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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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 al...

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>&lt;strong&gt;Required Materials&lt;/strong&gt;</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?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 &amp;micro;moles/m&amp;sup2;/s set at 9/15 hours light/dark at 18/21?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 &amp;micro;L to 1000 &amp;micro;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>&lt;strong&gt;Suggested Materials&lt;/strong&gt;</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>&lt;strong&gt;Biological Materials Used&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;em&gt;Arabidopsis thaliana&lt;/em&gt; seeds</td><td>any</td><td>N/A</td><td>We recommend Arabidopsis Biological Resource Center for seed stocks</td></tr><tr><td>&lt;em&gt;Arthrobacter nicotinovorans&lt;/em&gt;</td><td></td><td></td><td>Levy, &lt;em&gt;et al. &lt;/em&gt;2018</td></tr><tr><td>&lt;em&gt;Curtobacterium oceanosedimentum&lt;/em&gt;</td><td></td><td></td><td>Levy, &lt;em&gt;et al. &lt;/em&gt;2018</td></tr><tr><td>&lt;em&gt;Microbacterium oleivorans&lt;/em&gt;</td><td></td><td></td><td>Levy, &lt;em&gt;et al. &lt;/em&gt;2018</td></tr><tr><td>&lt;em&gt;Pseudomonas simiae &lt;/em&gt;WCS417r</td><td></td><td></td><td>Published in a similar system in Haney, &lt;em&gt;et al.&lt;/em&gt; 2015. Strain used developed in Cole, &lt;em&gt;et al. 2017&lt;/em&gt;</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....

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

Research

JoVE Journal

Immunology and Infection

10.5K 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

An Orthotopic Model of Serous Ovarian Cancer in Immunocompetent Mice for in vivo Tumor Imaging and Monitoring of Tumor Immune Responses

Background: Ovarian cancer is generally diagnosed at an advanced stage where the case/fatality ratio is high and thus remains the most lethal of all gynecologic malignancies among US women 1,2,3. Serous tumors are the most widespread forms of ovarian cancer and 4,5 the Tg-MISIIR-TAg transgenic represents the only mouse model that spontaneously develops this type of tumors. Tg-MISIIR-TAg mice express SV40 transforming region under control of the Mullerian Inhibitory Substance type II Receptor (MISIIR) gene promoter 6. Additional transgenic lines have been identified that express the SV40 TAg transgene, but do not develop ovarian tumors. Non-tumor prone mice exhibit typical lifespan for C57Bl/6 mice and are fertile. These mice can be used as syngeneic allograft recipients for tumor cells isolated from Tg-MISIIR-TAg-DR26 mice.
Objective: Although tumor imaging is possible 7, early detection of deep tumors is challenging in small living animals. To enable preclinical studies in an immunologically intact animal model for serous ovarian cancer, we describe a syngeneic mouse model for this type of ovarian cancer that permits in vivo imaging, studies of the tumor microenvironment and tumor immune responses.
Methods: We first derived a TAg+ mouse cancer cell line (MOV1) from a spontaneous ovarian tumor harvested in a 26 week-old DR26 Tg-MISIIR-TAg female. Then, we stably transduced MOV1 cells with TurboFP635 Lentivirus mammalian vector that encodes Katushka, a far-red mutant of the red fluorescent protein from sea anemone Entacmaea quadricolor with excitation/emission maxima at 588/635 nm 8,9,10. We orthotopically implanted MOV1Kat in the ovary 11,12,13,14 of non-tumor prone Tg-MISIIR-TAg female mice. Tumor progression was followed by in vivo optical imaging and tumor microenvironment was analyzed by immunohistochemistry.
Results: Orthotopically implanted MOV1Kat cells developed serous ovarian tumors. MOV1Kat tumors could be visualized by in vivo imaging up to three weeks after implantation (fig. 1) and were infiltrated with leukocytes, as observed in human ovarian cancers 15 (fig. 2).
Conclusions: We describe an orthotopic model of ovarian cancer suitable for in vivo imaging of early tumors due to the high pH-stability and photostability of Katushka in deep tissues. We propose the use of this novel syngeneic model of serous ovarian cancer for in vivo imaging studies and monitoring of tumor immune responses and immunotherapies.

