This work was supported by the USDA-NIFA grant: 2017-67013-26520. Tracy Debenport and Claire Fogan provided technical support and Ruby Ye and Casey Cosetta provide helpful comments on early versions of this manuscript.

1. Growing germ-free cabbages
<ol>
	<li>Preparing equipment for growing germ-free cabbages
	<ol>
		<li>Cleaning the calcined clay to remove fine dust particles
		<ol>
			<li>Rinse calcined clay (Table of Materials) at least 3x with tap water; drain off water. 
			CAUTION: Calcined clay produces very fine dust and it is recommended to wear a protective mask (Table of Materials) when washing.</li>
			<li>Spread calcined clay out as a thin layer (~4 cm) into an autoclave tray and autoclave on a dry cycle (121 &#176;C heating for 20 min and 20 min drying time) to sterilize.</li>
			<li>Allow the calcined clay to fully dry prior to use by spreading out on trays and placing in a warm incubator (30&#8722;37 &#176;C) for at least a week. Stir to mix every 3 days to fully dry the calcined clay so that it will absorb an even amount of Murashige and Skoog (MS) nutrient broth (section 1.2). 
			NOTE: Drying also helps to keep the volume of calcined clay even when it is weighed into tubes. Drying by other means, such as a drying oven, would also be suitable.</li>
		</ol>
		</li>
		<li>Cleaning the glassware for growing germ-free cabbages
		<ol>
			<li>Thoroughly clean and sterilize the glass tubes (Table of Materials) between each use. Soak tubes for 30 min in 30% bleach solution and rinse well with tap water before cleaning in an acid wash on a bacteriology setting. Acid-wash two-way test tube caps (Table of Materials) between uses.</li>
		</ol>
		</li>
		<li>Surface sterilizing cabbage seeds
		<ol>
			<li>Place up to 100 Napa cabbage (B. rapa var pekinensis) seeds in a 1.5 mL microcentrifuge tube. 
			NOTE: Adding more than 100 seeds to one microcentrifuge tube or changing the size of the tube may affect germination rates of the seeds due to lack of seed coat removal.</li>
			<li>Add 1 mL of 70% ethanol to the seeds and vortex for 5 min. Discard the ethanol using a pipette.</li>
			<li>Add 1 mL of 50% bleach and vortex for 5 min. Discard the bleach solution using a pipette.</li>
			<li>Add 1 mL of autoclaved deionized water and vortex for 5 min. Discard the deionized water using a pipette.</li>
			<li>Repeat step 1.1.3.4 3x to rinse off all bleach. Soak the seeds in sterile deionized water for 2&#8722;8 h prior to planting to soften the seed coat.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Growing germ-free cabbages 
	NOTE: Napa cabbages (B. rapa var pekinensis) are grown in glass tubes (15 cm x 2.5 cm) containing calcined clay soaked in Murashige and Skoog (MS) nutrient broth (Figure 1).
	<ol>
		<li>Weigh 10 g of clean calcined clay into a clean glass tube (15 cm x 2.5 cm).</li>
		<li>Prepare MS nutrient broth by dissolving 4.4 g of MS medium in 1 L of deionized water. Add MS nutrient broth (~9 mL) to each glass tube to cover the calcined clay using a pipette. 
		NOTE: Standing liquid in the tube will prevent the seed from germinating so it might benecessary to add slightly less MS broth to some tubes.</li>
		<li>Loosely cap glass tubes with 22 mm two-way test tube caps and autoclave (121 &#176;C for 60 min). When removing them from the autoclave, push caps onto the glass tubes to seal them. Cool tubes to room temperature before use.</li>
		<li>Gently place one sterile cabbage seed into the center of each tube using sterile, extra-long (25.4 cm) forceps. Place the tubes in a 7-way tray then place under light racks (full-spectrum T5 fluorescent bulbs or other illumination setup for plant growth) with a 16 h light cycle at 24 &#176;C. 
		NOTE: Seeds germinate overnight and develop their first true leaf after 5 days. A true leaf is the first vascular leaf after the cotyledons have formed. It has a more wrinkled edge and in Brassica rapa is covered in trichomes.</li>
	</ol>
	</li>
	<li>Testing for sterility of germ-free cabbages 
	NOTE: To test whether the cabbages are germ-free, select a few (5&#8722;10) cabbages from each batch and plate out to determine whether any culturable colonies are present.
	<ol>
		<li>Gently remove the cabbage from the glass tubes by gripping the base of the plant with sterilized forceps and pulling it out. Before removing the cabbage fully from the tube, carefully cut off the roots using sterilized dissection scissors. Compact the cabbage leaves into a 1.5 mL microcentrifuge tube. 
		NOTE: Larger cabbages might require removing one or two of the larger leaves while the cabbage is still in the tube to make it easier to get the cabbage into the 1.5 mL microcentrifuge tube. These larger leaves can be added to the 1.5 mL tube after the rest of the cabbage has been placed into the tube if the entire cabbage is required.</li>
