This work was supported by research funds provided by the Department of Energy Biological and Environmental Research (DOE-BER 0000217519 to E.A.S.), the National Science Foundation (INSPIRE IOS-1343020 to E.A.S). SLH was also supported by the National Science Foundation Graduate Research Fellowship Program. We thank Dr. Jeffery Dangl for providing bacterial strains and invaluable insight. We thank Dr. Andrew Klein and Matthew J. Powers for experimental suggestions. Finally, SLH would like to thank connections on social media for reminding us that disseminating science is a privilege and a responsibility, especially through creative and accessible means.

NOTE: The experimental setup is described for clarity and used to generate the representative results included in this report, but conditions can be modified as desired. All steps should be performed using PPE and following institutional and federal reccomendations for safety, according to the BSL status of the bacteria used.
1. Characterization of bacteria
<ol>
	<li>Determine the morphology of bacteria on the growth medium agar plate. Resuspend cells at an approximate OD600 = 0.5 and plate a 1 &#181;L volume onto agar medium of choice. Add X-gal to agar plates to a final concentration of 20 mg/mL to better differentiate individual members of the specific bacterial community. Grow at 24 &#176;C or 30 &#176;C until colonies form, then take pictures of and notes on colony morphology.</li>
	<li>Define the correlation between each bacterial strain&#8217;s optical density and the number of CFU (colony forming units) per mL16. Resuspend bacteria in 1 mL of water in a 24-well plate to an approximate OD600 = 5, perform two-fold serial dilutions, monitor OD600 of all dilutions, and plate each to determine the viable CFU/mL in each sample at multiple optical densities.</li>
	<li>Determine the maximum sonication tolerance for each bacterial strain. To do this, aliquot cells into a 24-well plate containing liquid medium, reserving some cells as an unsonicated control sample. Using an ultrasonicator with a 24-tip horn attachment, apply three rounds of 12 s of sonication at 40 amp with 2 s pulses. 
	NOTE: The use of a 24-well ultrasonicator is advised to facilitate the downstream multiplexing of processing plant samples, but if one is not available, use an ultrasonicator fitted with a microtip and perform each sample sonication independently. Always wear earmuffs rated to at least 25 NRR protection.</li>
	<li>Perform 10-fold serial dilutions of the sonicated and unsonicated samples and spot onto agar plates. Determine whether there is a reduction in viable cells after sonication. If so, use a fresh sample and repeat sonication step using a reduced total sonication time or amplitude until the treatment has no effect on final CFU/mL17.</li>
</ol>
2. Preparation of Arabidopsis thaliana seedlings on a plastic mesh
<ol>
	<li>Create disks of the plastic mesh using a standard hole puncher.
	<ol>
		<li>Collect the disks in a glass container with a loose cover of aluminum foil, and sterilize using an autoclave set to a 20 min dry cycle13.</li>
		<li>Using flame-sterilized tweezers, distribute approximately 40 sterilized mesh disks in a single layer across the surface of a plant-growth-medium agar plate. Use 0.5x Murashige and Skoog (MS) salts, containing 500 mg/L of MES buffer [2-(N-morpholino)ethanesulfonic acid] and 1.5% Bacto agar, as plant growth medium, with 50 &#181;g/mL benomyl added to the limit fungal contamination of the seedlings.</li>
	</ol>
	</li>
	<li>Prepare axenic seeds of A. thaliana as previously described17.
	<ol>
		<li>Place approximately 100&#8211;300 seeds each into individual centrifuge tubes in a rack and place into a resealable glass or heavy plastic container (&#8220;jar&#8221;) in a fume hood.</li>
		<li>Using caution, place a beaker of 100 mL of bleach into the jar, add 3 mL of concentrated HCl to the bleach, and immediately seal the jar and allow fumes to sterilize seeds for at least 4 h.</li>
		<li>Carefully remove the tubes of sterilized seeds from underneath the jar and seal.</li>
	</ol>
	</li>
	<li>Place two seeds at the center of each mesh. Seal plates with surgical tape and incubate for 2&#8211;6 days at 4 &#176;C in darkness to vernalize seeds.</li>
	<li>To germinate and grow seedlings, place the plate agar side down in a plant growth chamber for 8&#8211;10 days under short day settings: 9 h of light at 21 &#176;C and 15 h of dark at 18 &#176;C (Figure 1, step 2).</li>
</ol>
3. Colonization of plants in liquid bacterial growth medium
<ol>
	<li>Add 1 mL of bacterial growth medium to each well of a sterile 24-well plate, except for media-only control wells. Use Lennox Luria Broth (10 g of tryptone, 5 g of yeast extract, 5 g of NaCl) as the bacterial growth medium.</li>
	<li>Transfer the germinated seedlings embedded in mesh from agar plates to the liquid (Figure 1, step 3a).
	<ol>
		<li>Gently peel the mesh containing two germinated seedlings up and off the agar plate using flame-sterilized forceps. Choose mesh with equally sized and undamaged seedlings.</li>
		<li>If removal from the agar is not smooth, discard that mesh and plant. Transfer one float to each well of bacterial growth liquid, root side down.</li>
	</ol>
	</li>
	<li>Inoculate bacteria into wells containing floating seedlings.
	<ol>
		<li>Resuspend bacteria grown overnight on agar plates to an OD600 equivalent to 108 CFU/mL in the bacterial growth medium liquid. Add 10 &#181;L of bacterial suspension to each well for a final concentration of 106 CFU bacteria per well.</li>
		<li>If preparing a mix of bacteria, resuspend each to the OD600 equivalent to 108 CFU/mL, mix in equal proportions, and add 10 &#181;L of the final mix per well of liquid.</li>
	</ol>
	</li>
	<li>Seal the plate for sterile growth. Without touching the sticky side, carefully press the gas-permeable film across the plate. Ensure that each well has been individually sealed by applying pressure around each of the rings made by the wells. Replace the plate&#8217;s plastic lid snuggly over the plate and gas-permeable film (Figure 1, step 3b).</li>
	<li>Incubate the plates for 18 h in a plant growth chamber, under the same conditions as the seedlings were originally germinated, except on an orbital plate shaker set to 220 rpm.</li>
</ol>
4. Maintenance of bacterial colonization
<ol>
	<li>To rinse all floats (plants on mesh), add 1 mL of sterile water to wells of a new 24-well plate. Remove gas-permeable film. Using sterile forceps, transfer floats to wells with water (Figure 1, step 4a). Rinse by resting for 10 min at room temperature (RT) without agitation. 
	NOTE: To determine bacterial colonization efficiency of roots rather than their ability to maintain colonization over time, plants can be sacrificed at this step by taking them directly to step 5.1.</li>
	<li>Fill the wells of a new 24-well plate with 1 mL of plant growth medium. Transfer one mesh to each well. Cover with a gas-permeable seal and incubate for 72 h on the orbital plate shaker at 220 rpm in plant growth chamber (Figure 1, step 4b).</li>
	<li>Repeat the rinsing as performed in step 4.1 with floats after the 72 h incubation period.</li>
</ol>
5. Collection of bacteria for viable cell counts
NOTE: The number of bacteria per seedling root can be determined at any incubation timepoint. Colonization can be monitored between 0 h and 18 h, while maintenance can be monitored from 18 h onwards. Plants destined for imaging can proceed directly to section 6.
<ol>
	<li>Remove the seedlings from the mesh (Figure 1, step 5). Gently place flame-sterilized forceps below the leaves (but on the leaf side of the mesh), and lightly pinch the stem. Wiggle the seedlings up and away from the mesh to dislodge the root without breaking it. If the root breaks, gently scrape the mesh bottom to collect the full length.</li>
	<li>Remove bacteria from plant roots. Transfer the bacteria to wells of a 24-well plate containing 1 mL of ddH20. Sonicate the samples as described in step 1.3. 
	NOTE: Using a microscope, look for any remaining bacteria on the root surface on a sonicated sample. If bacteria remain, increase the total sonication time or intensity until no bacteria remain bound, up to the highest level of sonication that does not affect viable cell counts as determined in section 1.</li>
	<li>Quantify the bacteria on roots.
	<ol>
		<li>Perform serial 10-fold dilutions of the sonicated samples up to a 10-6 dilution in bacterial growth medium. Add 50 &#181;L of each dilution to individual agar plates and spread with sterile glass beads (or bacterial spreader). Incubate plates at the optimal temperature for bacteria until individual colonies are countable.</li>
		<li>Once distinguishable, count the number of each colony morphology (as determined in section 1), and calculate CFU of each bacterial species per seedling. Discard any samples showing contamination, as contamination during colonization or maintenance may affect bacterial presence.</li>
	</ol>
	</li>
</ol>
6. Collection of intact plant roots for microscopy
<ol>
	<li>Using forceps, remove the seedlings from mesh as in section 5.</li>
	<li>Transfer each plant to microscope slides.
	<ol>
		<li>Place the tip of the root on the slide and drag away from the tip to set the shoot down flush with the slide, ensuring a straightened root for best imaging. Add a drop of water or sterile plant growth medium to the samples to hydrate interfaces between the coverslips and slides.</li>
		<li>Place a glass coverslip just above the root crown (uppermost boxed region in Figure 1) and below the shoot leaves to avoid slanting of the coverslip (to allow for root crown imaging), and press down gently17.</li>
	</ol>
	</li>
	<li>If using fluorescent bacteria, image using appropriate excitation/emission filters to differentiate bacteria from each other and the plant root18.</li>
</ol>