An Orthotopic Model of Serous Ovarian Cancer in Immunocompetent Mice for in vivo Tumor Imaging and Monitoring of Tumor Immune Responses

To study in vivo tumor growth and tumor microenvironment, we used a syngeneic and orthotopic mouse model of ovarian cancer in immunocompetent animals. We transduced a mouse tumor cell line (MOV1) with Katushka fluorescent protein (MOV1KAT) and here we show its orthotopic implantation in ovary and in vivo imaging.

Background: Ovarian cancer is generally diagnosed at an advanced stage where the case/fatality ratio is high and thus remains the most lethal of all ...

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, phosphorus, sulfate and trace elements that plants acquire from the soil. If we want to develop sustainable approaches to producing high crop yield, we need to better understand how the root develops, takes up a wide spectrum of nutrients, and interacts with symbiotic and pathogenic organisms. To accomplish these goals, we need to be able to explore roots in microscopic detail over time periods ranging from minutes to days.
We developed the RootChip, a polydimethylsiloxane (PDMS)- based microfluidic device, which allows us to grow and image roots from Arabidopsis seedlings while avoiding any physical stress to roots during preparation for imaging1 (Figure 1). The device contains a bifurcated channel structure featuring micromechanical valves to guide the fluid flow from solution inlets to each of the eight observation chambers2. This perfusion system allows the root microenvironment to be controlled and modified with precision and speed. The volume of the chambers is approximately 400 nl, thus requiring only minimal amounts of test solution.
Here we provide a detailed protocol for studying root biology on the RootChip using imaging-based approaches with real time resolution. Roots can be analyzed over several days using time lapse microscopy. Roots can be perfused with nutrient solutions or inhibitors, and up to eight seedlings can be analyzed in parallel. This system has the potential for a wide range of applications, including analysis of root growth in the presence or absence of chemicals, fluorescence-based analysis of gene expression, and the analysis of biosensors, e.g. FRET nanosensors3.

This article provides a protocol for cultivation of Arabidopsis seedlings in the RootChip, a microfluidic imaging platform that combines automated control of growth conditions with microscopic root monitoring and FRET-based measurement of intracellular metabolite levels.

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, ...

Bioengineering

A Flexible Low Cost Hydroponic System for Assessing Plant Responses to Small Molecules in Sterile Conditions

A wide range of studies in plant biology are performed using hydroponic cultures. In this work, an in vitro hydroponic growth system designed for assessing plant responses to chemicals and other substances of interest is presented. This system is highly efficient in obtaining homogeneous and healthy seedlings of the C3 and C4 model species Arabidopsis thaliana and Setaria viridis, respectively. The sterile cultivation avoids algae and microorganism contamination, which are known limiting factors for plant normal growth and development in hydroponics. In addition, this system is scalable, enabling the harvest of plant material on a large scale with minor mechanical damage, as well as the harvest of individual parts of a plant if desired. A detailed protocol demonstrating that this system has an easy and low-cost assembly, as it uses pipette racks as the main platform for growing plants, is provided. The feasibility of this system was validated using Arabidopsis seedlings to assess the effect of the drug AZD-8055, a chemical inhibitor of the target of rapamycin (TOR) kinase. TOR inhibition was efficiently detected as early as 30 min after an AZD-8055 treatment in roots and shoots. Furthermore, AZD-8055-treated plants displayed the expected starch-excess phenotype. We proposed this hydroponic system as an ideal method for plant researchers aiming to monitor the action of plant inducers or inhibitors, as well as to assess metabolic fluxes using isotope-labeling compounds which, in general, requires the use of expensive reagents.

A simple, versatile, and low-cost in vitro hydroponic system was successfully optimized, enabling large-scale experiments under sterile conditions. This system facilitates the application of chemicals in a solution and their efficient absorption by roots for molecular, biochemical, and physiological studies.

A wide range of studies in plant biology are performed using hydroponic cultures. In this work, an in vitro hydroponic growth system designed for ...

Environment

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 ...