		<li>Add 400 &#181;L of 1x phosphate buffer saline (PBS) to each 1.5 mL microcentrifuge tube. Using a sterile micropestle, homogenize the cabbage by pestling 30x.</li>
		<li>Plate 100 &#181;L of the cabbage homogenate onto agar plates to determine whether there are any contaminants present in the sample. Most bacteria found in the phyllosphere will grow on tryptic soy (TS) agar plates. Use wide orifice pipette tips when plating cabbage homogenate as the cabbage homogenate is thick and can clog regular pipette tips.</li>
	</ol>
	</li>
</ol>
2. Inoculating the phyllosphere with microbial solutions
<ol>
	<li>Making glycerol stocks of inoculation strains 
	NOTE: Table 1 lists the microbial isolates that can be used in this step. Other phyllosphere isolates could also be used here.
	<ol>
		<li>Densely streak out individual colonies from a fresh streak, onto two/three new plates of the same media to get many colonies.</li>
		<li>Let streaks grow for 2&#8722;5 days then scrape colonies from all plates into a 15 mL conical tube containing 15 mL of 15% glycerol, and vortex to mix thoroughly.</li>
		<li>Transfer an aliquot of 1 mL of the well-mixed glycerol stock into a 1.5 mL microcentrifuge tube and store glycerol stocks at -80 &#176;C until use. Save the remaining 14 mL of glycerol stock at -80 &#176;C as relatively large volumes of inoculation solution are required when inoculating cabbages.</li>
		<li>One week before use, thaw the 1.5 mL tube containing 1 mL of glycerol stock (from step 2.1.3) on ice, dilute, and plate at several different dilutions (e.g., 10-4, 10-5, and 10-6) to determine the concentration (colony-forming unit [CFU] per &#181;L) of the 14 mL of inoculation solution.</li>
	</ol>
	</li>
	<li>Sterilizing inoculation spray bottles
	<ol>
		<li>Disassemble the amber round Boston pump bottles (59 mL) and soak all components (pump, tube, cap and bottle) in 30% bleach solution for 30 min in a large plastic container with a tightly fitting lid.</li>
		<li>After soaking, carefully pour out all bleach from the container by lifting just one corner of the lid of the container.</li>
		<li>Rinse the bottles by filling the plastic container with autoclaved deionized water (~1 L depending on the container size) and carefully pour out deionized water, again by lifting the lid at one corner.</li>
		<li>Sterilize a biosafety cabinet by spraying with 70% ethanol solution and turning on the UV light for 30 min. 
		NOTE: Continue this work in the biosafety cabinet so that there is no risk of microbial contamination of the bottles as they air dry.</li>
		<li>Remove bottles from the large plastic container and fill each bottle with autoclaved deionized water using a pipette. Reassemble the pumps and place one in each bottle. Pump the deionized water through each bottle (10 sprays per bottle) to remove bleach from the pump component of the bottle.</li>
		<li>Repeat step 2.2.5 to ensure that all bleach is removed from the glass bottles.</li>
		<li>Test whether bottles are sterile by placing a number on each bottle (sticking lab tape to the side of the bottle when it is fully dry) then add 10 mL of 1x PBS to each of the bottles and pump 3 sprays onto a TS agar plate. After spraying, incubate the plates for one week at room temperature. If any colonies grow on a plate it indicates that the respective bottle was not sterile and should not be used for experiments.</li>
		<li>Before storing the sterile bottles, remove all remaining PBS and allow the bottles to dry thoroughly in the biosafety cabinet. Store sterile bottles in a sterile plastic container (typically the container used for bleaching the bottles) until use.</li>
	</ol>
	</li>
	<li>Preparing the microbial inoculum and spraying germ-free cabbages 
	CAUTION: All steps should be performed in a biosafety cabinet, as spraying aerosolizes the microbial solutions which could contaminate work surfaces or pose a health risk if carried out on a lab bench. 
	NOTE: Cabbages will form true leaves after 5 days, so it is advisable to wait one week after planting the cabbage before inoculating with any microbial solutions. As the tubes are sealed, there is no need to water the cabbages. Experiments are best performed within a month of planting, as the small tubes restrict the cabbage&#39;s growth.
	<ol>
		<li>Thaw glycerol stocks on ice and dilute in 1x PBS to the desired inoculation concentration (concentration determined by thawing and plating a 1 mL aliquot in step 2.1.4). 
		NOTE: A variety of different inoculation levels can be used, but phyllosphere isolates can grow from 104 to 108 CFUs/mL of cabbage slurry in 10 days.</li>