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R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plant+growth-promoting+rhizobacteria+promote+plant+size+inequality.">Plant growth-promoting rhizobacteria promote plant size inequality.</a> Science Reports. 8 (1), 13828 (2018).</li><li>Levy, A., 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=Genomic+features+of+bacterial+adaptation+to+plants.">Genomic features of bacterial adaptation to plants.</a> Nature Genetics. 50 (1), 138-150 (2018).</li><li>Mart&#237;nez-Hidalgo, P., Maymon, M., Pule-Meulenberg, F., Hirsch, A. 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The authors have nothing to disclose.

Plants in all environments interact with thousands to millions of different bacteria and fungi5,7. These interactions can either negatively and positively impact plant health, with potential effects on crop yield and food production. Recent work also suggests that variable colonization of crops by PGPBs may account for unpredictable plant size and crop yield in field trials22. Understanding the mechanisms behind these interactions might allow us to directly manipulate plant-associated microbial communities to aid in healthy plant development, even under stress24.
Because bacterial colonization of roots and their maintenance in the rhizosphere is critical for plant-microbe interactions9, we wanted to build a system to reproducibly visualize and quantify these bacterial behaviors. This hydroponic, floating seedling growth system allows for microscopic imaging and quantification of bacterial populations on the roots of A. thaliana.
The described plant-microbe interaction assay integrates beneficial elements of existing experimental protocols. The floating mesh method was based on that from Haney et al.13, which measured initial colonization of P. simiae WCS417r on static, floating A. thaliana seedlings. In evaluating this system, strong colonization of A. thaliana roots by P. simiae was validated, even though a different growth medium from Haney et al. was utilized and included orbital shaking during colonization. The inclusion of orbital shaking during both colonization and maintenance facilitates bacterial interactions that might not occur in static culture, as well as reduce anoxic conditions that can inhibit both bacterial growth and plant root health13. We also integrated aspects of microbiology-focused protocols designed to support plant root colonization by various bacterial species8,15,25. This included a crucial transfer step out of the medium optimized for bacterial colonization and into a medium optimized for plant growth. This transfer to fresh medium also will allow the mechanisms underlying bacterial maintenance on roots to begin to be addressed, an approach that may provide insights into the erratic maintenance of PGPB in field trials6.
This assay was optimized for rapid processing of multiple samples to allow for environmental variables and mixed bacterial communities to be assayed within one multi-well biological replicate. While ultrasonication has been previously shown to be sufficient for disruption and collection of rhizosphere bacteria, the 24-well plate and multi-prong horn attachment quickens sample processing. This cell viability calculation approach to quantifying bacterial presence could be complemented by, or expanded to include, qPCR or MiSeq 16S rRNA gene community profiling to determine the relative abundance of more diverse communities of colonizing bacteria20. In addition, during imaging, the utility of focusing on just three regions of each plant root to speed the visualization of bacterial presence and localization on the roots is highlighted. The colonization of these different root regions has been shown to differ among bacterial species14. Imaging can be performed with either naturally autofluorescent bacteria or genetically tractable bacteria engineered to express a fluorescent protein.
The methodology described here allows for fast and reproducible evaluation of root colonization by PGPB bacteria, but there are limitations to the conclusions that can be drawn from these experiments. For instance, the ability for bacteria to chemotax towards the root is known to be important for many bacterial species&#8217; colonization of plant roots, but this process may not required within this shaking inoculation system. That said, for studies specifically interested in chemotaxis, the colonization step could be performed in static liquid culture or on the surface of a soft agar medium, where bacteria could be plated distant from the plant, requiring them to actively move towards the root. In addition, a relatively rich growth medium during colonization was used to promote bacterial growth and plant attachment; however, these comparatively high nutrient concentrations may prohibit the examination of bacterial utilization of or competition for plant-derived carbon during colonization. Again, depending on the growth requirements of the bacteria being studied, the colonization medium can be varied to best suit the particular research questions of specific researchers.
This system was designed to be easily amenable to different bacterial and plant growth conditions and to the addition of different environmental stressors and timepoints. However, the methods described here are best suited for measuring bacterial interactions with the roots of A. thaliana seedlings. Collection has been optimized for this plant, and larger or more sensitive plants may be intractable in the floating multi-well-plate system. Finally, while the bacteria of interest used here colonize the plant root in liquid culture, for other bacteria it may be necessary to inoculate the plant roots by dip-inoculation or plating on a solid agar medium instead19.

