We thank Prof. Dr. Nicola Zamboni and Prof. Dr. Uwe Sauer from ETH Z&#252;rich, Switzerland, for determining the root exudation profiles with direct injection and Prof. Dr. Klaus Schl&#228;ppi from the University of Basel for the A. thaliana commensal bacteria. Further, we acknowledge the Swiss National Science Foundation (PR00P3_185831 to J.S., supporting S.M., A.S., E.M.S.) and the PSC-Syngenta Fellowship program (granted to Prof. Dr. Klaus Schl&#228;ppi and J.S., supporting C.J.).

1. Preparation of seedlings: Surface sterilization of seeds
NOTE: The surface sterilization of seeds and all following steps must be done in sterile conditions unless noted otherwise. Steps 1.1 to 1.4 are specific for surface sterilization of A. thaliana seeds. Other plant species have alternative variations in tube size (depending on the number of seeds and seed size), time in solutions, and sterilization solutions (Table 1).
<ol>
	<li>Add seeds to a microcentrifuge tube (maximum 20 mg of seeds) and cover the seeds with 70% ethanol. 
	CAUTION: Ethanol is flammable. Do not use near open flame.</li>
	<li>Move the closed microcentrifuge tubes to a shaker for 15 min, and set the rotation such that the seeds are gently agitated.</li>
	<li>Carefully remove 70% ethanol with a pipette and replace it with 100% ethanol. Repeat step 1.2.</li>
	<li>Remove 100% ethanol and dry the seeds by leaving the microcentrifuge tube lids open in a sterile air flow.</li>
	<li>Once the seeds are dry, spread them out evenly onto 0.5x Murashige and Skoog (MS) medium with 0.7% phyto agar by flicking the tube or using sterile forceps. Close the agar plates and seal them with 1.25 cm micropore tape.</li>
	<li>Place the plates horizontally on a shelf in the growth chamber (16 h light/8 h dark, 22 &#176;C day/18 &#176;C night, 150-160 &#956;mol m-2&#160;s-1).</li>
</ol>
2. Preparation of seedlings: Transfer of germinated seeds to fresh plates
NOTE: The following steps are specific for A. thaliana and are not needed for other species.
<ol>
	<li>After germination, transfer 5 to 6 seedlings to new 0.7% phyto agar plates with 0.5x MS medium by placing the seedlings linearly, approximately 3 cm from the top (see the position of the rosette in Figure 2D).</li>
	<li>Seal agar plates with 1.25 cm micropore tape and grow vertically in the growth chamber (Figure 2D) until the seedlings are ready to be transferred into jars at 15-18 days after germination (Table 2).</li>
</ol>
3. Preparation of hydroponic system: Jar setup
<ol>
	<li>Add 150 mL of clean 5 mm glass beads to each clean jar and close with the lid (Figure 1A). 
	NOTE: In this protocol, 5 mm glass beads are suitable for most species22, but the bead size can be adjusted if desired.</li>
	<li>Place enough aluminum foil over the closed jars to cover the lid-to-jar junction and autoclave (121 &#176;C for 20 min) (Figure 1B).</li>
	<li>Keep the prepared jars covered until the plants are ready to be transferred (Table 2).</li>
</ol>
4. Preparation of hydroponic system: Addition of seedlings
<ol>
	<li>When the seedlings are ready to be transferred into jars (Table 2), remove the aluminum foil and lids of the jars in the sterile bench.</li>
	<li>If the jars are to be inoculated with bacteria, perform the following steps before moving on to step 4.3; otherwise, skip step 4.2.
	<ol>
		<li>Using a sterile inoculation loop, pick single colonies from pure bacterial cultures on agar plates and suspend them in 750 &#181;L of sterile 10 mM MgCl2. Repeat until the solution is turbid.</li>
		<li>Optional: Wash the suspension 3x with 750 &#181;L of sterile 10 mM MgCl2 by 5 min centrifugation at 1,000 &#215; g and room temperature followed by the removal of the supernatant and resuspension of the pellet in 750 &#181;L of 10 mM MgCl2.</li>
		<li>Optional: If inoculating several different bacterial species, mix the suspensions of single strains prepared in steps 4.2.1-4.2.2 in equal ratios before proceeding.</li>
		<li>Adjust the optical density at wavelength 600 nm (OD600) to 0.2-0.4 in 0.5x MS medium (pH 5.7 to 5.9, adjusted with KOH) or another growth medium of choice (see step 4.3). Measure the OD600 again to check if the solution was diluted correctly. 
		CAUTION: KOH is corrosive; wear gloves and lab coat.</li>
		<li>Dilute the suspension further to OD600 0.002-0.004 in the same medium (1:100 dilution). 
		NOTE: Final cell densities will depend on the bacterial strain used, but in our experience, this inoculum corresponds to approximately 3-6 &#215; 105 cells mL-1.</li>
		<li>Add 35 mL of the final dilution per jar. Add 35 mL of sterile growth medium to control jars. Stir the beads to evenly coat with 0.5x MS media before plant transfer so that the roots do not dry. Proceed to step 4.4.</li>
	</ol>
	</li>
	<li>Add 35 mL of 0.5x MS medium to each jar (pH 5.7 to 5.9, adjust with KOH). Stir the beads to evenly coat with 0.5x MS media before plant transfer so roots do not dry. 