Rhizobacterial Colonization Detection on Arabidopsis thaliana

Measuring Bacterial Colonization on Arabidopsis thaliana Roots in Hydroponic Condition

Measuring bacterial colonization on Arabidopsis thaliana root is one of the most frequent experiments in plant-microbe interaction studies. A standardized method for measuring bacterial colonization in the rhizosphere is necessary to improve reproducibility. We first cultured sterile A.thaliana in hydroponic conditions and then inoculated the bacterial cells in the rhizosphere at a final concentration of OD600 of 0.01. At 2 days post-inoculation, the root tissue was harvested and washed three times in sterile water to remove the uncolonized bacterial cells. The roots were then weighed, and the bacterial cells colonized on the root were collected by vortex. The cell suspension was diluted in a gradient with a phosphate-buffered saline (PBS) buffer, followed by plating onto a Luria-Bertani (LB) agar medium. The plates were incubated at 37 &#176;C for 10 h, and then, the single colonies on LB plates were counted and normalized to indicate the bacterial cells colonized on roots. This method is used to detect bacterial colonization in the rhizosphere in mono-interaction conditions, with good reproducibility.

Author Spotlight: Enhancing Rhizobacteria Colonization on Plant Roots for Improved Microbial Fertilizer Efficiency

Colonization of plant growth-promoting rhizobacteria (PGPR) in the rhizosphere is essential for its growth-promoting effect. It is necessary to standardize the method of detection of bacterial rhizosphere colonization. Here, we describe a reproducible method for quantifying bacterial colonization on the root surface.

Measuring bacterial colonization on Arabidopsis thaliana root is one of the most frequent experiments in plant-microbe interaction studies. A standardized ...

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 ...

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

A Hydroponic Co-cultivation System for Simultaneous and Systematic Analysis of Plant/Microbe Molecular Interactions and Signaling

An experimental design mimicking natural plant-microbe interactions is very important to delineate the complex plant-microbe signaling processes. Arabidopsis thaliana-Agrobacterium tumefaciens provides an excellent model system to study bacterial pathogenesis and plant interactions. Previous studies of plant-Agrobacterium interactions have largely relied on plant cell suspension cultures, the artificial wounding of plants, or the artificial induction of microbial virulence factors or plant defenses by synthetic chemicals. However, these methods are distinct from the natural signaling in planta, where plants and microbes recognize and respond in spatial and temporal manners. This work presents a hydroponic cocultivation system where intact plants are supported by metal mesh screens and cocultivated with Agrobacterium. In this cocultivation system, no synthetic phytohormone or chemical that induces microbial virulence or plant defense is supplemented. The hydroponic cocultivation system closely resembles natural plant-microbe interactions and signaling homeostasis in planta. Plant roots can be separated from the medium containing Agrobacterium, and the signaling and responses of both the plant hosts and the interacting microbes can be investigated simultaneously and systematically. At any given timepoint/interval, plant tissues or bacteria can be harvested separately for various &#34;omics&#34; analyses, demonstrating the power and efficacy of this system. The hydroponic cocultivation system can be easily adapted to study: 1) the reciprocal signaling of diverse plant-microbe systems, 2) signaling between a plant host and multiple microbial species (i.e.&#160;microbial consortia or microbiomes), 3) how nutrients and chemicals are implicated in plant-microbe signaling, and 4) how microbes interact with plant hosts and contribute to plant tolerance to biotic or abiotic stresses.

The described hydroponic cocultivation system supports intact plants with metal mesh screens and cocultivates them with bacteria. Plant tissue, bacteria, and secreted molecules can then be separately harvested for downstream analyses, simultaneously allowing for the molecular responses of both plant hosts and interacting microbes or microbiomes to be investigated.

An experimental design mimicking natural plant-microbe interactions is very important to delineate the complex plant-microbe signaling processes. ...