		<li>Add 10 mL of diluted glycerol stock to the sterile pump bottle and pump 5 sprays into a large waste collection beaker to remove any residual PBS from the bottle pump component.</li>
		<li>Remove the lid from the cabbage tube, tilt the cabbage towards the spray bottle, and spray each cabbage with 3 pumps of the inoculation solution, which provides ~600 &#181;L of inoculum.</li>
		<li>After inoculating, harvest a subset of the cabbages to assess the actual input inoculation concentration. Remove the cabbage from a tube with sterilized forceps. Cut off the roots with sterile dissection scissors and then carefully place the cabbage in a preweighed sterile 1.5 mL microcentrifuge tube. Record the weight of the cabbage for future calculations if CFUs/g of cabbage is required for calculations.</li>
		<li>Add 400 &#181;L of 1x PBS to each 1.5 mL microcentrifuge tube containing cabbage and use a sterile micropestle to homogenize the cabbage into the 1x PBS by grinding it 30x.</li>
		<li>Dilute cabbage homogenate (if required) and plate out the pestled cabbage mixture. Use wide orifice tips for pipetting the cabbage slurry because it will be thick and full of plant tissue pieces.</li>
	</ol>
	</li>
</ol>
3. Preparing sterile vegetable extract
NOTE: This method is a modified version of cabbage sterile media production18,19.
<ol>
	<li>Purchase a cabbage from a supermarket. In the lab, remove and discard the outermost leaves of the cabbage. Chop all remaining cabbage to fit into a blender and homogenize cabbage to a fine pulp, i.e., the cabbage will not get any finer with further blending. 
	NOTE: Any blender which can chop cabbage to a smooth homogeneous pulp should be suitable for this method.</li>
	<li>Weigh the blended cabbage homogenate and add 2 mL of distilled water per gram of cabbage. Filter the blended cabbage slurry through 2 layers of basket coffee filters (unbleached paper).</li>
	<li>Dispense the cabbage slurry into centrifuge tubes (size is dependent on the centrifuge). Centrifuge the filtered cabbage slurry at 20,000 x g for 20 min until large particles settle out of solution. 
	NOTE: It is essential to centrifuge the cabbage slurry for a long period of time as cabbage particles rapidly clog the filter sterilizer.</li>
	<li>Using a serological pipette, remove the supernatant from the pelleted cabbage debris taking care not to disturb the pelleted cabbage. If aiming to recreate fermentation conditions where standard salt concentrations are used, add 2% w/v NaCl at this step (i.e., before filter sterilization).</li>
	<li>Filter sterilize the vegetable extract using a 0.2 &#181;m filter (500 mL or 1 L) attached to a vacuum. Dispense into sterile tubes (either 50 mL centrifuge tubes or 15 mL centrifuge tubes) and freeze at -80 &#176;C until use.</li>
</ol>
4. Inoculation of sterile vegetable extract
<ol>
	<li>Thaw SVE and dispense 490 &#181;L into 1.5 mL microcentrifuge tubes. Use sufficient tubes to have at least five replicates per treatment per timepoint as each timepoint measurement is destructive.</li>
	<li>Thaw glycerol stocks of microbial isolates on ice and dilute with 1x PBS to the desired concentration. The concentration of lactic acid bacteria can be as low as 5,000 CFU per mL of SVE. To achieve this concentration, dilute stocks to 250 CFUs/&#181;L because 10 &#181;L will be used for inoculation of a total volume of 500 &#181;L.</li>
	<li>Inoculate SVE with 10 &#181;L of diluted microbial isolate. Pipette up and down a few times to thoroughly mix. Incubate at desired temperature (14 &#176;C for kimchi production temperature or 24 &#176;C for warmest sauerkraut fermentation).</li>
	<li>Measure rate of growth of microbial isolate in the SVE by harvesting replicate tubes on day 1, day 2, day 4, day 7 and day 14. 
	NOTE: Fermentation proceeds rapidly at the outset and slows over time. Therefore, having more initial timepoints gives greater resolution to the dynamics of how fermentation proceeds.</li>
	<li>At each timepoint, mix the inoculated SVE well by pipetting up and down a few times. Serially dilute the inoculated SVE in 1x PBS and plate onto agar plates. Incubate the agar plates for 4&#8722;7 days before counting colonies. 
	NOTE: Man, Rogosa, and Sharpe (MRS) agar should be used to enumerate all lactic acid bacteria, yeast peptone dextrose (YPD) should be used for yeast, and TS agar for most other bacteria isolates from the phyllosphere.</li>
	<li>Record the pH of the samples at each timepoint using a micro pH probe. 
	NOTE: This step should be carried out after plating because the pH probe will transfer cells between tubes/treatments.</li>
</ol>

