content in a continuum to gather and correlate otherwise scattered pieces of information. Collected data are destined to feed more elaborate scenarios of glycan function. Glycoinformatics introduced here is, therefore, proposed as a means to either strengthen, shape, or refute assumptions on the specificity of a protein glycome in a given context.

This protocol illustrates how to explore, compare, and interpret human protein glycomes with online resources.

The Glyco@Expasy initiative was launched as a collection of interdependent databases and tools spanning several aspects of knowledge in glycobiology. In ...