	NOTE: The growth medium can be modified, for example, by the removal of nutrients, the addition of osmolytes to induce salt stress, or the use of different pH values or growth media. CAUTION: KOH is corrosive; wear gloves and lab coat.</li>
	<li>Place 3-5 A. thaliana plants into a jar by placing the root systems between the glass beads. 
	NOTE: The number of plants per jar is different for other species (Table 2).</li>
	<li>Move the beads around with sterile forceps or spoons to cover the roots and lift the leaves out of the media (Figure 1C). 
	NOTE: Carefully move the plants and beads to avoid breaking the roots and stressing the plants.</li>
	<li>Add 2 strips of 1.25 cm micropore tape across the top of the jars and gently place the lid on top (Figure 1D-F). 
	NOTE: Do not push down on the lid to avoid obstructing air exchange.</li>
	<li>Cover the gap between the lid and jar with 2.5 cm micropore tape to maintain sterility while allowing air exchange, and place the jars into a growth chamber (Figure 1G,H).</li>
	<li>Set up 4-8 jars per experimental condition, depending on the biological question and the variability expected.</li>
	<li>Make sure to include biological negative controls (e.g., jars without plants) to account for the metabolite background of the experimental system. Set up the same number of replicates for negative controls as for experimental treatments (e.g., quadruplicates).</li>
	<li>For extended growth periods, replace the growth medium weekly: remove the remainder of the growth medium with a 25 mL volumetric pipet and add in 35 mL of fresh medium.</li>
</ol>
5. Collection of root exudates
<ol>
	<li>When the plants reach the desired age, remove the 2.5 cm micropore tape and lid in the sterile bench. 
	NOTE: For A. thaliana in our growth conditions, 21 days after germination represents a mature vegetative stage before flowering onset. The stage of vegetative development is specific to the plant species, and the timing of the growth period changes with plant species; this timing can be modified depending on the research question.</li>
	<li>For sterility testing, aliquot 20 &#181;L of growth medium and pipet onto LB agar plates.
	<ol>
		<li>For inoculated jars, aliquot 100 &#181;L of growth medium to use for Colony-forming Unit (CFU) counts (e.g., in a dilution series23). 
		NOTE: In our experience, bacterial densities range within 105-108 live cells mL-1&#160;of growth medium.</li>
	</ol>
	</li>
	<li>Remove as much growth medium as possible with a 25 mL volumetric pipet, avoiding damage to the roots.</li>
	<li>Add 50 mL of collection medium (e.g., 20 mM ammonium acetate; pH 5.7-5.9 adjusted with HCl) by pipetting along the walls of the jar to avoid wetting the leaves. Close the jars with lids and incubate them in sterile conditions for the desired time of exudate collection. For time-sensitive experiments, 2 h is a good collection window. 
	CAUTION: HCl is corrosive; wear gloves and lab coat. For longer root exudate collection times, wrap the lids again with tape and move the jars to the growth chamber.</li>
	<li>After 2 h, remove the desired amount of collection media into tubes (e.g., 50 mL for assays or analyses using concentrated exudates, or 2 mL for direct use). Store the collected root exudates at -80 &#176;C until further processing or analysis.
	<ol>
		<li>When working with inoculated jars, centrifuge the collected exudates for 10 min at 5,000 &#215; g and 4 &#176;C and only use supernatants for the analysis. Alternatively, filter-sterilize the exudates with a 0.22 &#956;m or 0.45 &#956;m filter.</li>
	</ol>
	</li>
	<li>If the experiment is part of a time series, remove all remaining collection medium from jars and add 35 mL of 0.5x MS medium. Replace the lid and 2.5 cm tape before returning the jars to the growth chamber.</li>
	<li>Measure the root and shoot fresh weight of all the plants in one jar to later normalize root exudates by plant weight. 
	NOTE: Plant tissues can be sampled in addition if desired.</li>
</ol>
6. Processing root exudates for mass spectrometry
<ol>
	<li>Depending on the analytical method, freeze-dry the root exudates to concentrate and remove the collection medium (-80 &#176;C at 0.37 mbar until no liquid is present). Store at -80 &#176;C until ready to analyze. 
	NOTE: The data presented here was generated with flow-injection TOF-MS analysis on an accurate-mass quadrupole time-of-flight instrument.</li>
	<li>Prior to injection into the analytical instrument, reconstitute freeze-dried exudates or dilute unprocessed root exudates with collection medium according to the root fresh weight. 
	NOTE: The collection medium was chosen to be compatible with the mass spectrometer. However, depending on the analytical method, desalting steps might be necessary before injection.</li>
</ol>

Semihydroponic Glass Jar System for Exudate Analysis in Multiple Plant Species

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The authors have no conflicts of interest to disclose.