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The authors have nothing to disclose.

Germ-free Napa cabbage plants have been used to study dispersal limitation of lactic acid bacteria in the Napa cabbage phyllosphere17. Germ-free Napa cabbages can also be used to test individual or pair-wise growth in the phyllosphere (Figure 1). Methods for making sterile vegetable extract has been tested for three different varieties of cabbage: red, green and Napa. Each of these SVEs act as a reliable growth media; inoculated microbes grow consistently across the d...

Microbial diversity of the phyllosphere plays an important role in maintaining plant health and can also influence the ability of plants to withstand environmental stress1,2,3,4,5. In turn, the health of crops directly impacts food safety and quality6,7. Plants play a role in ecosystem functioning and their associated microbiomes both affect the ability of plants to carry out these activities as well as directly influencing the environment themselves8. While scientists have begun to decipher the function and composition of the phyllosphere, the ecological processes that influence phyllosphere microbial community assembly are not fully understood9,10. The phyllosphere microbiome is an excellent experimental system for studying the ecology of microbiomes11. These communities are relatively simple and many of the community members can be grown on standard lab media10,12,13.
Fermented vegetables are one system where the community structure of the phyllosphere has important consequences. In both sauerkraut and kimchi, the microbes that naturally occur on vegetable leaves (the phyllosphere of Brassica species) serves as the inoculum for fermentation14,15. Lactic acid bacteria (LAB) are considered ubiquitous members of vegetable microbiomes, however they can be in low abundance in the phyllosphere16. Strong abiotic selection during fermentation drives a shift in microbial community composition enabling lactic acid bacteria to increase in abundance. As LAB grow, they produce lactic acid which creates the acidic environment of fermented vegetable products17. The link between the phyllosphere and the ferment provides an opportunity to use vegetables as a model to understand how microbiomes are structured.
We have developed methods to grow germ-free Napa cabbages and to inoculate them with specific microbial communities using spray bottles. This is an inexpensive and reliable method of evenly inoculating the cabbage with either individual microbes or mixed communities. A sterile vegetable extract (SVE) has also been developed from three different cabbage types/varieties: red and green cabbage (Brassica oleracea) and Napa cabbage (B. rapa). The addition of salt to these SVEs replicates the fermentation environment and allows for small-scale and relatively high-throughput experimental studies of fermentation microbiome assembly. These methods can be used to study microbial community assembly in the phyllosphere and how microbial community dynamics in the phyllosphere can be linked to the success of vegetable fermentation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5 mL microcentrifuge tubes</td>
			<td>VWR</td>
			<td>20170-650</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL conical tubes</td>
			<td>Falcon</td>
			<td>352096</td>
			<td></td>
		</tr>
		<tr>
			<td>7-way tray tray</td>
			<td>Sigma Magenta</td>
			<td>T8654</td>
			<td></td>
		</tr>
		<tr>
			<td>Amber Round Boston Glass Bottle</td>
			<td></td>
			<td>GPS 712OZSPPK12BR</td>
			<td>Ordered on Amazon.com from various suppliers</td>
		</tr>
		<tr>
			<td>Basket coffee filters</td>
			<td>If you care</td>
			<td></td>
			<td>(unbleached paper) Purchased from Wholefoods</td>
		</tr>
		<tr>
			<td>Bleach (mercury-free)</td>
			<td>Austin&#39;s</td>
			<td>50-010-45</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate Glass tubes</td>
			<td>VWR</td>
			<td>47729-586</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcined clay</td>
			<td>Turface</td>
			<td>MVP</td>
			<td>Ordered on Amazon.com from Root Naturally 6 Quart Bags. Particle size approximately 3-5 mm</td>
		</tr>
		<tr>
			<td>Cuisinart blender</td>
			<td>Cuisinart</td>
			<td></td>
			<td>Cuisinart Mini-Prep Plus Food Processor, 3-Cup</td>
		</tr>
		<tr>
			<td>Dissection scissors</td>
			<td>7-389-A</td>
			<td>American Educational Products</td>
			<td>Ordered on Amazon.com</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>VWR</td>
			<td>89125-172</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Aven</td>
			<td>18434</td>
			<td>Ordered on Amazon.com</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Fisher Scientific</td>
			<td>56-81-5</td>
			<td></td>
		</tr>
		<tr>
			<td>KleenGuard M10</td>
			<td>Kimberley-Clark</td>
			<td>64240</td>
			<td></td>
		</tr>
		<tr>
			<td>Large plastic container</td>
			<td>Rubbermaid</td>
			<td></td>
			<td>Ordered on Amazon.com</td>
		</tr>
		<tr>
			<td>Light racks</td>
			<td>Gardner&#39;s Supply</td>
			<td>39-357</td>
			<td>full-spectrum T5 fluorescent bulbs</td>
		</tr>
		<tr>
			<td>Magenta tm 2-way caps</td>
			<td>Millipore Sigma</td>
			<td>C1934</td>
			<td></td>
		</tr>
		<tr>
			<td>Man, Rogosa, and Sharpe</td>
			<td>Fisher Scientific</td>
			<td>DF0881-17-5</td>
			<td>This media is for broth and 15 g of agar is added to make plates</td>
		</tr>
		<tr>
			<td>Micro pH probe</td>
			<td>Thermo Scientific</td>
			<td>8220BNWP</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropestle</td>
			<td>Carolina</td>
			<td>215828</td>
			<td>Also called Pellet Pestle</td>
		</tr>
		<tr>
			<td>MS nutrient broth</td>
			<td>Millipore Sigma</td>
			<td>M5519</td>
			<td>Murashige and Skoog Basal Medium</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma Aldrich</td>
			<td>S9888</td>
			<td></td>
		</tr>
		<tr>
			<td>Napa cabbage seeds</td>
			<td>Johnny&#39;s Select Seeds</td>
			<td>2814G</td>
			<td>B. rapa var pekinensis (Bilko)</td>
		</tr>
		<tr>
			<td>Petri dish 100 mm x 15 mm</td>
			<td>Fisher</td>
			<td>FB0875712</td>
			<td>Used to make agar plates</td>
		</tr>
		<tr>
			<td>Phosphate buffer saline</td>
			<td>Fisher Scientific</td>
			<td>50-842-941</td>
			<td>Teknova</td>
		</tr>
		<tr>
			<td>Plant tissue culture box</td>
			<td>Sigma</td>
			<td>Magenta GA-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Serologial pipettes</td>
			<td>VWR</td>
			<td>89130-900</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile dowel</td>
			<td>Puritan</td>
			<td>10805-018</td>
			<td>Autoclave before use to sterilize</td>
		</tr>
		<tr>
			<td>Sterilizing 0.2 &#181;m filter</td>
			<td>Nalgene</td>
			<td>974103</td>
			<td></td>
		</tr>
		<tr>
			<td>Tryptic soy agar</td>
			<td>Fisher Scientific</td>
			<td>DF0370-17-3</td>
			<td>This media is for broth and 15 g of agar is added to make plates</td>
		</tr>
		<tr>
			<td>Wide orifice pipette tips</td>
			<td>Rainin</td>
			<td>17007102</td>
			<td></td>
		</tr>
		<tr>
			<td>Yeast, peptone and dextrose</td>
			<td>Fisher Scientific</td>
			<td>DF0428-17-5</td>
			<td>This media is suitable but media can also be made using yeast, peptone and dextrose, add 15 g of agar when making plates</td>
		</tr>
	</tbody>
</table>