Biology

A Simple Protocol for Mapping the Plant Root System Architecture Traits

Comprehensive knowledge of plant root system architecture (RSA) development is critical for improving nutrient use efficiency and increasing crop cultivar tolerance to environmental challenges. An experimental protocol is presented for setting up the hydroponic system, plantlet growth, RSA spreading, and imaging. The approach used a magenta box-based hydroponic system containing polypropylene mesh supported by polycarbonate wedges. Experimental settings are exemplified by assessing the RSA of the plantlets under varying nutrient (phosphate [Pi]) supply. The system was established to examine the RSA of Arabidopsis, but it is readily adaptable to study other plants like Medicago sativa (Alfalfa). Arabidopsis thaliana (Col-0) plantlets are used in this investigation as an example to understand the plant RSA. Seeds are surface sterilized by treating ethanol and diluted commercial bleach, and kept at 4 &#176;C for stratification. The seeds are germinated and grown on a liquid half-MS medium on a polypropylene mesh supported by polycarbonate wedges. The plantlets are grown under standard growth conditions for the desired number days, gently picked out from the mesh, and submersed in water-containing agar plates. Each root system of the plantlets is spread gently on the water-filled plate with the help of a round art brush. These Petri plates are photographed or scanned at high resolution to document the RSA traits. The root traits, such as primary root, lateral roots, and branching zone, are measured using the freely available ImageJ software. This study provides techniques for measuring plant root characteristics in controlled environmental settings. We discuss how to (1) grow the plantlets, and collect and spread root samples, (2) obtain pictures of spread RSA samples, (3) capture the images, and (4) use image analysis software to quantify root attributes. The advantage of the present method is the versatile, easy, and efficient measurement of the RSA traits.

We use simple laboratory tools to examine the root system architecture (RSA) of Arabidopsis and Medicago. The plantlets are grown hydroponically over mesh and spread using an art brush to reveal the RSA. Images are taken using scanning or a high-resolution camera, then analyzed with ImageJ to map traits.

Comprehensive knowledge of plant root system architecture (RSA) development is critical for improving nutrient use efficiency and increasing crop cultivar ...

Semihydroponic Glass Jar System for Exudate Analysis in Multiple Plant Species

A Versatile Glass Jar System for Semihydroponic Root Exudate Profiling

Root exudates shape the plant-soil interface, are involved in nutrient cycling&#160;and modulate interactions with soil organisms. Root exudates are dynamic and shaped by biological, environmental, and experimental conditions. Due to their wide diversity and low concentrations, accurate exudate profiles are challenging to determine, even more so in natural environments where other organisms are present, turning over plant-derived compounds and producing additional compounds themselves. The semihydroponic glass jar experimental system introduced here allows control over biological, environmental, and experimental factors. It allows the growth of various phylogenetically distinct plant species for up to several months with or without microbes, in a variety of different growth media. The glass-based design offers a low metabolite background for high sensitivity and low environmental impact as it can be reused. Exudates can be sampled nondestructively, and conditions can be altered over the course of an experiment if desired. The setup is compatible with mass spectrometry analytics and other downstream analytical procedures. In summary, we present a versatile growth system suited for sensitive root exudate analysis in a variety of conditions.

Author Spotlight: Exploring Plant-Microbe Interactions Through Root Exudates in a Novel Growth System 

We present a protocol for a glass-based, semihydroponic experimental system supporting the growth of a variety of phylogenetically distinct plants with or without microbes. The system is compatible with different growth media and permits nondestructive root exudate sampling for downstream analysis.

Root exudates shape the plant-soil interface, are involved in nutrient cycling&#160;and modulate interactions with soil organisms. Root exudates are ...

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

Summary

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Chapters in this video

An Orthotopic Model of Serous Ovarian Cancer in Immunocompetent Mice for in vivo Tumor Imaging and Monitoring of Tumor Immune Responses

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

A Flexible Low Cost Hydroponic System for Assessing Plant Responses to Small Molecules in Sterile Conditions

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

Measuring Bacterial Colonization on Arabidopsis thaliana Roots in Hydroponic Condition

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

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

A Hydroponic Co-cultivation System for Simultaneous and Systematic Analysis of Plant/Microbe Molecular Interactions and Signaling

A Simple Protocol for Mapping the Plant Root System Architecture Traits

A Versatile Glass Jar System for Semihydroponic Root Exudate Profiling