The experimental system presented here is based on glass jars and glass beads and thus provides a simple, low-maintenance, and versatile semihydroponic system to study root exudation in various contexts. It has been used in studies investigating the exudation profiles of different plant species25, the responses of exudation to different growth conditions25, as well as the influence of soil physiochemical properties on exudation22. The system is suitable for all plant species tested here for extended growth periods, ranging from weeks to months. The maintenance of sterile conditions is straightforward, as is the inoculation with bacteria, which persist over the analyzed 2-week growth period. Thus, the experimental system not only allows for a controlled collection of root exudates in sterile conditions, but it can also be used to study plant-microbe interactions. Furthermore, the plant growth media can be varied to study metabolic responses to different nutrient levels, and growth periods can be adjusted by adapting the light conditions or using different-sized jars.
Studying root exudates in hydroponic or semihydroponic conditions remains standard in the field&#160;mainly because of the enhanced resolution of low-concentration metabolites11. Many hydroponic approaches rely on Petri dishes, multi-well plates, or other small containers allowing sterility and high throughput but restricting experimentation to small plants or seedlings grown in high-humidity environments17,18,26,27. In the presented glass jar setup, sufficient head space is provided by the comparably large jars, allowing extended growth periods. Micropore tape stripes secure air exchange whilst maintaining sterility. Thus, even tall monocots such as barley and maize can be grown in the glass jar setup for multiple weeks. Small plants such as A. thaliana and clover can be studied for 4-5 weeks after germination, including vegetative and reproductive stages.
Alternative hydroponic setups are available for larger plants also, but these often require custom-made boxes and inlets made out of mesh, foam boards, and engraftment baskets for plant support15,28,29,30. In addition, these devices usually are not set up to be sterile, or they require challenging setup and maintenance procedures to keep them free from microbial and/or chemical contaminations. Setup and maintenance of sterility in the presented experimental system are straightforward. In addition, the use of glass for jars and beads reduces the presence of contaminants leaching from plastics and saves resources as it can easily be washed and reused.
Glass beads have been applied previously to mimic soil particles. They induce natural root development in root exudation sampling devices such as exudation traps31&#160;or other semihydroponic systems19. The glass-jar setup takes advantage of this development and introduces the beads as a colonization surface for microbes. In soil, the microbiome around plant roots evolves in a semisolid environment, with compact particles and spaces filled with air or water. Even though the glass jar setup does not include active aeration of the growth medium due to which the lower liquid phase likely does not contain optimal oxygen levels, the combination of a larger bead volume with a smaller growth medium volume creates a humid yet aerated upper phase where microbes can grow under oxic conditions. Others have proposed to shake growth containers26,28&#160;or use tubing coupled to air pumps19,29&#160;to maintain air supply in hydroponic growth systems. However, those systems either are set up to not be sterile, or require specialized material and constant surveillance to maintain sterility. In addition, in the case of shaking, much care to avoid submerging of shoots in growth solutions and damage to root systems. Nevertheless, if desired, the experimental setup presented could be adapted with additional material for aeration.
A crucial aspect to consider in all plant-microbe interaction studies investigating metabolism is that microbes degrade plant-derived compounds and produce metabolites on their own. Without a specialized sterile experimental setup, it is not possible to distinguish between plant- and microbe-derived metabolites. To inhibit microbial activity and enrich plant-derived compounds, Oburger et al. proposed to chemically sterilize the root exudate sampling solution to inhibit bacterial degradation32. The effect of chemical inhibitors could be studied in the presented experimental system, comparing exudation profiles of sterile versus nonsterile plants treated with or without the inhibitor.
A main limitation of the presented glass jar setup is that the growth conditions remain very artificial compared to soil. Exudates from soil grown-plants are often either collected from percolation systems13, where solvent flowthroughs are gathered at the base of growth containers, or soil-hydroponic hybrid systems, where plants are initially grown in soil and then transferred to hydroponic conditions16,33. In contrast to the glass jar setup, these procedures usually are destructive, not allowing for multiple collections over time in changing growth environments. Furthermore, whilst in percolating systems, the soil background is sampled together with the exudates, in soil-hydroponic hybrid systems the problem of high soil metabolic background is circumvented with the transfer to hydroponic conditions for exudate collection. Although recovery times have been implemented to reduce metabolite leakage via wounded roots11, the plant transfer is very disruptive and wounds are likely to persist, and plant metabolism might change in response to transfer to hydroponic conditions. Moreover, in many instances, an osmotic shock is induced by transferring plants to water instead of a suitable growth solution16,33. In the presented protocol, the growth solution is exchanged with an equimolar solution to maintain osmotic balance, still allowing to capture exudation within a short, defined time window. The change of growth solution is common practice in many published studies and can easily be achieved in hydroponic setups without root wounding12,16,26,34. Due to its versatility, the experimental system presented can easily be adapted to mimic more natural conditions, for example, by using sterile or nonsterile soil extract as a growth solution with or without the presence of solid soil particles. The gradual change towards natural conditions allows for the study of the impact of the different physiochemical soil properties and microbial presence on plant metabolism and physiology. Before the scientific community has a good understanding of exudation in various environments, it is desirable to employ soil-based and hydroponic systems in parallel, as both setups have their advantages and limitations13.
In conclusion, the presented semihydroponic, glass-based experimental setup stands out because of its simplicity combined with high versatility of applications. It presents an accessible, low-cost way to collect and study exudation in sterile conditions, or in combination with microbes and plant-microbe interactions.