a gnotobiotic system for studying microbiome assembly in the phyllosphere and in vegetable fermentation

The phyllosphere, the above ground portion of the plant that can be colonized by microbes, is a useful model system to identify processes of microbial community assembly. This protocol outlines a system for studying microbial community dynamics in the phyllosphere of Napa cabbage plants. It describes how to grow germ-free plants in test tubes with a calcined clay and nutrient broth substrate. Inoculation of germ-free plants with specific microbial cultures provides opportunities to measure microbial growth and community dynamics in the phyllosphere. Through the use of sterile vegetable extract produced from cabbages shifts in microbial communities that occur during fermentation can also be assessed. This system is relatively simple and inexpensive to set up in the lab and can be used to address key ecological questions in microbial community assembly. It also provides opportunities to understand how phyllosphere community composition can impact the microbial diversity and quality of vegetable fermentations. This approach for developing gnotobiotic cabbage phyllosphere communities could be applied to other wild and agricultural plant species.

Growth rates of Napa cabbages 
The seed sterilization method was tested with several different Napa cabbages (B. rapa var pekinese; Supplemental Figure 1) from a number of different suppliers and all grew consistently with similar growth rates. However, testing the methods with different species of Brassica (B. rapa: Turnip Purple Top; B. oleracea: Cairo Hybrid, Tropic Giant Hybrid; B. campestris: Pak Choi...

Watch this Scientific Journal Video about A Gnotobiotic System for Studying Microbiome Assembly in the Phyllosphere and in Vegetable Fermentation at JoVE.com

A Gnotobiotic System for Studying Microbiome Assembly in the Phyllosphere and in Vegetable Fermentation

A method of growing germ-free Napa cabbages has been developed which enables researchers to evaluate how single microbial species or multispecies microbial communities interact on cabbage leaf surfaces. A sterile vegetable extract is also presented which can be used to measure shifts in community composition during vegetable fermentation.

The phyllosphere, the above ground portion of the plant that can be colonized by microbes, is a useful model system to identify processes of microbial ...

a-gnotobiotic-system-for-studying-microbiome-assembly-phyllosphere

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7.1K Views.  Tufts University. A method of growing germ-free Napa cabbages has been developed which enables researchers to evaluate how single microbial species or multispecies microbial communities interact on cabbage leaf surfaces. A sterile vegetable extract is also presented which can be used to measure shifts in community composition during vegetable fermentation.