Crop destruction by bacterial and fungal diseases results in lowered food production and can severely disrupt global stability1. Based on the discovery that microbes in suppressive soils are responsible for increasing plant health2, scientists have asked whether the plant microbiome can be leveraged to support plant growth by modifying the presence and abundance of particular bacterial species3. Bacteria found to aid in plant growth or development are collectively termed plant growth-promoting bacteria (PGPB). More recently, studies have shifted from simply identifying potential PGPB to understanding how interkingdom interactions in the soil, around roots, or in the rhizosphere (the area directly surrounding and including the root surface) may be impacting PGPB activity4.
Rhizosphere colonization by PGPB can increase the health or the development of host plants in response to diverse stressors relative to uncolonized plants5. However, results are often more variable in native soil conditions compared to those observed in the closely controlled greenhouse and laboratory settings6. One hypothesis for this difference is that the growth or behavior of PGPB may be inhibited by native soil bacteria or fungi in the fields7,8. Beneficial effects by rhizosphere bacteria generally depend on the ability of the bacteria to 1) locate and move towards the root, 2) colonize the root through biofilm formation, and 3) interact with the host plant or pathogens via production of small molecule metabolites7,9. Any of these colonization behaviors may be affected by the presence and activity of neighboring microbes10.
We designed a system to quantify and visualize these distinct bacterial colonization stages of the rhizosphere (Figure 1). This approach will facilitate studies investigating why long-term PGPB maintenance is sometimes not observed following transfer of plants into new environments, such as during the planting of pre-inoculated seedlings. Arabidopsis thaliana as were chosen as a plant model due to its extensive use in laboratory studies as well as the ample data available about its microbial interactions11. There are three stages in the system: 1) A. thaliana growth, 2) bacterial colonization, and 3) bacterial maintenance (see Figure 1). Because A. thaliana is a terrestrial plant, it was important to ensure that it was not suffering undue water stress in the hydroponic system12. Inspired by the methods used by Haney et al.13, the seedlings are grown on plastic mesh to separate the shoot from the liquid growth medium. This system does not appear to compromise the health and development of the plant host, and it improves A. thaliana growth in liquid11. As the plant shoot floats above the surface, the roots are fully exposed to colonization by bacteria inoculated into the liquid bacterial growth medium. This permits bacteria of interest to be examined for colonization in nutrients that are most conducive to growth, while then shifting conditions to allow the plant to continue growing in a nutrient medium designed to support its growth. Both stages include steady shaking to prevent anoxia of the root13. Bacteria can be visualized or quantified from the plant roots following transfer from either the colonization medium or the maintenance medium. This hydroponic system is very flexible, allowing experimental conditions and applied stresses to be easily altered depending on interests of the researchers.
This described method is important in the context of the larger body of literature about plant-microbe interactions because it provides a robust system for studying these interactions at the root surface while also being customizable to the growth preferences of different bacteria. Plant biology labs often perform plant-microbe colonization experiments on solid agar, allowing for only planar movement (if that) of bacteria while also requiring the potentially destructive manipulation of plants during subsequent transfer. In contrast, microbiology labs have frequently prioritized the health of the bacteria within their experiments, to the detriment of the plants14,15. These different priorities of plant- and microbiology-focused labs have historically made it difficult to compare results between these groups, since each typically optimizes experimental conditions to optimize their organism of interest15.&#160;The floating-mesh-plant-growth system described here prevents full plant submersion, a notable advantage to previous microbiology-oriented studies, while also temporarily optimizing the growth and survival of bacteria to facilitate colonization. Thus, the assay we present here may address concerns from both plant biologists (about over-hydration and tactile manipulation of the plant) while satisfying the criteria of microbiologists (allowing for different bacterial growth conditions and multiple species&#8217; interactions)7. This protocol is designed to be adaptable for use with various bacteria, plants, and environmental conditions.