Within densely populated soils, the rhizosphere presents a carbon-rich niche. It is shaped by plant roots via the exudation of up to 20% of assimilated carbon and harbors microbial communities that are distinct from the resident soil microbiome1,2,3,4,5,6. As researchers are tapping into the beneficial functions of root-associated microbes and the potential for sustainable agriculture that goes with it7, this observation, often termed as the rhizosphere effect, has been the focus of growing scientific efforts. However, thus far, the chemical dialog between microorganisms and plants, which is proposed to be the driver of the rhizosphere effect, remains poorly understood and hence, the mechanistic understanding for the development of reliable microbial solutions in agriculture is limited8,9,10.
Deciphering root exudates in soil environments where metabolites are readily absorbed by soil particles and quickly turned over by microbial communities is not straightforward, especially for plant species with fine root systems such as the model plant Arabidopsis thaliana11. This is why, in most studies, root exudates are sampled from hydroponic systems. In these microcosms, aerial parts of plants are held in place by customized plant-holders or more low-key materials such as mesh, agar, and glass beads. The containers used range from Petri dishes over multi-well plates to various custom and commercial boxes with or without aeration filters12,13,14,15,16,17,18,19. Depending on the system, the plant-growth conditions will vary greatly and reflect natural conditions to a greater or lesser extent.
Here, we present a glass-based, semihydroponic system that is experimentally amenable and produces highly reproducible results. It is straightforward to assemble and use and is based on commonly available materials. The system is based on a glass jar filled with glass beads, taking advantage of the reusable nature and low-binding properties of glassware (Figure 1). The beads provide physical support for the growing plant and simulate mechanical impedance, contributing to more soil-like root architecture when compared to hydroponic setups19,20,21. If inoculated with microbes, the glass beads present surfaces for bacteria to attach to.
The glass jar can be closed to maintain sterility and the system is designed to allow for sufficient headspace and air circulation, avoiding an environment saturated in humidity. The jars are suitable for the prolonged growth of different plant species and can be scaled up and down using different-sized jars. Here, applications for six plant species are shown, covering C3 and C4 grasses, dicots, and legumes. Among them are the model species A. thaliana (dicot), Brachypodium distachyon (C3 monocot), Medicago truncatula (legume),&#160;as well as crop species such as Solanum lycopersicum (tomato, dicot), Triticum aestivum (wheat, C3 monocot), and Sorghum bicolor (sorghum, C4 monocot). The protocol presented includes the experimental setup of the system, seed sterilization and germination of six plant species, transplantation of seedlings to jars, different growth media, microbe inoculation, root exudate sampling, and exudate processing for analytics.