Video: A Gnotobiotic System for Studying Microbiome Assembly in the Phyllosphere and in Vegetable Fermentation

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail millet. Recently, the 510 Mb genome of foxtail millet, S. italica, has been sequenced 1,2 and a 25x coverage genome sequence of the weedy relative S. viridis is in progress. S. viridis has a number of characteristics that make it a potentially excellent model genetic system including a rapid generation time, small stature, simple growth requirements, prolific seed production 3 and developed systems for both transient and stable transformation 4. However, the genetics of S. viridis is largely unexplored, in part, due to the lack of detailed methods for performing crosses. To date, no standard protocol has been adopted that will permit rapid production of seeds from controlled crosses.
The protocol presented here is optimized for performing genetic crosses in S. viridis, accession A10.1. We have employed a simple heat treatment with warm water for emasculation after pruning the panicle to retain 20-30 florets and labeling of flowers to eliminate seeds resulting from newly developed flowers after emasculation. After testing a series of heat treatments at permissive temperatures and varying the duration of dipping, we have established an optimum temperature and time range of 48 &deg;C for 3-6 min. By using this method, a minimum of 15 crosses can be performed by a single worker per day and an average of 3-5 outcross progeny per panicle can be recovered. Therefore, an average of 45-75 outcross progeny can be produced by one person in a single day. Broad implementation of this technique will facilitate the development of recombinant inbred line populations of S. viridis X S. viridis or S. viridis X S. italica, mapping mutations through bulk segregant analysis and creating higher order mutants for genetic analysis.

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

We have developed a methodology for performing crosses in Setaria viridis (S. viridis). The method involves pruning the panicle prior to a hot water treatment to kill viable pollen. Crosses are performed following a well-controlled growth regime and typically result in the recovery of 1 to 7 cross-pollinated seed/s per panicle.

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail ...

Technique for Studying Arthropod and Microbial Communities within Tree Tissues

Phloem tissues of pine are habitats for many thousands of organisms. Arthropods and microbes use phloem and cambium tissues to seek mates, lay eggs, rear young, feed, or hide from natural enemies or harsh environmental conditions outside of the tree. Organisms that persist within the phloem habitat are difficult to observe given their location under bark. We provide a technique to preserve intact phloem and prepare it for experimentation with invertebrates and microorganisms. The apparatus is called a &#8216;phloem sandwich&#8217; and allows for the introduction and observation of arthropods, microbes, and other organisms. This technique has resulted in a better understanding of the feeding behaviors, life-history traits, reproduction, development, and interactions of organisms within tree phloem. The strengths of this technique include the use of inexpensive materials, variability in sandwich size, flexibility to re-open the sandwich or introduce multiple organisms through drilled holes, and the preservation and maintenance of phloem integrity. The phloem sandwich is an excellent educational tool for scientific discovery in both K-12 science courses and university research laboratories.

We provide a technique to preserve intact tree phloem and prepare it for observation. We create an apparatus called a phloem sandwich that allows for the introduction and observation of arthropods, microbes, and other organisms that inhabit phloem tissues.

Phloem tissues of pine are habitats for many thousands of organisms.  Arthropods and microbes use phloem and cambium tissues to seek mates, lay eggs, rear ...

Transient Gene Expression in Tobacco using Gibson Assembly and the Gene Gun

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately using complex regulatory strategies including differential promoters and/or signal sequences. Metabolic engineers and synthetic biologists interested in targeting enzymes to a particular organelle are faced with a challenge: For a protein that is to be localized to more than one organelle, the engineer must clone the same gene multiple times. This work presents a solution to this strategy: harnessing alternative splicing of mRNA. This technology takes advantage of established chloroplast and peroxisome targeting sequences and combines them into a single mRNA that is alternatively spliced. Some splice variants are sent to the chloroplast, some to the peroxisome, and some to the cytosol. Here the system is designed for multiple-organelle targeting with alternative splicing. In this work, GFP was expected to be expressed in the chloroplast, cytosol, and peroxisome by a series of rationally designed 5&#8217; mRNA tags. These tags have the potential to reduce the amount of cloning required when heterologous genes need to be expressed in multiple subcellular organelles. The constructs were designed in previous work11, and were cloned using Gibson assembly, a ligation independent cloning method that does not require restriction enzymes. The resultant plasmids were introduced into Nicotiana benthamiana epidermal leaf cells with a modified Gene Gun protocol. Finally, transformed leaves were observed with confocal microscopy.

This work describes a novel method for selectively targeting subcellular organelles in plants, assayed using the BioRad Gene Gun.

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately ...