<table>
	<tbody>
		<tr>
			<td>Required Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL eppendorf tubes</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>24-well plates</td>
			<td>BD Falcon</td>
			<td>1801343</td>
			<td></td>
		</tr>
		<tr>
			<td>Aeraseal</td>
			<td>Excel Scientific</td>
			<td>BE255A2</td>
			<td></td>
		</tr>
		<tr>
			<td>Autoclave</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Bacteria of Interest</td>
			<td>any</td>
			<td>N/A</td>
			<td>Stored at -80&#730;C in 40% glycerol preferred</td>
		</tr>
		<tr>
			<td>BactoAgar</td>
			<td>BD</td>
			<td>2306428; REF 214010</td>
			<td></td>
		</tr>
		<tr>
			<td>bleach</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Conviron</td>
			<td>any</td>
			<td>N/A</td>
			<td>Short Day Light-Dark Cycles: 460-600 &#181;moles/m&#178;/s set at 9/15 hours light/dark at 18/21&#730;C, with inner power outlet</td>
		</tr>
		<tr>
			<td>Dessicator Jar: glass or heavy plastic</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Flame</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>any</td>
			<td>N/A</td>
			<td>At optimal temperature for growth of specified bacteria</td>
		</tr>
		<tr>
			<td>Hydrochloric Acid</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Lennox LB Broth</td>
			<td>RPI</td>
			<td>L24066-1000.0</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipetters</td>
			<td>any</td>
			<td>N/A</td>
			<td>Volumes 5 &#181;L to 1000 &#181;L</td>
		</tr>
		<tr>
			<td>Microscope (preferably fluorescence)</td>
			<td>any</td>
			<td>N/A</td>
			<td>Could be light if best definition not important</td>
		</tr>
		<tr>
			<td>MS Salts + MES</td>
			<td>RPI</td>
			<td>M70300-50.0</td>
			<td></td>
		</tr>
		<tr>
			<td>Orbital Plate Shaker</td>
			<td>any</td>
			<td>N/A</td>
			<td>Capable of running at 220 rpm for at least 96 hours</td>
		</tr>
		<tr>
			<td>Petri Dishes</td>
			<td>any</td>
			<td>N/A</td>
			<td>50 mL total volume</td>
		</tr>
		<tr>
			<td>Reservoirs</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectrophotometer</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard Hole Punch</td>
			<td>any</td>
			<td>N/A</td>
			<td>Approximately 7mm punch diameter</td>
		</tr>
		<tr>
			<td>Sterile water</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Tape</td>
			<td>3M</td>
			<td>MMM1538-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Teflon Mesh</td>
			<td>McMaster-Carr</td>
			<td>1100t41</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonicator</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex Mixer</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>X-gal</td>
			<td>GoldBio</td>
			<td>x4281c</td>
			<td>other vendors available</td>
		</tr>
		<tr>
			<td>Suggested Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>24 Prong Ultrasonicator attachment</td>
			<td>any</td>
			<td>N/A</td>
			<td>For sonicating multiple samples at once. Can be done individually</td>
		</tr>
		<tr>
			<td>Alumaseal II</td>
			<td>Excel Scientific</td>
			<td>FE124F</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass beads</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Multipetter/Repetter</td>
			<td>any</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile 96-well plates</td>
			<td>any</td>
			<td>N/A</td>
			<td>For serial dilutions. Can be replaced by eppendorf tubes</td>
		</tr>
		<tr>
			<td>Biological Materials Used</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Arabidopsis thaliana seeds</td>
			<td>any</td>
			<td>N/A</td>
			<td>We recommend Arabidopsis Biological Resource Center for seed stocks</td>
		</tr>
		<tr>
			<td>Arthrobacter nicotinovorans</td>
			<td></td>
			<td></td>
			<td>Levy, et al. 2018</td>
		</tr>
		<tr>
			<td>Curtobacterium oceanosedimentum</td>
			<td></td>
			<td></td>
			<td>Levy, et al. 2018</td>
		</tr>
		<tr>
			<td>Microbacterium oleivorans</td>
			<td></td>
			<td></td>
			<td>Levy, et al. 2018</td>
		</tr>
		<tr>
			<td>Pseudomonas simiae WCS417r</td>
			<td></td>
			<td></td>
			<td>Published in a similar system in Haney, et al. 2015. Strain used developed in Cole, et al. 2017</td>
		</tr>
	</tbody>
</table>

monitoring bacterial colonization and maintenance on arabidopsis thaliana roots in a floating hydroponic system

Bacteria form complex rhizosphere microbiomes shaped by interacting microbes, larger organisms, and the abiotic environment. Under laboratory conditions, rhizosphere colonization by plant growth-promoting bacteria (PGPB) can increase the health or the development of host plants relative to uncolonized plants. However, in field settings, bacterial treatments with PGPB often do not provide substantial benefits to crops. One explanation is that this may be due to loss of the PGPB during interactions with endogenous soil microbes over the lifespan of the plant. This possibility has been difficult to confirm, since most studies focus on the initial colonization rather than maintenance of PGPB within rhizosphere communities. It is hypothesized here that the assembly, coexistence, and maintenance of bacterial communities are shaped by deterministic features of the rhizosphere microenvironment, and that these interactions may impact PGPB survival in native settings. To study these behaviors, a hydroponic plant-growth assay is optimized using Arabidopsis thaliana to quantify and visualize the spatial distribution of bacteria during initial colonization of plant roots and after transfer to different growth environments. This system&#8217;s reproducibility and utility are then validated with the well-studied PGPB Pseudomonas simiae. To investigate how the presence of multiple bacterial species may affect colonization and maintenance dynamics on the plant root, a model community from three bacterial strains (an Arthrobacter, Curtobacterium, and Microbacterium species) originally isolated from the A. thaliana rhizosphere is constructed. It is shown that the presence of these diverse bacterial species can be measured using this hydroponic plant-maintanence assay, which provides an alternative to sequencing-based bacterial community studies. Future studies using this system may improve the understanding of bacterial behavior in multispecies plant microbiomes over time and in changing environmental conditions.