<table><tbody><tr><td>Agar powder for bacteriology</td><td>VWR</td><td>20767.298</td><td></td></tr><tr><td>Aluminum foil</td><td>FORA GmbH</td><td></td><td></td></tr><tr><td>Ammonium acetate</td><td>Sigma-Aldrich</td><td>32301-1KG</td><td>ACS reagent, Eur &gt;- 98%</td></tr><tr><td>Autoclave VX-150</td><td>Systec</td><td>1150</td><td></td></tr><tr><td>Balance</td><td>Sartorius</td><td>QUINTIX64-1S</td><td></td></tr><tr><td>Centrifuge</td><td>Hermle Labortechnik GmbH</td><td>305.00 V05</td><td></td></tr><tr><td>Cuvettes</td><td>Greiner Bio-One</td><td>613101</td><td></td></tr><tr><td>Difco LB Broth, Lennox</td><td>BD</td><td>240210</td><td></td></tr><tr><td>Ethanol</td><td>Reuss-Chemie AG</td><td>RC-A15-A-005L</td><td></td></tr><tr><td>Filtered deionized water</td><td>Merck Millipore</td><td>Milli-Q IQ7000</td><td></td></tr><tr><td>Glass beads</td><td>Carl Roth</td><td>HH56.1</td><td>5 mm</td></tr><tr><td>Hydrochloric acid</td><td>Merk</td><td>1.00317.1000</td><td></td></tr><tr><td>Inoculation loop</td><td>Karl Hammacher GmbH</td><td>HWO_070-21</td><td></td></tr><tr><td>Jars</td><td>Weck</td><td>105741</td><td>850 mL</td></tr><tr><td>Lyophilizer</td><td>Christ</td><td>Alpha 2-4 LSCplus</td><td></td></tr><tr><td>Magnesium chloride hexahydrate</td><td>Carl Roth</td><td>2189.1</td><td></td></tr><tr><td>Matrix Orbital thermoshaker</td><td>IKA</td><td>10006248</td><td></td></tr><tr><td>Microcentrifuge tube</td><td>Sarstedt AG &amp;amp; Co. KG</td><td>72.695.500</td><td>SafeSeal reaction tube, 2 mL, PP</td></tr><tr><td>Micropore tape&amp;nbsp;</td><td>3M</td><td>1530-0</td><td>1.25 cm x 9.1 m</td></tr><tr><td>Micropore tape&amp;nbsp;</td><td>3M</td><td>1530-1</td><td>2.5 cm x 9.1 m</td></tr><tr><td>Murashige &amp;amp; Skoog Medium (MS)</td><td>Duchefa Biochemie</td><td>M0221.0050</td><td></td></tr><tr><td>Growth chamber</td><td>Percival</td><td>SE41-TLCU4</td><td>16 hour light/8 dark. 22 &amp;deg;C day/18 night</td></tr><tr><td>Phyto agar</td><td>Duchefa Biochemie</td><td>P1003.1000</td><td></td></tr><tr><td>Potassium hydroxide</td><td>Sigma-Aldrich</td><td>8.14353.0100</td><td></td></tr><tr><td>SmartSpec Plus Spectrophotometer</td><td>Bio-Rad</td><td>170-2525</td><td></td></tr><tr><td>Sodium hypochlorite solution, 12% Cl</td><td>Carl Roth</td><td>9062.4</td><td></td></tr><tr><td>Square petri dish</td><td>Greiner Bio-One</td><td>688102</td><td>120x120x17 mm, with vents</td></tr><tr><td>Stericup Quick release</td><td>Millipore</td><td>S2GPU05RE</td><td>0.22 &amp;micro;m PES, 500 mL</td></tr><tr><td>Sterile bench</td><td>FASTER S.r.l.</td><td>FlowFast H 18</td><td></td></tr></tbody></table>

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.

The experimental system introduced here allows control of experimental and environmental factors altering root exudation profiles. We compared A. thaliana growth under different lighting conditions, plant age, and plant densities across two different laboratories (Figure 3). Plants looked healthy across laboratories (Figure 3A,B). Short-day conditions (10 h lights vs. 16 h light, Figure 3) resulted in higher root mass compared to long-day conditions (Figure 3C,D). Similarly, the total root mass of all plants grown in long-day conditions was smaller than in short-day (Figure 3C). Overall, the variability of growth within and between jars was low across laboratories.
The glass jar system is compatible with a variety of plant species and developmental stages. The endpoint for root exudate analysis is typically 21 days, as in our experimental conditions, many plant species are in a mature vegetative stage before transitioning to the reproductive stage (Figure 4A-E). Plants can easily be removed from the experimental system without visible damage to the root system. Thus, determining tissue weights or using tissues for downstream analysis is straightforward. The highest shoot weights were found for wheat, followed by sorghum and tomato. The highest root weights were found for sorghum, followed by wheat and M. truncatula. The root:shoot ratio differs between species. Overall, tissue weights varied between 100 mg and 800 mg.
A central aspect of experimental systems allowing root exudate collection is the controlled inoculation with microbes to study plant-microbe interactions in a defined environment. The experimental system presented can be kept sterile even with repeated manipulation (see&#160;&#39;control&#39; condition in Figure 5), and bacteria can be added and maintained for extended periods. When 33-day-old A. thaliana plants were&#160;inoculated with a mixture of commensal bacteria at OD600 0.004, bacteria persisted for 12 days, which was the full duration of the experiment (Figure 5). Colony-forming units even increased 100-fold in the growth medium and also colonized the roots (Figure 5C). Phenotypes of inoculated plants were indistinguishable from those of sterile plants (Figure 5A,B).
The presented experimental system allows exudate collection in many experimental conditions. Here, we present exudation profiles of A. thaliana Col-0 collected either in ammonium acetate or in water from the same plants consecutively (Figure 6A). Exudates were stored at -80 &#176;C until analysis, normalized by root weight, and analyzed by mass spectrometry. Samples of jars containing plants were filtered against experimental controls (jars without plants). Of 2,163 metabolites, 436 showed a signal above the background (20.16%) and were retained for analysis. Of these, 416 or 95% of compounds showed distinct abundance between the experimental conditions. However, 26 metabolites could not be attributed to any kind of compound and are thus not defined. Most metabolites (406 or 98%) were more abundant in water-collected exudates. The consecutive collection of exudates first in ammonium acetate and later in water might impact the exudation profile, as metabolic signals might dilute with time. However, the almost exclusively higher exudation signal in water collected as a second timepoint does not support this hypothesis: Exudate collection in pure water likely creates an osmotic shock for plants, which causes the increased metabolite abundances compared to a collection solvent equimolar to the growth solution (20 mM ammonium acetate is equimolar to 0.5x MS medium). Investigation of chemical classes of identified compounds of both growth conditions showed that the majority of the compounds are organic acids and derivatives (28.6%) followed by organic oxygen compounds (18%), organoheterocyclic compounds (14.2%), and lipids (13.2%). Only a small subset of metabolites belong to phenylpropanoids and polyketides (8.7%) and benzenoids (6%). Organic nitrogen compounds, nucleosides, organosulfur compounds, alkaloids, lignans, and related compounds represent between 2% and 0.5% of the classified compounds (Figure 6B). The distribution of metabolites depicted here corresponds to already published data using other types of root exudates collection systems24.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66070/66070fig01v2.jpg" /> 
Figure 1: Glass jar setup. (A) Jar with glass beads. (B) Jar with glass beads ready to be autoclaved. (C) Seedlings in jars (top view). (D) Seedlings in a jar (top view) with 1.25 cm micropore tape. (E) Seedlings in jars (side view) with 1.25 cm micropore tape. (F) Seedlings in jars (side view) with 1.25 cm micropore tape and lid. (G) Complete jar setup with seedlings in jars (side view) with 1.25 cm micropore tape, lid, and 2.5 cm micropore tape. (H) Complete jar setup (top view). (I) Plants 21 days old and ready for harvest (top view). Jar size 147 mm height x 100 mm diameter. <a href="https://www.jove.com/files/ftp_upload/66070/66070fig01largev2.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/66070/66070fig02v2.jpg" /> 
Figure 2: Surface-sterilized seedlings on 0.5x Murashige and Skoog medium agar plates in a growth chamber. (A) Arabidopsis thaliana (6 days old), (B) Brachypodium distachyon (6 days old), (C) Medicago truncatula (6 days old), (D) A. thaliana (18 days old). Agar plate size 120 mm x 120 mm. <a href="https://www.jove.com/files/ftp_upload/66070/66070fig02largev2.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/66070/66070fig03.jpg" /> 
Figure 3: Arabidopsis thaliana Col-0 plants grown in jars under short- and long-day light conditions. (A) Jar with three 33-day-old plants grown in short-day conditions (10 h light/14 h dark, 220 &#181;mol m-2 s-1 light intensity, 21 &#176;C day/18 &#176;C night) and (B) jar with five 21-day-old plants grown in long-day conditions (16 h light/8 h dark, 150-160 &#181;mol m-2 s-1 light intensity, 22 &#176;C day/18 &#176;C night). Root weight of (C) all plants of one jar and (D) of single plants. ** represent significance values of t-test (p &#60; 0.05). <a href="https://www.jove.com/files/ftp_upload/66070/66070fig03large.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/66070/66070fig04v3.jpg" /> 
Figure 4: Five phylogenetically distinct species at day 21 in the sterile glass jar system setup. (A) Model monocot Brachypodium distachyon, (B) model legume Medicago truncatula, (C) dicot Solanum lycopersicum (tomato), (D) dicot Sorghum bicolor, (E) monocot Triticum aestivum (wheat). (F) Fresh weight of roots and shoots of the species grown in glass jars. <a href="https://www.jove.com/files/ftp_upload/66070/66070fig04largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66070/66070fig05v3.jpg" /> 
Figure 5: Arabidopsis thaliana Col-0 (33-day-old) grown in short-day conditions. (A) In a sterile setup or (B) inoculated for 12 days with a consortium of commensal bacteria. (C) Colony-forming units for sterile (control) and inoculated jars of the growth substrate (left) and the root (right). N = 4 jars for the sterile control condition, and n = 8 jars for the inoculated condition. <a href="https://www.jove.com/files/ftp_upload/66070/66070fig05largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66070/66070fig6v3.jpg" /> 
Figure 6: Distinct root exudate profiles for 21-day-old A. thaliana Col-0. Exudates were collected for 2 h in sterile 20 mM ammonium acetate (pH 5.7) followed by 2 h collection in sterile filtered deionized water. Metabolites were detected by mass spectrometry using direct injection. (A) Principal component analysis of 436 metabolites detected above background level (comparison of jars with plants against jars without plants). PC: principal component with the amount of variance explained. Blue: water-collected exudates, yellow: ammonium acetate-collected exudates. (B) Pie chart of 416 metabolites significantly different between experimental conditions (Tukey test) colored according to the superclass (https://cfb.fiehnlab.ucdavis.edu/). <a href="https://www.jove.com/files/ftp_upload/66070/66070fig06largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Species</td>
			<td>Seed preparation</td>
			<td>Time in 70% ethanol (min)</td>
			<td colspan="2">Concentration of Sodium hypochlorite (NaClO) (v/v % bleach)</td>
			<td>Time in bleach (min)</td>
		</tr>
		<tr>
			<td colspan="2">Arabidopsis thaliana</td>
			<td></td>
			<td>15</td>
			<td colspan="2">None; 100% ethanol&#160;</td>
			<td>15</td>
		</tr>
		<tr>
			<td colspan="2">Brachypodium distachyon*</td>
			<td>Dehusk</td>
			<td>0.5</td>
			<td colspan="2">6</td>
			<td>5</td>
		</tr>
		<tr>
			<td colspan="2">Medicago truncatula*a</td>
			<td></td>
			<td>30</td>
			<td colspan="2">6</td>
			<td>30</td>
		</tr>
		<tr>
			<td colspan="2">Solanum lycopersicum*</td>
			<td></td>
			<td>0.5</td>
			<td colspan="2">6</td>
			<td>5</td>
		</tr>
		<tr>
			<td colspan="2">Sorghum bicolor*</td>
			<td>Dehusk</td>
			<td>0.5</td>
			<td colspan="2">6</td>
			<td>30</td>
		</tr>
		<tr>
			<td colspan="2">Triticum aestivum*</td>
			<td>Dehusk</td>
			<td>0.5</td>
			<td colspan="2">12</td>
			<td>20</td>
		</tr>
	</tbody>
</table>
Table 1: Surface seed sterilization methods for multiple species. *Washed 4-5x with filtered deionized water between ethanol and bleach and at the end; aincubation for 3-6 h after bleach, replacing with filtered deionized water every 30 min.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Species</td>
			<td>Day to jars</td>
			<td>Number of plants</td>
		</tr>
		<tr>
			<td>Brachypodium distachyon</td>
			<td>4 to 5</td>
			<td>3</td>
		</tr>
		<tr>
			<td>Sorghum bicolor</td>
			<td>4 to 5</td>
			<td>3</td>
		</tr>
		<tr>
			<td>Triticum aestivum</td>
			<td>4 to 5</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Medicago truncatula</td>
			<td>5 to 6</td>
			<td>3</td>
		</tr>
		<tr>
			<td>Solanum lycopersicum</td>
			<td>7 to 8</td>
			<td>3</td>
		</tr>
		<tr>
			<td>Arabidopsis thaliana</td>
			<td>17</td>
			<td>3 to 5</td>
		</tr>
	</tbody>
</table>
Table 2: Age of seedlings in days for transfer to jars and number of plants per jar for various plant species.

Watch this Scientific Journal Video about Exploring Plant-Microbe Interactions Through Root Exudates in a Novel Growth System at JoVE.com

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.

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 ...

a-versatile-glass-jar-system-for-semihydroponic-root-exudate-profiling

Research

JoVE Journal

Biology

1.6K Views.  University of Zürich. 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.

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

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and detritus, and excreting particulate and dissolved compounds, they serve as important agents for benthic-pelagic coupling. Accurately measuring the compounds removed and excreted by suspension feeders (such as sponges, ascidians, polychaetes, bivalves) is crucial for the study of their physiology, metabolism, and feeding ecology, and is fundamental to determine the ecological relevance of the nutrient fluxes mediated by these organisms. However, the assessment of the rate by which suspension feeders process particulate and dissolved compounds in nature is restricted by the limitations of the currently available methodologies. Our goal was to develop a simple, reliable, and non-intrusive method that would allow clean and controlled water sampling from a specific point, such as the excurrent aperture of benthic suspension feeders, in situ. Our method allows simultaneous sampling of inhaled and exhaled water of the studied organism by using minute tubes installed on a custom-built manipulator device and carefully positioned inside the exhalant orifice of the sampled organism. Piercing a septum on the collecting vessel with a syringe needle attached to the distal end of each tube allows the external pressure to slowly force the sampled water into the vessel through the sampling tube. The slow and controlled sampling rate allows integrating the inherent patchiness in the water while ensuring contamination free sampling. We provide recommendations for the most suitable filtering devices, collection vessel, and storing procedures for the analyses of different particulate and dissolved compounds. The VacuSIP system offers a reliable method for the quantification of undisturbed suspension feeder metabolism in natural conditions that is cheap and easy to learn and apply to assess the physiology and functional role of filter feeders in different ecosystems.

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

We introduce the VacuSIP, a simple, non-intrusive, and reliable method for clean and accurate point sampling of water. The system was developed and evaluated for the simultaneous collection of the water inhaled and exhaled by benthic suspension feeders in situ, to cleanly measure removal and excretion of particulate and dissolved compounds.

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and ...

Environment

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 ...

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

Developmental Biology

Inoculation Strategies to Infect Plant Roots with Soil-Borne Microorganisms

The rhizosphere harbors a highly complex microbial community in which plant roots are constantly challenged. Roots are in close contact with a wide variety of microorganisms, but studies on soil-borne interactions are still behind those performed on aboveground organs. Although some inoculation strategies for infecting model plants with model root pathogens are described in the literature, it remains difficult to get a comprehensive methodological overview. To address this problem, three different root inoculation systems are precisely described that can be applied to gain insights into the biology of root-microbe interactions. For illustration, Verticillium species (namely, V. longisporum and V. dahliae) were employed as root invading model pathogens. However, the methods can be easily adapted to other root colonizing microbes - both pathogenic and beneficial. By colonizing the plant xylem, vascular soil-borne fungi such as Verticillium spp. exhibit a unique lifestyle. After root invasion, they spread via the xylem vessels acropetally, reach the shoot, and elicit disease symptoms. Three representative plant species were chosen as model hosts: Arabidopsis thaliana, economically important oilseed rape (Brassica napus), and tomato (Solanum lycopersicum). Step-by-step protocols are given. Representative results of pathogenicity assays, transcriptional analyses of marker genes, and independent confirmations by reporter constructs are shown. Furthermore, the advantages and disadvantages of each inoculation system are thoroughly discussed. These proven protocols can assist in providing approaches for research questions on root-microbe interactions. Knowing how plants cope with microbes in the soil is crucial for developing new strategies to improve agriculture.

This protocol presents a detailed summary of strategies to inoculate plant roots with soil-borne microbes. Exemplified for the fungi Verticillium longisporum and Verticillium dahliae, three different root infection systems are described. Potential applications and possible downstream analyses are highlighted, and advantages or disadvantages are discussed for each system.

The rhizosphere harbors a highly complex microbial community in which plant roots are constantly challenged. Roots are in close contact with a wide ...

Dissolved Solute Sampling Across an Oxic-Anoxic Soil-Water Interface Using Microdialysis Profilers

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface in response to disturbances. Deciphering the rapid biogeochemical changes requires in situ, minimally invasive tools with high spatial and temporal sampling resolution. However, the available passive sampling devices are not very useful in many cases either due to their disposable nature or the complexity and extensive workload for sample preparation.
To address this problem, a microdialysis profiler with 33 individual polyethersulfone nanomembrane tubes (semipermeable, &#60;20 nm pore size) integrated into the one-dimensional skeleton (60 mm) was established to iteratively sample the dissolved compounds in porewater across the soil-water interface at a high resolution of 1.8 mm (outer diameter plus one spacing, i.e., 0.1 mm between probes). The sampling mechanism is based on the principle of concentration gradient diffusion. The automatic loading of degassed water allows minimal disturbance to the chemical species across the oxic-anoxic interface.
This paper describes the procedures of device setup and continuous porewater sampling across the soil-water interface on a daily basis. Concentration-depth profiles were selectively measured before (on Day 6) and after (on Day 7) disturbances induced by irrigation. The results showed that concentration-depth profiles were undergoing rapid changes, especially for redox-sensitive elements (i.e., iron and arsenic). These protocols can help investigate the biogeochemical responses across the soil-water interface under various disturbances caused by physical, chemical, and biological factors. The paper thoroughly discusses the advantages and disadvantages of this method for potential use in the environmental sciences.

A microdialysis profiler is described to sample dissolved porewater solutes across an oxic-anoxic soil-water interface in situ with minimal disturbance. This device is designed to capture rapid changes in concentration-depth profiles induced by disturbances at the soil-water interface and beyond.

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface ...

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 ...

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 ...

Hydroponics: A Versatile System to Study Nutrient Allocation and Plant Responses to Nutrient Availability and Exposure to Toxic Elements

Hydroponic systems have been utilized as one of the standard methods for plant biology research and are also used in commercial production for several crops, including lettuce and tomato. Within the plant research community, numerous hydroponic systems have been designed to study plant responses to biotic and abiotic stresses. Here we present a hydroponic protocol that can be easily implemented in laboratories interested in pursuing studies on plant mineral nutrition.
This protocol describes the hydroponic system set up in detail and the preparation of plant material for successful experiments. Most of the materials described in this protocol can be found outside scientific supply companies, making the set up for hydroponic experiments less expensive and convenient.
The use of a hydroponic growth system is most advantageous in situations where the nutrient media need to be well controlled and when intact roots need to be harvested for downstream applications. We also demonstrate how nutrient concentrations can be modified to induce plant responses to both essential nutrients and toxic non-essential elements.

Here, we present an easy-to-follow protocol to establish a successful hydroponic system for plant nutrition studies. This protocol has been extensively tested in Arabidopsis and can easily be adapted to other plant species to study specific nutritional requirements or the effect of non-essential elements on plant growth and development.

Hydroponic systems have been utilized as one of the standard methods for plant biology research and are also used in commercial production for several ...

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.

A Versatile Glass Jar System for Semihydroponic Root Exudate Profiling

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Chapters in this video

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

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

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Inoculation Strategies to Infect Plant Roots with Soil-Borne Microorganisms

Dissolved Solute Sampling Across an Oxic-Anoxic Soil-Water Interface Using Microdialysis Profilers

Measuring Bacterial Colonization on Arabidopsis thaliana Roots in Hydroponic Condition

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

Hydroponics: A Versatile System to Study Nutrient Allocation and Plant Responses to Nutrient Availability and Exposure to Toxic Elements

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