LC-MS Analysis of Human Platelets as a Platform for Studying Mitochondrial Metabolism

Perturbed mitochondrial metabolism has received renewed interest as playing a causative role in a range of diseases. Probing alterations to metabolic pathways requires a model in which external factors can be well controlled, allowing for reproducible and meaningful results. Many studies employ transformed cellular models for these purposes; however, metabolic reprogramming that occurs in many cancer cell lines may introduce confounding variables. For this reason primary cells are desirable, though attaining adequate biomass for metabolic studies can be challenging. Here we show that human platelets can be utilized as a platform to carry out metabolic studies in combination with liquid chromatography-tandem mass spectrometry analysis. This approach is amenable to relative quantification and isotopic labeling to probe the activity of specific metabolic pathways. Availability of platelets from individual donors or from blood banks makes this model system applicable to clinical studies and feasible to scale up. Here we utilize isolated platelets to confirm previously identified compensatory metabolic shifts in response to the complex I inhibitor rotenone. More specifically, a decrease in glycolysis is accompanied by an increase in fatty acid oxidation to maintain acetyl-CoA levels. Our results show that platelets can be used as an easily accessible and medically relevant model to probe the effects of xenobiotics on cellular metabolism.

Here we show isolated human platelets can be used as an accessible ex vivo model to study metabolic adaptations in response to the complex I inhibitor rotenone. This approach employs isotopic tracing and relative quantification by liquid chromatography-mass spectrometry and can be applied to a variety of study designs.

Perturbed mitochondrial metabolism has received renewed interest as playing a causative role in a range of diseases. Probing alterations to metabolic ...

Measuring and Mapping Patterns of Soil Erosion and Deposition Related to Soil Carbonate Concentrations Under Agricultural Management

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonate (CaCO3) profiles. The objective is to describe a simple conceptual model and detailed protocol for repeatable field and laboratory measurements of these quantities. Here, accurate elevation is measured using a ground-based differential global positioning system (GPS); other data acquisition methods could be applied to the same basic method. Soil samples are collected from prescribed depth intervals and analyzed in the lab using an efficient and precise modified pressure-calcimeter method for quantitative analysis of inorganic carbon concentration. Standard statistical methods are applied to point data, and representative results show significant correlations between changes in soil surface layer CaCO3 and changes in elevation consistent with the conceptual model; CaCO3 generally decreased in depositional areas and increased in erosional areas. Maps are derived from point measurements of elevation and soil CaCO3 to aid analyses. A map of erosional and depositional patterns at the study site, a rain-fed winter wheat field cropped in alternating wheat-fallow strips, shows the interacting effects of water and wind erosion affected by management and topography. Alternative sampling methods and depth intervals are discussed and recommended for future work relating soil erosion and deposition to soil CaCO3.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonates. Repeatable methods for field and laboratory measurements of these quantities and data analysis methods are described here.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes ...

Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Life tables provide a means of measuring the schedules of birth and death from populations over time. They also can be used to quantify the sources and rates of mortality in populations, which has a variety of applications in ecology, including agricultural ecosystems. Horizontal, or cohort-based, life tables provide for the most direct and accurate method of quantifying vital population rates because they follow a group of individuals in a population from birth to death. Here, protocols are presented for conducting and analyzing cohort-based life tables in the field that takes advantage of the sessile nature of the immature life stages of a global insect pest, Bemisia tabaci. Individual insects are located on the underside of cotton leaves and are marked by drawing a small circle around the insect with a non-toxic pen. This insect can then be observed repeatedly over time with the aid of hand lenses to measure development from one stage to the next and to identify stage-specific causes of death associated with natural and introduced mortality forces. Analyses explain how to correctly measure multiple mortality forces that act contemporaneously within each stage and how to use such data to provide meaningful population dynamic metrics. The method does not directly account for adult survival and reproduction, which limits inference to dynamics of immature stages. An example is presented that focused on measuring the impact of bottom-up (plant quality) and top-down (natural enemies) effects on the mortality dynamics of B. tabaci in the cotton system.

Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Life tables allow quantification of the sources and rates of mortality in insect populations and contribute to understanding, predicting and manipulating population dynamics in agroecosystems. Methods for conducting and analyzing cohort-based life tables in the field for an insect with sessile immature life stages are presented.

Life tables provide a means of measuring the schedules of birth and death from populations over time. They also can be used to quantify the sources and ...

Techniques for Investigating the Anatomy of the Ant Visual System

This article outlines a suite of techniques in light microscopy (LM) and electron microscopy (EM) which can be used to study the internal and external eye anatomy of insects. These include traditional histological techniques optimized for work on ant eyes and adapted to work in concert with other techniques such as transmission electron microscopy (TEM) and scanning electron microscopy (SEM). These techniques, although vastly useful, can be difficult for the novice microscopist, so great emphasis has been placed in this article on troubleshooting and optimization for different specimens. We provide information on imaging techniques for the entire specimen (photo-microscopy and SEM) and discuss their advantages and disadvantages. We highlight the technique used in determining lens diameters for the entire eye and discuss new techniques for improvement. Lastly, we discuss techniques involved in preparing samples for LM and TEM, sectioning, staining, and imaging these samples. We discuss the hurdles that one might come across when preparing samples and how best to navigate around them.