The well-characterized PGPB P. simiae WCS417r is known to colonize the roots of A. thaliana in hydroponic culture. This naturally fluorescent bacterium can easily be visualized using microscopy on the roots of seedlings following colonization (Figure 2). Although it is possible to image the full length of these A. thaliana seedlings&#8217; (4&#8211;6 mm length) roots, doing so for many plants would take a prohibitive amount of time. Because most variation across timepoints and species of bacteria can be captured by imaging the crown, middle, and tip of the root14 (indicated by red boxes in Figure 1), these regions were prioritized for imaging rather than imaging full root lengths. In the bright-field images of P. simiae-colonized A. thaliana roots (Figure 2), it is possible to visualize the outline of the roots and root hairs; however, at 18 h of colonization, it is not possible to clearly differentiate colonized versus non-colonized roots using bright-field images. While P. simiae displays autofluorescence, we used a strain also engineered to express a yellow fluorescent protein (YFP)19 with excitation/emission wavelengths of 490-510/520-550 nm18. A magnification of 100x was sufficient to clearly identify individual and small aggregates of P. simiae cells on A. thaliana roots. As shown in Figure 2, laboratories with access to either high-resolution confocal microscopes or less expensive benchtop microscopes can both visualize the presence and distribution of bacteria along the root.
While informative in terms of spatial distribution, microscopy images are not well-suited for quantification of bacterial cells. We thus collected bacteria from the surface of roots using ultrasonication as previously described and validated9,20. Three rounds of 12 s of ultrasonication21 at an amplitude of 40 were sufficient to disrupt the outer surface of the root seedlings (Supplemental Figure 1) and remove all bacteria while not impacting the bacterial viability. Sonication was used rather than bead-beating methods9 to better promote dispersal of bacterial aggregates/biofilms. Quantifying CFU/root after 18 h of colonization and an additional 72 h of maintenance showed that P. simiae both colonizes and is maintained on the roots of A. thaliana in our hydroponic, floating seedling system (Figure 3). The number of CFU/seedling at either timepoint showed good reproducibility across biological replicates performed on different days (Figure 3). The variation observed is common among root colonization assays22 and is likely due to minor variations of timing, environmental conditions, or plant root size, even among seedlings germinated at the same time and selected to be similar in size. We observe an increase in the number of CFU/seedling after 72 h in maintenance medium as compared to the numbers observed at the post-colonization 18 h timepoint (Figure 3). This indicates active growth of the colonized bacteria on the plant root occurred during the maintenance stage.
In addition to the utility of this hydroponic assay to quantify individual bacterial colonization and maintenance, it is also applicable to monitoring the association of multiple species on plant roots. To demonstrate this, three species of bacteria isolated from A. thaliana grown in natural soil under laboratory conditions were chosen20. The isolates were strains of Arthrobacter nicotinovorans, Microbacterium oleivorans, and Curtobacterium oceanosedimentum23. This simplified community was chosen due to these species&#8217; ability to coexist in liquid bacterial growth media in shaking culture (unpublished data). In addition, these three species can be clearly differentiated on media containing X-gal due to differences in colony morphology and color (Figure 4A). The X-gal does not affect relative growth of any of these bacterial species (unpublished data). These differences in morphology and colony appearance on X-gal allowed us to count the CFU/seedling of each species without antibiotic selection, even in multi-species coculture.
A. nicotinovorans, M. oleivorans, and C. oceanosedimentum were all colonized and maintained on the root, whether alone or in bacterial coculture (Figure 4B). Each species showed trends that were similar across different biological and technical replicates, even within mixed communities. This demonstrates that the assay protocol can be used to measure both relative or total CFU/root of each species. Interestingly, when grown alone, no individual species showed an appreciable increase in abundance during the maintenance stage, but the overall CFU/root of the combined community increased in cocultures, indicating that these bacteria do not prohibit the colonization of the other strains.
For all experiments, plants grown in liquid media without the addition of bacteria as negative controls were always included. No bacteria were visible on these control roots during microscopy (Figure 2), nor were any bacteria detected via plating for CFU (unpublished data). This indicates that sterilization of seeds and using the sterile techniques during this assay were sufficient to keep plants axenic unless purposely colonized.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59517/59517fig1.jpg" /> 
Figure 1: Assay for bacterial colonization and maintenance on A. thaliana roots. A. thaliana seedlings grown on sterilized plastic mesh were transferred to a growth medium optimized for bacteria [here, 0.1 x LB (Luria Broth) Lennox]. Bacteria then colonized the root over 18 h while the plant floated in shaking liquid. Following a rinse, the colonized float was transferred to a growth medium optimized for plants (0.5x MS + MES) for 72 h to test for maintenance of bacteria on the roots. The float was then rinsed, and the plant with any attached microbes was removed for analysis (quantification of CFU/seedling or imaging by microscopy). <a href="https://www.jove.com/files/ftp_upload/59517/59517fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59517/59517fig2.jpg" /> 
Figure 2: Visualizing P. simiae colonization of roots with fluorescent microscopy. P. simiae (false-colored green) colonized A. thaliana roots and was maintained on the root following transfer to plant-growth medium. Root crown (left), mid-length (center), and tip (right) at 40x magnification are shown from areas indicated in Figure 1. The top two rows show the bright-field and fluorescent images of roots colonized by P. simiae (imaged by epifluorescent microscopy). The same roots were also imaged by a confocal microscope (center two rows). The no-bacteria negative control in the two bottom rows showed no colonization. Scale bars represent 50 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59517/59517fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59517/59517fig3.jpg" /> 
Figure 3: Quantification of P. simiae on A. thaliana roots. Total number of P. simiae viable cells recovered per A. thaliana seedling following 18 h of colonization or 72 h of maintenance. Three individual biological replicates are shown, each containing three technical replicates of two seedlings per float. The numbers shown are the means from the technical replicates, while bars represent standard errors of the means. <a href="https://www.jove.com/files/ftp_upload/59517/59517fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59517/59517fig4v2.jpg" /> 
Figure 4: Quantification of the colonization and maintenance of a mixed bacterial community on A. thaliana roots. (A) Colonies of A. nicotinovorans, M. oleivorans, and C. oceanosedimentum can be differentiated on X-gal-containing agar medium by colony morphology and color. (B) Roots of 10 day-old seedlings were inoculated with approximately 3x105 CFU/mL of each of the three strains. Shown are total CFU/seedling recovered of each species following 18 h of colonization or 72 h of maintenance when colonized either alone or in a three-member bacterial community. Two biological replicates, each comprising two technical replicates of two seedlings per float, are shown. The numbers shown are the means from the two technical replicates, while bars represent standard errors of the means . <a href="https://www.jove.com/files/ftp_upload/59517/59517fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure 1: Ultrasonication disrupts the root surface. To dislodge bacteria from the surface of the root, whole plants were sonicated, and the bacteria was released into the liquid, which was serially diluted and plated for quantification of CFU/seedling. (A) An intact seedling is (B) structurally disrupted following ultrasonication. <a href="https://www.jove.com/files/ftp_upload/59517/Supplemental_Figure.eps">Please click here to download this file.</a>

Watch this Scientific Journal Video about Monitoring Bacterial Colonization and Maintenance on Arabidopsis thaliana Roots in a Floating Hydroponic System at JoVE.com

Monitoring Bacterial Colonization and Maintenance on Arabidopsis thaliana Roots in a Floating Hydroponic System

Here we describe a hydroponic plant growth assay to quantify species presence and visualize the spatial distribution of bacteria during initial colonization of plant roots and after their transfer into different growth environments.

Monitoring Bacterial Colonization and Maintenance on Arabidopsis thaliana Roots in a Floating Hydroponic System

Bacteria form complex rhizosphere microbiomes shaped by interacting microbes, larger organisms, and the abiotic environment. Under laboratory conditions, ...

monitoring-bacterial-colonization-maintenance-on-arabidopsis-thaliana

Universidad San Sebastian

Research

JoVE Journal

Immunology and Infection

10.4K Views.  University of North Carolina at Chapel Hill. Here we describe a hydroponic plant growth assay to quantify species presence and visualize the spatial distribution of bacteria during initial colonization of plant roots and after their transfer into different growth environments.

Video: Monitoring Bacterial Colonization and Maintenance on Arabidopsis thaliana Roots in a Floating Hydroponic System

Quantification of Fungal Colonization, Sporogenesis, and Production of Mycotoxins Using Kernel Bioassays

The rotting of grains by seed-infecting fungi poses one of the greatest economic challenges to cereal production worldwide, not to mention serious risks to human and animal health. Among cereal production, maize is arguably the most affected crop, due to pathogen-induced losses in grain integrity and mycotoxin seed contamination. The two most prevalent and problematic mycotoxins for maize growers and food and feed processors are aflatoxin and fumonisin, produced by Aspergillus flavus and Fusarium verticillioides, respectively.
Recent studies in molecular plant-pathogen interactions have demonstrated promise in understanding specific mechanisms associated with plant responses to fungal infection and mycotoxin contamination1,2,3,4,5,6. Because many labs are using kernel assays to study plant-pathogen interactions, there is a need for a standardized method for quantifying different biological parameters, so results from different laboratories can be cross-interpreted. For a robust and reproducible means for quantitative analyses on seeds, we have developed in-lab kernel assays and subsequent methods to quantify fungal growth, biomass, and mycotoxin contamination. Four sterilized maize kernels are inoculated in glass vials with a fungal suspension (106) and incubated for a predetermined period. Sample vials are then selected for enumeration of conidia by hemocytometer, ergosterol-based biomass analysis by high performance liquid chromatography (HPLC), aflatoxin quantification using an AflaTest fluorometer method, and fumonisin quantification by HPLC.

The devastation of cereal crops by seed-infecting fungi has prompted numerous research efforts to better understand plant-pathogen interactions. To study seed-fungal interactions in a laboratory setting, we developed a robust method for the quantification of fungal reproduction, biomass, and mycotoxin contamination using kernel bioassays.

The rotting of grains by seed-infecting fungi poses one of the greatest economic challenges to cereal production worldwide, not to mention serious risks ...

Investigating the Effects of Probiotics on Pneumococcal Colonization Using an In Vitro Adherence Assay

Adherence of Streptococcus pneumoniae (the pneumococcus) to the epithelial lining of the nasopharynx can result in colonization and is considered a prerequisite for pneumococcal infections such as pneumonia and otitis media. In vitro adherence assays can be used to study the attachment of pneumococci to epithelial cell monolayers and to investigate potential interventions, such as the use of probiotics, to inhibit pneumococcal&#160;colonization.&#160;The protocol described here is used to investigate the effects of the probiotic Streptococcus salivarius on the adherence of pneumococci to the human epithelial cell line CCL-23 (sometimes referred to as HEp-2 cells). The assay involves three main steps: 1) preparation of epithelial and bacterial cells, 2) addition of bacteria to epithelial cell monolayers, and 3) detection of adherent pneumococci by viable counts (serial dilution and plating) or quantitative real-time PCR (qPCR). This technique is relatively straightforward and does not require specialized equipment other than a tissue culture setup. The assay can be used to test other probiotic species and/or potential inhibitors of pneumococcal colonization and can be easily modified to address other scientific questions regarding pneumococcal adherence and invasion.

Investigating the Effects of Probiotics on Pneumococcal Colonization Using an In Vitro Adherence Assay

In vitro adherence assays can be used to study the attachment of Streptococcus pneumoniae to epithelial cell monolayers and to investigate potential interventions such as the use of probiotics for inhibiting pneumococcal colonization.

Adherence of Streptococcus pneumoniae (the pneumococcus) to the epithelial lining of the nasopharynx can result in colonization and is considered a ...

Monitoring the Assembly of a Secreted Bacterial Virulence Factor Using Site-specific Crosslinking

This article describes a method to detect and analyze dynamic interactions between a protein of interest and other factors in vivo. Our method is based on the amber suppression technology that was originally developed by Peter Schultz and colleagues1. An amber mutation is first introduced at a specific codon of the gene encoding the protein of interest. The amber mutant is then expressed in E. coli together with genes encoding an amber suppressor tRNA and an amino acyl-tRNA synthetase derived from Methanococcus jannaschii. Using this system, the photo activatable amino acid analog p-benzoylphenylalanine (Bpa) is incorporated at the amber codon. Cells are then irradiated with ultraviolet light to covalently link the Bpa residue to proteins that are located within 3-8 &#197;. Photocrosslinking is performed in combination with pulse-chase labeling and immunoprecipitation of the protein of interest in order to monitor changes in protein-protein interactions that occur over a time scale of seconds to minutes. We optimized the procedure to study the assembly of a bacterial virulence factor that consists of two independent domains, a domain that is integrated into the outer membrane and a domain that is translocated into the extracellular space, but the method can be used to study many different assembly processes and biological pathways in both prokaryotic and eukaryotic cells. In principle interacting factors and even specific residues of interacting factors that bind to a protein of interest can be identified by mass spectrometry.

This article illustrates the use of pulse-chase radio labeling in combination with site-specific photocrosslinking to monitor interactions between a protein of interest and other factors in E. coli. Unlike traditional chemical cross-linking methods, this approach generates high resolution &#8220;snapshots&#8221; of an ordered assembly pathway in a living cell.

This article describes a method to detect and analyze dynamic interactions between a protein of interest and other factors in vivo. Our method is based on ...

Assessing Bacterial Invasion of Cardiac Cells in Culture and Heart Colonization in Infected Mice Using Listeria monocytogenes

Listeria monocytogenes is a Gram-positive facultative intracellular pathogen that is capable of causing serious invasive infections in immunocompromised patients, the elderly, and pregnant women. The most common manifestations of listeriosis in humans include meningitis, encephalitis, and fetal abortion. A significant but much less documented sequelae of invasive L. monocytogenes infection involves the heart. The death rate from cardiac illness can be up to 35% despite treatment, however very little is known regarding L. monocytogenes colonization of cardiac tissue and its resultant pathologies. In addition, it has recently become apparent that subpopulations of L. monocytogenes have an enhanced capacity to invade and grow within cardiac tissue. This protocol describes in detail in vitro and in vivo methods that can be used for assessing cardiotropism of L. monocytogenes isolates. Methods are presented for the infection of H9c2 rat cardiac myoblasts in tissue culture as well as for the determination of bacterial colonization of the hearts of infected mice. These methods are useful not only for identifying strains with the potential to colonize cardiac tissue in infected animals, but may also facilitate the identification of bacterial gene products that serve to enhance cardiac cell invasion and/or drive changes in heart pathology. These methods also provide for the direct comparison of cardiotropism between multiple L. monocytogenes strains.

Assessing Bacterial Invasion of Cardiac Cells in Culture and Heart Colonization in Infected Mice Using Listeria monocytogenes

Listeria monocytogenes causes fetal infections in pregnant women and meningitis in susceptible populations. Subpopulations of bacteria can colonize cardiac tissue, causing myocarditis in patients and laboratory animals. Here we present a protocol that describes how to assess L. monocytogenes cardiac cell invasion in vitro and cardiac colonization in infected animals.

Listeria monocytogenes is a Gram-positive facultative intracellular pathogen that is capable of causing serious invasive infections in immunocompromised ...

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence factors, which can subvert and manipulate key host functions. The study of host/pathogen interactions is therefore extremely important to understand bacterial infections and develop alternative strategies to counter infectious diseases. This approach however, requires the development of new high-throughput assays for the unbiased, automated identification and characterization of bacterial virulence determinants. Here, we describe a method for the generation of a GFP-tagged mutant library by transposon mutagenesis and the development of high-content screening approaches for the simultaneous identification of multiple transposon-associated phenotypes. Our working model is the intracellular bacterial pathogen Coxiellaburnetii, the etiological agent of the zoonosis Q fever, which is associated with severe outbreaks with a consequent health and economic burden. The obligate intracellular nature of this pathogen has, until recently, severely hampered the identification of bacterial factors involved in host pathogen interactions, making of Coxiella the ideal model for the implementation of high-throughput/high-content approaches.

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Coxiella burnetii is an obligate intracellular Gram-negative bacterium responsible for the zoonotic disease Q fever. Here we describe methods for the generation of Coxiella fluorescent transposon mutants as well as the automated identification and analysis of the resulting internalization, replication and cytotoxic phenotypes.

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence ...

Bacterial Leaf Infiltration Assay for Fine Characterization of Plant Defense Responses using the Arabidopsis thaliana-Pseudomonas syringae Pathosystem

In the absence of specialized mobile immune cells, plants utilize their localized programmed cell death and Systemic Acquired Resistance to defend themselves against pathogen attack. The contribution of a specific Arabidopsis gene to the overall plant immune response can be specifically and quantitatively assessed by assaying the pathogen growth within the infected tissue. For over three decades, the hemibiotrophic bacterium Pseudomonas syringae pv. maculicola ES4326 (Psm ES4326) has been widely applied as the model pathogen to investigate the molecular mechanisms underlying the Arabidopsis immune response. To deliver pathogens into the leaf tissue, multiple inoculation methods have been established, e.g., syringe infiltration, dip inoculation, spray, vacuum infiltration, and flood inoculation. The following protocol describes an optimized syringe infiltration method to deliver virulent Psm ES4326 into leaves of adult soil-grown Arabidopsis plants and accurately screen for enhanced disease susceptibility (EDS) towards this pathogen. In addition, this protocol can be supplemented with multiple pre-treatments to further dissect specific immune defects within different layers of plant defense, including Salicylic Acid (SA)-Triggered Immunity (STI) and MAMP-Triggered Immunity (MTI).

Bacterial Leaf Infiltration Assay for Fine Characterization of Plant Defense Responses using the Arabidopsis thaliana-Pseudomonas syringae Pathosystem

Quantification of pathogen growth is a powerful tool to characterize various Arabidopsis thaliana (hereafter: Arabidopsis) immune responses. The method described here presents an optimized syringe infiltration assay to quantify the Pseudomonas syringae pv. maculicola ES4326 growth in adult Arabidopsis leaves.

In the absence of specialized mobile immune cells, plants utilize their localized programmed cell death and Systemic Acquired Resistance to defend ...

Implementation of a Permeable Membrane Insert-based Infection System to Study the Effects of Secreted Bacterial Toxins on Mammalian Host Cells

Many bacterial pathogens secrete potent toxins to aid in the destruction of host tissue, to initiate signaling changes in host cells or to manipulate immune system responses during the course of infection. Though methods have been developed to successfully purify and produce many of these important virulence factors, there are still many bacterial toxins whose unique structure or extensive post-translational modifications make them difficult to purify and study in in vitro systems. Furthermore, even when pure toxin can be obtained, there are many challenges associated with studying the specific effects of a toxin under relevant physiological conditions. Most in vitro cell culture models designed to assess the effects of secreted bacterial toxins on host cells involve incubating host cells with a one-time dose of toxin. Such methods poorly approximate what host cells actually experience during an infection, where toxin is continually produced by bacterial cells and allowed to accumulate gradually during the course of infection. This protocol describes the design of a permeable membrane insert-based bacterial infection system to study the effects of Streptolysin S, a potent toxin produced by Group A Streptococcus, on human epithelial keratinocytes. This system more closely mimics the natural physiological environment during an infection than methods where pure toxin or bacterial supernatants are directly applied to host cells. Importantly, this method also eliminates the bias of host responses that are due to direct contact between the bacteria and host cells. This system has been utilized to effectively assess the effects of Streptolysin S (SLS) on host membrane integrity, cellular viability, and cellular signaling responses. This technique can be readily applied to the study of other secreted virulence factors on a variety of mammalian host cell types to investigate the specific role of a secreted bacterial factor during the course of infection.

Here, a method using a permeable membrane insert-based infection system to study the effects of Streptolysin S, a secreted toxin produced by Group A Streptococcus, on keratinocytes is described. This system can be readily applied to the study of other secreted bacterial proteins on various host cell types during infection.

Many bacterial pathogens secrete potent toxins to aid in the destruction of host tissue, to initiate signaling changes in host cells or to manipulate ...

A Murine Model of Group B Streptococcus Vaginal Colonization

Streptococcus agalactiae (group B Streptococcus, GBS), is a Gram-positive, asymptomatic colonizer of the human gastrointestinal tract and vaginal tract of 10 - 30% of adults. In immune-compromised individuals, including neonates, pregnant women, and the elderly, GBS may switch to an invasive pathogen causing sepsis, arthritis, pneumonia, and meningitis. Because GBS is a leading bacterial pathogen of neonates, current prophylaxis is comprised of late gestation screening for GBS vaginal colonization and subsequent peripartum antibiotic treatment of GBS-positive mothers. Heavy GBS vaginal burden is a risk factor for both neonatal disease and colonization. Unfortunately, little is known about the host and bacterial factors that promote or permit GBS vaginal colonization. This protocol describes a technique for establishing persistent GBS vaginal colonization using a single &#946;-estradiol pre-treatment and daily sampling to determine bacterial load. It further details methods to administer additional therapies or reagents of interest and to collect vaginal lavage fluid and reproductive tract tissues. This mouse model will further the understanding of the GBS-host interaction within the vaginal environment, which will lead to potential therapeutic targets to control maternal vaginal colonization during pregnancy and to prevent transmission to the vulnerable newborn. It will also be of interest to increase our understanding of general bacterial-host interactions in the female vaginal tract.

A Murine Model of Group B Streptococcus Vaginal Colonization

The purpose of this protocol is to imitate human group B Streptococcus (GBS) vaginal colonization in a murine model. This method may be used to investigate host immune responses and bacterial factors contributing to GBS vaginal persistence, as well as to test therapeutic strategies.

Streptococcus agalactiae (group B Streptococcus, GBS), is a Gram-positive, asymptomatic colonizer of the human gastrointestinal tract and vaginal tract of ...

Detection of Enterohemorrhagic Escherichia Coli Colonization in Murine Host by Non-invasive In Vivo Bioluminescence System

Enterohemorrhagic E. coli (EHEC) O157:H7, which is a foodborne pathogen that causesdiarrhea, hemorrhagic colitis (HS), and hemolytic uremic syndrome (HUS), colonize to the intestinal tract of humans. To study the detailed mechanism of EHEC colonization in vivo, it is essential to have animal models to monitor and quantify EHEC colonization. We demonstrate here a mouse-EHEC colonization model by transforming the bioluminescent expressing plasmid to EHEC to monitor and quantify EHEC colonization in living hosts. Animals inoculated with bioluminescence-labeled EHEC show intense bioluminescent signals in mice by detection with a non-invasive in vivo imaging system. After 1 and 2 days post infection, bioluminescent signals could still be detected in infected animals, which suggests that EHEC colonize in hosts for at least 2 days. We also demonstrate that these bioluminescent EHEC locate to mouse intestine, specifically in the cecum and colon, from ex vivo images. This mouse-EHEC colonization model may serve as a tool to advance the current knowledge of the EHEC colonization mechanism.

Detection of Enterohemorrhagic Escherichia Coli Colonization in Murine Host by Non-invasive In Vivo Bioluminescence System

A detailed protocol of a mouse model for enterohemorrhagic E. coli (EHEC) colonization by using bioluminescence-labeled bacteria is presented. The detection of these bioluminescent bacteria by a non-invasive in vivo imaging system in live animals can advance our current understanding of EHEC colonization.

Enterohemorrhagic E. coli (EHEC) O157:H7, which is a foodborne pathogen that causesdiarrhea, hemorrhagic colitis (HS), and hemolytic uremic syndrome ...

A High-throughput Shigella-specific Bactericidal Assay

Serum bactericidal assays (SBAs) measure the functional activity of antibodies and have been used for many decades. SBAs directly measure antibody killing activity by assessing the ability of antibodies in serum to bind to bacteria and activate complement. This complement activation results in the lysis and killing of the target bacteria. These assays are valuable because they go beyond quantifying antibody production to elucidate the biological functions that these antibodies have, allowing researchers to study the role that antibodies may play in preventing infection. SBAs have been used to study immune responses for many human pathogens, but there is no widely accepted methodology for Shigella at present. Historically, SBAs have been very labor-intensive, requiring many time-consuming steps to accurately quantify surviving bacteria. This protocol describes a simple, robust, and high-throughput assay that measures functional antibodies specific for Shigella in serum in vitro. The method described here offers many advantages over traditional SBAs, including the use of frozen bacterial stocks, 96 well assay plates, a micro-culture system, and automated colony-counting. All of these modifications make this assay less labor-intensive and more high-throughput. This protocol is simpler and faster to perform than traditional SBAs while still using simple technologies and readily available reagents. The protocol has been successfully applied in multiple independent laboratories and the assay is robust and reproducible. The assay can be used to assess immune responses in pre-clinical as well as clinical studies. Quantifying shigellacidal antibody titers both before and after antigen exposure (either by immunization or infection) allows for a broader understanding of how functional antibody responses are generated and their contribution to protective immunity. The development of this standardized, well-characterized assay may greatly facilitate Shigella vaccine design.

A High-throughput Shigella-specific Bactericidal Assay

Here we present a protocol to measure Shigellacidal activity of antibodies in serum. Serum is mixed with bacteria and exogenous complement, incubated, and the reaction mixture is plated on agar plates. Viable bacteria form colonies which are counted, using an automated colony enumerator, and used to determine the bactericidal titer.

Serum bactericidal assays (SBAs) measure the functional activity of antibodies and have been used for many decades. SBAs directly measure antibody killing ...