This article outlines a suite of techniques in light and electron microscopy to study the internal and external eye anatomy of insects. These include several traditional techniques optimized for work on ant eyes, detailed troubleshooting, and suggestions for optimization for different specimens and regions of interest.

This article outlines a suite of techniques in light microscopy (LM) and electron microscopy (EM) which can be used to study the internal and external eye ...

The Plant Infection Test: Spray and Wound-Mediated Inoculation with the Plant Pathogen Magnaporthe Grisea

Plants possess a powerful system to defend themselves against potential threats by pathogenic fungi. For agriculturally important plants, however, current measures to combat such pathogens have proved too conservative and, thus, not sufficiently effective, and they can potentially pose environmental risks. Therefore, it is extremely necessary to identify host-resistance factors to assist in controlling plant diseases naturally through the identification of resistant germplasm, the isolation and characterization of resistance genes, and the molecular breeding of resistant cultivars. In this regard, there is need to establish an accurate, rapid, and large-scale inoculation method to breed and develop plant resistance genes. The rice blast fungal pathogen Magnaporthe grisea causes severe disease symptoms and yield losses. Recently, M. grisea has emerged as a model organism for studying the mechanisms of plant-fungal pathogen interactions. Hence, we report the development of a plant virulence test method that is specific for M. grisea. This method provides for both spray inoculation with a conidial suspension and wounding inoculation with mycelium cubes or droplets of conidial suspension. The key step of the wounding inoculation method for detached rice leaves is to make wounds on plant leaves, which avoids any interference caused by host penetration resistance. This spray/wounding protocol contributes to the rapid, accurate, and large-scale screening of the pathotypes of M. grisea isolates. This integrated and systematic plant infection method will serve as an excellent starting point for gaining a broad perspective of issues in plant pathology.

The Plant Infection Test: Spray and Wound-Mediated Inoculation with the Plant Pathogen Magnaporthe Grisea

Here, we present a protocol to test plant virulence with the plant pathogen Magnaporthe grisea. This report will contribute to the large-scale screening of the pathotypes of fungi isolates and serve as an excellent starting point for understanding the resistant mechanisms of plants during molecular breeding.

Plants possess a powerful system to defend themselves against potential threats by pathogenic fungi. For agriculturally important plants, however, current ...

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 ...

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Although solitary bees provide crucial pollination services for wild and managed crops, this species-rich group has been largely overlooked in pesticide regulation studies. The risk of exposure to fungicide residues is likely to be especially high if the spray occurs on, or near host plants while the bees are collecting pollen to provision their nests. For species of Osmia that consume pollen from a select group of plants (oligolecty), the inability to use pollen from non-host plants can increase their risk factor for fungicide-related toxicity. This manuscript describes protocols used to successfully rear oligolectic mason bees, Osmia ribifloris sensu lato, from egg to prepupal stage within cell culture plates under standardized laboratory conditions. The in vitro-reared bees are subsequently used to investigate the effects of fungicide exposure and pollen source on bee fitness. Based on a 2 &#215; 2 fully crossed factorial design, the experiment examines the main and interactive effects of fungicide exposure and pollen source on larval fitness, quantified by prepupal biomass, larval developmental time, and survivorship. A major advantage of this technique is that using in vitro-reared bees reduces natural background variability and allows the simultaneous manipulation of multiple experimental parameters. The described protocol presents a versatile tool for hypotheses testing involving the suite of factors affecting bee health. For conservation efforts to be met with significant, lasting success, such insights into the complex interplay of physiological and environmental factors driving bee declines will prove to be critical.

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Fungicide sprays on flowering plants may expose solitary bees to high concentrations of pollen-borne fungicide residues. Using laboratory-based experiments involving in vitro-reared bee larvae, this study investigates the interactive effects of consuming fungicide-treated pollen derived from host and non-host plants.

Although solitary bees provide crucial pollination services for wild and managed crops, this species-rich group has been largely overlooked in pesticide ...

A Gnotobiotic System for Studying Microbiome Assembly in the Phyllosphere and in Vegetable Fermentation

Summary

Explore More Videos

Chapters in this video

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

Technique for Studying Arthropod and Microbial Communities within Tree Tissues

Transient Gene Expression in Tobacco using Gibson Assembly and the Gene Gun

LC-MS Analysis of Human Platelets as a Platform for Studying Mitochondrial Metabolism

Measuring and Mapping Patterns of Soil Erosion and Deposition Related to Soil Carbonate Concentrations Under Agricultural Management

Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Techniques for Investigating the Anatomy of the Ant Visual System

The Plant Infection Test: Spray and Wound-Mediated Inoculation with the Plant Pathogen Magnaporthe Grisea

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

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors