Name: exploring the root microbiome: extracting bacterial community data from the soil, rhizosphere, and root endosphere
Uploaded: 2018-05-02T14:30:00.000+00:00
Duration: 9 min 55 s
Description: Watch this Scientific Journal Video about Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere at JoVE.com

This work was funded by the USDA-ARS (CRIS 2030-21430-008-00D). TS is supported by the NSF Graduate Research Fellowship Program.

1. Collection and Separation of Root Endosphere, Rhizosphere, and Soil Samples
<ol>
	<li>Prior to entering the field, autoclave ultrapure water (at least 90 mL of water per sample) to sterilize. Prepare epiphyte removal buffer (at least 25 mL per sample) by adding 6.75 g of KH2PO4, 8.75 g of K2HPO4, and 1 mL of Triton X-100, to 1 L of sterile water. Sterilize the buffer using a vacuum filter with 0.2 &#181;m pore size.
	<ol>
		<li>For steps 1.2 to 1.5, wear clean gloves sterilized with ethanol at all times and replace the gloves between each sample to prevent contamination. Sterilize all equipment with 70% ethanol and wipe clean all equipment between samples. Before sampling, determine the optimal sampling depth for your experiment, and be consistent with all soil and root collections.</li>
	</ol>
	</li>
	<li>To collect bulk soil samples, use an ethanol-sterilized soil core collector to obtain soil that is free of plant roots by collecting a core approximately 23 to 30 cm from the base of the plant.</li>
	<li>Transfer the soil to a plastic bag, homogenize the soil by gentle shaking, and transfer an aliquot of the soil sample (approximately 600 mg) to fill one 2 mL tube. Immediately place the 2 mL tube on dry ice or flash freeze the tube in liquid N2 until ready to proceed with DNA extraction (step 2).
	<ol>
		<li>In some environments, the surrounding soil can contain plant material. In this case, use a sterilized 2 mm sieve to separate the plant debris from the soil prior to placing it in the plastic bag.</li>
	</ol>
	</li>
	<li>To collect the root and rhizosphere, use an ethanol-sterilized shovel to dig up the plant, taking care to obtain as much of the root as possible. Depth is dependent upon the plant. While small plants such as wheat can be removed by digging several centimeters, larger plants such as sorghum may require 30 cm or more. Gently shake off excess soil from the roots until there is approximately 2 mm of soil adhering to the root surface. 
	NOTE: Take care when working with small plants, with fragile roots, or in dry, high-clay content soils. Ideally, there should only be a thin layer of soil remaining on the roots after shaking. If large aggregates of soil remain, a rubber mallet can be used to dislodge the soil by gently hitting the base of the shoot. If the amount of soil remaining after this process exceeds or falls short of 2 mm, the approximate thickness should be noted.</li>
	<li>For large plants, use sterile scissors and/or shears to cut a representative subsection of roots and place a minimum of 500 mg of root tissue into a 50 mL conical vial. For smaller grasses, place the entire root system into the vial. Add enough epiphyte removal buffer to cover the roots, then immediately place the sample on dry ice or flash freeze the sample in liquid N2. 
	NOTE: Take care not to overfill the 50 mL conical vial, as it will make washing step difficult. There should be enough empty space such that the epiphyte buffer is able to flow to the bottom, surround the roots throughout the vial, and cover the top. Because some grasses have more root biomass than will fit into a 50 mL conical vial, a subsection of the roots should be collected. However, it should be noted that cutting the roots could lead to endophytic bacteria being washed out into the rhizosphere fraction, so breaking roots should be minimized. If samples are not processed immediately after returning to lab, they can be stored at -80 &#176;C.</li>
	<li>To separate the rhizosphere from the roots, thaw the root sample on ice, then sonicate the root samples at 4 &#176;C for 10 min with pulses of 160 W for 30 s, separated by 30 s. Transfer the roots into a chilled (4 &#176;C), clean 50 mL tube using sterile forceps. Do not dispose the original tube with buffer and soil, which is the rhizosphere fraction (Figure 1).</li>
	<li>Centrifuge the tube containing buffer and rhizosphere for 10 min at 4 &#176;C, 4,000 x g. Decant the supernatant, flash freeze the tube containing the rhizosphere fraction in liquid N2, and store the rhizosphere fraction at -80 &#176;C until ready to proceed with DNA extraction (step 2).</li>
	<li>To wash the roots, add approximately 20 mL of chilled (4 &#176;C) sterile water to the root fraction. Wash the root by shaking vigorously (by hand or mixer, for 15 - 30 s), and then drain the water.</li>
	<li>Repeat this step at least twice, until no soil remains on the root surface. If the DNA extraction (step 2) is not performed immediately, wrap the roots in sterile aluminum foil, flash freeze the roots in liquid N2, and store the samples at -80 &#176;C until ready to proceed with DNA extraction.</li>
</ol>
2. DNA Extraction
NOTE: Throughout steps 2 and 3, clean gloves sterilized with ethanol should be worn at all times and all work should be performed on a surface sterilized with ethanol.
<ol>
	<li>Extract DNA from the soil and rhizosphere samples.
	<ol>
		<li>Use a sterile spatula to quickly transfer 250 mg of soil and rhizosphere from steps 1.3 and 1.7 into separate collection tubes provided in a commercial DNA isolation kit designed for extraction from soil, then proceed with DNA isolation using the kit supplier&#39;s protocol.</li>
		<li>After eluting the DNA in the elution buffer supplied by the DNA isolation kit, store the DNA at -20 &#176;C until ready to proceed with step 3.</li>
	</ol>
	</li>
	<li>Extract DNA from the root samples.
	<ol>
		<li>Chill a sterilized mortar and pestle using liquid N2. Measure out 600 to 700 mg of root tissue and place the tissue into the mortar. Carefully, add enough liquid N2 to cover the roots.</li>
		<li>Grind the roots into small pieces. Continue the process of adding liquid N2 and grinding (at least two times, be consistent between samples), until the roots are a fine powder. Ensure that the root tissue does not thaw during this step. 
		Caution: Use appropriate personal protective equipment (lab coat, protective eyewear, and cryogenic gloves) when working with liquid N2. 
		NOTE: For a low-quality DNA extraction, it can be beneficial to grind excess roots into powder and store the powder at -80 &#176;C.</li>
		<li>Quickly, before the root powder begins to thaw, use a sterile spatula to transfer the root powder into pre-weighed 1.5 mL tubes on ice. Record the weight of the tube and powder. Typically, 300 - 400 mg of powder is transferred.</li>
		<li>Use a sterile spatula to quickly transfer 150 mg of root powder to the collection tube provided in a commercial DNA isolation kit designed for extraction from soil, then proceed with DNA isolation using the kit supplier&#39;s protocol. 
		NOTE: For some root samples, there can be a high concentration of organics remaining in the DNA pellet, which prevents the amplification of the DNA during PCR, especially when a different DNA extraction protocol (e.g., CTAB extraction) is used. If necessary, clean the DNA by following the instructions provided in the environmental DNA clean-up kit.</li>
	</ol>
	</li>
	<li>Measure the concentration of all DNA samples using a high-sensitivity benchtop fluorometer.
	<ol>
		<li>Add 1 - 20 &#181;L of each eluted DNA sample into tubes provided in the dsDNA high-sensitivity assay kit. Add fluorometer working solution (1:200 dye:buffer) up to 200 &#181;L.</li>
		<li>Prepare two additional tubes containing 10 &#181;L of DNA standard 1 (0 ng/&#181;L DNA) or 10 &#181;L of standard 2 (100 ng/&#181;L), and add 190 &#181;L of fluorometer working solution to each standard.</li>
		<li>Measure the concentration of the standards and each sample. If it is not done automatically, calculate the DNA concentration from the absorbance output by a linear regression of the two standards.</li>
	</ol>
	</li>
</ol>
3. Amplicon Library Preparation and Submission
<ol>
	<li>Set up materials for the amplification reaction.
	<ol>
		<li>Thaw DNA samples at 4 &#176;C and keep them on ice throughout step 3. Randomize the order of DNA samples to minimize bias due to the location on the PCR plate (Table 2.)</li>
		<li>In a 96-well PCR plate, dilute DNA from each sample in molecular-grade water to 5 ng/&#181;L in a total volume of 20 &#181;L. Add 20 &#181;L of molecular-grade water to the four corner wells as negative controls for amplification (blanks) (Table 2).</li>
		<li>Arrange the barcoded primers (10 &#181;M) in either PCR strip tubes or a 96-well plate such that they can be added with a multi-channel pipette (Figure 2).</li>
		<li>Prepare sufficient PCR master mix to amplify each DNA sample in triplicate. Prepare 1.5 &#181;L of BSA (20 mg/mL), 37.5 &#181;L of pre-made 2x master mix (composed of PCR buffer, MgCl2, dNTPs, and&#160;Taq&#160;DNA polymerase), 0.57 &#181;L of chloroplast PNA (100 &#181;M), 0.57 &#181;L of mitochondrial PNA (100 &#181;M), and 25.86 &#181;L of molecular grade water.</li>
		<li>Pour the master mix into a sterile 25 mL of multichannel pipette reservoir and distribute 66 &#181;L of master mix into each well of a new 96-well PCR plate using a multichannel pipette. 
		NOTE: When calculating reagent volumes for the master mix, make sure to also include the 4 blank wells per plate.</li>
		<li>Using a multi-channel pipette, add 6 &#181;L of 5 ng/&#181;L DNA (from the normalized DNA plate) to the master mix plate. Then add to the master plate 1.5 &#181;L of 10 &#181;M forward primer such that each column has a different forward barcode, and 1.5 &#181;L of 10 &#181;M reverse primer such that each row has a different reverse barcode (Figure 2). 
		NOTE: Prior to adding primers, the randomized plates and master mix could be used to amplify the ITS or ITS2 fungal genes if different primers were added. If this is the case, a similar primer design can be used.</li>
		<li>Spin down the plate briefly at 3,000 x g. Use a multi-channel pipette to mix gently, then divide into three plates with 25 &#181;L of reaction mixture. 
		NOTE: Although three replicates are not strictly necessary, it decreases the impact of technical variability.</li>
	</ol>
	</li>
	<li>Amplify the DNA in each plate using a thermocycler set to the following conditions: 180 s at 98 &#176;C, 30 cycles of: 98 &#176;C for 45 s (denaturing), 78 &#176;C for 10 s (PNA annealing), 55 &#176;C for 60 s (primer annealing), and 72 &#176;C for 90 s (extension), then 600 s at 72 &#176;C followed by a 4 &#176;C hold step. After the amplification, pool the three replicate plates into one single 96-well plate.</li>
	<li>Quantify the DNA using high-sensitivity fluorometer reagents in a 96-well plate reader.
	<ol>
		<li>Add 2 &#181;L of each PCR product to a 96-well microplate, along with 98 &#181;L of fluorometer working solution (1:200 dye:buffer). Include 4 wells as standards: 5 &#181;L of DNA standard 1 (0 ng/&#181;L DNA), 1 &#181;L of standard 2 (10 ng/&#181;L), 2 &#181;L of standard 2 (20 ng/&#181;L), and 5 &#181;L of standard 2 (50 ng/&#181;L). Then add fluorometer working solution for a final volume of 100 &#181;L. 
		NOTE: Each sample can be measured using a benchtop fluorometer as described in step 2.3 if a plate reader is not available.</li>
		<li>Calculate the DNA concentration from the absorbance output by a linear regression of the four standards.</li>
		<li>For the successfully amplified barcoded products (those that have a concentration greater than 15 ng/&#181;L), pool 100 ng of each sample into a single 1.5 mL tube (Table 2).</li>
		<li>Calculate the average volume of samples added to the pool by using the =AVERAGE() function in a spreadsheet program. Add the volume of the &#34;blank&#34; PCR products to the pooled samples. 
		NOTE: Since the &#34;blank&#34; PCR products have their own unique barcode combinations, they can be sequenced to check for any laboratory contaminants.</li>
	</ol>
	</li>
	<li>Measure the concentration of the pooled product using a benchtop fluorometer as described in step 2.3, and take 600 ng of DNA and dilute in molecular-grade water to a final volume of 100 &#181;L in a 1.5 mL tube. Store the remaining pooled product at -20 &#176;C.</li>
	<li>Wash the 600 ng DNA aliquot by following the established PCR purification process with paramagnetic purification beads in a 96-well format with a few exceptions.
	<ol>
		<li>Make a fresh 600 &#181;L aliquot of 70% ethanol. Shake the bottle of magnetic beads to re-suspend beads that settle to the bottom.</li>
		<li>Add 1x volume (100 &#181;L) of bead solution to the 600 ng aliquot of DNA. Mix thoroughly by pipetting 10 times. Incubate for 5 min at room temperature.</li>
		<li>Place the tube onto the magnetic stand for 2 min (or until solution is clear) to separate beads from solution. While the tube is still in the magnetic stand, aspirate the clear supernatant carefully without touching the magnetic beads, and discard the clear supernatant. 
		NOTE: At this point, the amplicon products are bound to the magnetic beads. Any beads that are disturbed or lost during aspiration will result in a loss of DNA.</li>
		<li>Leave the tube in the magnetic stand and add 300 &#181;L of 70% ethanol to the tube; incubate at room temperature for 30 s. Aspirate out the ethanol and discard. Repeat this process, and remove all ethanol after the second wash. Remove the tube from the magnetic stand, and air dry for 5 min.</li>
		<li>Add 30 &#181;L of molecular-grade water to the dried beads and mix by pipetting 10 times. Incubate at room temperature for 2 min. Return the tube to the magnetic stand for 1 min to separate the beads from solution. Transfer the eluate to a new tube. 
		NOTE: Magnetic beads will not affect downstream reactions.</li>
	</ol>
	</li>
	<li>Measure the final concentration of cleaned, pooled DNA using a benchtop fluorometer as described in step 2.3. Dilute an aliquot to 10 nM in a final volume of 30 &#181;L, or to the concentration and volume preferred by the sequencing facility.</li>
	<li>Utilize the services of a sequencing facility to sequence the DNA on a NGS platform, 2 x 300 bp paired-end sequencing.</li>
</ol>

The authors have nothing to disclose.

This protocol demonstrates an established pipeline for exploring root endosphere, rhizosphere, and soil microbial community compositions, from field sampling to sample processing and downstream sequencing. Studying root-associated microbiomes presents unique challenges, due in part to the inherent difficulties in sampling from soil. Soils are highly variable in terms of physical and chemical properties, and different soil conditions can be separated by as little as a few millimeters28,29. This can lead to the samples which are collected from adjacent sampling sites having considerably different microbial community compositions and activities30,31. Thus, using soil core collectors and shovels to maintain consistent sampling depths and homogenization prior to DNA extraction are essential to the reproducibility within root microbiome studies. It is also essential to efficiently separate the rhizosphere and root fractions; using a harsh method of root surface sterilization can potentially lyse endophytes within roots prior to DNA extraction, while a more conservative wash may not remove all microbes from the root surface7. Another key factor that can negatively impact or disrupt sequencing results is bacterial contamination, which can come from many sources and is sometimes impossible to distinguish from the sampled environmental bacteria32,33. For this reason, careful sterilization of sampling tools, experimental materials, and working environments are vital in order to avoid contamination.
After sampling, obtaining high quality DNA is a high priority for successful downstream analyses. In our experience, DNA extraction from field grown root samples through alternative methods, such as through CTAB-based extraction, often contain substantially greater quantities of humic acids and other compounds compared to rhizosphere and soil samples. These compounds can prevent the enzymatic activity of the DNA polymerase during PCR amplification, even at low concentrations34,35. Using DNA extraction kits designed for soils on root samples, as opposed to a CTAB extraction followed by a phenol chloroform clean-up, can effectively rid samples of humic acids and will result in high quality DNA36,37,38,39. Accordingly, we recommend using a commercially available DNA extraction kit for root samples as well. It should be noted that the goal is to obtain microbial genomic DNA from plant roots. Thus, thorough and consistent root grinding is important to break down the plant tissue and lyse the microbial cells to release microbial DNA without introducing bias between samples due to the variation in grinding pressure and time.
Following careful extraction of DNA from samples, there are two main sources for problems during amplification: 1) contamination of plant tissues with plant endosymbionts (chloroplast and mitochondria) and 2) selection of 16S rRNA region to amplify. The amplification from chloroplast or mitochondria 16S rRNA sequences can generate &#62;80% of the sequences in root samples40, and more in leaf tissues, though the amount of contamination is dependent on the choice of primers. Thus, PNA clamps are necessary during the PCR step to suppress plant host chloroplast and mitochondrial 16S contamination27,41. However, different plant species can have variation in the chloroplast and mitochondrial 16S sequence27; therefore, it is essential to confirm the sequence of the chloroplast and mitochondrial 16S genes of the plant being studied prior to library sequencing, in order to determine if alternate PNA oligos are needed (Figure 3). Additionally, the 16S rRNA gene consists of nine hypervariable regions flanked by nine conservative regions; different results can be obtained from the same community depending upon which hypervariable region is amplified42. Previous studies have found the V4 region to be one of the most reliable for assigning taxonomy43 and it has been used for other extensive microbiome surveys11. Lengthening the target to the V3-V4 region is suggested here to increase variability and improve taxonomic resolution.
In this protocol, we demonstrated a pipeline to perform 16S rRNA amplicon sequencing via next generation sequencing (NGS) for studying microbial community compositions of environmental samples12. We recommend using amplicon sequencing as a tool for phylogenetic profiling, because it is relatively inexpensive, high-throughput, and does not require extensive computational expertise or resources to analyze. While our method focuses on analyzing the bacterial fraction of the microbiome, it can easily be adapted to investigate fungi. The protocol is identical through step 2, and the only difference in step 3 is what primers would be used during the amplification. However, it is worth nothing that amplicon based profiling is not without limitations. By sequencing a single marker gene, no information is obtained regarding the functional capacity of the community. Additionally, the taxonomic resolution can be quite low, especially when sequencing from environments with a high percentage of uncharacterized microbes. However, sequencing technologies are rapidly evolving, and we anticipate the potential to deal with some of these shortcomings by adapting this protocol for use with other sequencing platforms. Finally, as mentioned in the introduction, shotgun metagenomics and metatranscriptomics can easily be performed on soil and rhizosphere samples, and methods to eliminate plant contamination from plant tissues are currently being explored. Experimental designs which pair amplicon-based approaches and other metagenomic techniques can be particularly effective in complex communities where high species diversity and uneven representation of taxa can prevent shotgun data from accurately characterizing the less dominant members.

Plant-associated microbiomes consist of dynamic and complex microbial communities comprised of bacteria, archaea, viruses, fungi, and other eukaryotic microorganisms. In addition to their well-studied role in causing plant disease, plant-associated microbes can also positively influence plant health by improving tolerance to biotic and abiotic stresses, promoting nutrient availability, and enhancing plant growth through the production of phytohormones. For this reason, particular interest exists in characterizing the taxa that associate with plant root endospheres, rhizospheres, and the surrounding soil. While some microbes can be cultured in isolation on laboratory generated media, many cannot, in part because they may rely on symbiotic relationships with other microbes, grow very slowly, or require conditions that cannot be replicated in a lab environment. Because it circumvents the need for cultivation and is relatively inexpensive and high-throughput, sequence-based phylogenetic profiling of environmental and host-associated microbial samples has become a preferred method for assaying microbial community composition.
The selection of appropriate sequencing technologies provided by various next generation sequencing (NGS) platforms1 is dependent on the users&#39; needs, with important factors including: desired coverage, amplicon length, expected community diversity, as well as sequencing error-rate, read-length, and the cost-per-run/megabase. Another variable that needs to be considered in amplicon-based sequencing experiments is what gene will be amplified and what primers will be used. When designing or choosing primers, researchers are often forced to make tradeoffs between the universality of amplification and the taxonomic resolution achievable from the resulting amplicons. For this reason, these types of studies often chose primers and markers that selectively target specific subsets of the microbiome. Evaluating the composition of bacterial communities is commonly accomplished by sequencing one or more of the hypervariable regions of the bacterial 16S rRNA gene2,3. In this study, we describe an amplicon based sequencing protocol developed for a NGS platform that targets the 500 bp V3-V4 region of the bacterial 16S rRNA gene, which allows for broad amplification of bacterial taxa while also providing sufficient variability to distinguish between different taxa. Additionally, this protocol can easily be adapted for the use with other primer sets, such as those targeting the ITS2 marker of fungi or the 18S rRNA subunit of eukaryotes.
While other approaches such as shotgun metagenomics, metatranscriptomics, and single-cell sequencing, offer other advantages including resolved microbial genomes and more direct measurement of community function, these techniques are typically more expensive and computationally intensive than the phylogenetic profiling described here4. Additionally, performing shotgun metagenomics and metatranscriptomics on root samples yields a large percentage of reads belonging to the host plant genome, and methods to overcome this limitation are still being developed5,6.
As with any experimental platform, amplicon-based profiling can introduce a number of potential biases which should be considered during the experimental design and data analysis. These include the methods of sample collection, DNA extraction, selection of PCR primers, and how library preparation is performed. Different methods can significantly impact the amount of usable data generated, and can also hinder the efforts to compare results between studies. For example, the method of removing rhizosphere bacteria7 and the use of different extraction techniques or choice of DNA extraction kits8,9 have been shown to significantly impact downstream analysis, which leads to different conclusions regarding which microbes are present and their relative abundances. Since amplicon-based profiling can be customized, making comparisons across studies can be challenging. The Earth Microbiome Project has suggested that researchers studying complex systems such as the plant-associated microbiome would benefit from the development of standardized protocols as a means of minimizing the variability caused by the application of different methods between studies10,11. Here, we discuss many of the above topics and offer suggestions as to best practices where appropriate.
The protocol demonstrates the process of collecting soil, rhizosphere, and root samples from Sorghum bicolor and extracting DNA using a well-established DNA isolation kit11. Additionally, our protocol includes a detailed amplicon sequencing workflow, using a commonly utilized NGS platform, to determine the structure of the bacterial communities12,13,14. This protocol has been validated for the use in a wide range of plant hosts in a recent published study of the roots, rhizosphere, and associated-soils of 18 monocot species including Sorghum bicolor, Zea mays, and Triticum aestivum15. This method has also been validated for use with other marker genes, as demonstrated by its successful application to studying the fungal ITS2 marker gene in studies of the agave microbiome16,17 and strawberry microbiome18.

<table><tbody><tr><td>0.1-10/20 &amp;micro;L filtered micropipette tips</td><td>USA Scientific</td><td>1120-3810</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>1.5 mL microcentrifuge tubes</td><td>USA Scientific</td><td>1615-5510</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>10 &amp;micro;L multi-channel pipette</td><td>Eppendorf</td><td>3122000027</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>10 &amp;micro;L, 100 &amp;micro;L, and 1000 &amp;micro;L micropipettes</td><td>Eppendorf</td><td>3120000909</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>100 &amp;micro;L multi-channel pipette</td><td>Eppendorf</td><td>3122000043</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>1000 &amp;micro;L filtered micropipette tips</td><td>USA Scientific</td><td>1122-1830</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>2 mL microcentrifuge tubes</td><td>USA Scientific</td><td>1620-2700</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>2 mm soil sieve</td><td>Forestry Suppliers</td><td>60141009</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>200 &amp;micro;L filtered micropipette tips</td><td>USA Scientific</td><td>1120-8810</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>25 mL reservoirs</td><td>VWR International LLC</td><td>89094-664</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>50 mL conical vials</td><td>Thermo Fisher Scientific</td><td>352098</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>500 mL vacuum filters (0.2 &amp;micro;m pore size)</td><td>VWR International LLC</td><td>156-4020</td><td></td></tr><tr><td>96-well microplates</td><td>USA Scientific</td><td>655900</td><td></td></tr><tr><td>96-well PCR plates</td><td>BioRad</td><td>HSP9631</td><td></td></tr><tr><td>Agencourt AMPure XP beads</td><td>Thermo Fisher Scientific</td><td>NC9933872</td><td>Instructions for use:&lt;br/&gt; https://www.beckmancoulter.com/wsrportal/ajax/downloadDocument/B37419AA.pdf?autonomyId=TP_DOC_150180&lt;br/&gt;&amp;amp;documentName=B37419AA.pdf</td></tr><tr><td>Aluminum foil</td><td>Boardwalk</td><td>7124</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Analytical scale with 0.001 g resolution</td><td>Ohaus Pioneer</td><td>PA323</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Bioruptor Plus ultrasonicator</td><td>Diagenode</td><td>B01020001</td><td></td></tr><tr><td>Bovine Serum Albumin (BSA) 20 mg/mL</td><td>New England Biolabs</td><td>B9000S</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5811000908</td><td>Including 50mL and 96-well plate bucket adapters</td></tr><tr><td>Cryogenic gloves</td><td>Millipore Sigma</td><td>Z183490</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>DNeasy PowerClean kit (optional)</td><td>Qiagen Inc.</td><td>12877-50</td><td>Previously MoBio</td></tr><tr><td>DNeasy PowerSoil kit</td><td>Qiagen Inc.</td><td>12888-100</td><td>Previously MoBio</td></tr><tr><td>Dry ice</td><td>Any</td><td>NA</td><td></td></tr><tr><td>DynaMag-2 magnet</td><td>Thermo Fisher Scientific</td><td>12321D</td><td>Do not substitute</td></tr><tr><td>Ethanol</td><td>VWR International LLC</td><td>89125-188</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Gallon size freezer bags</td><td>Ziploc</td><td>NA</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Gemini EM Microplate Reader</td><td>Molecular Devices</td><td>EM</td><td>Can use another fluorometer that reads 96-well plates from the top.</td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>P3786</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>P5655</td><td></td></tr><tr><td>Lab coat</td><td>Workrite</td><td>J1367</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Liquid N&lt;sub&gt;2&lt;/sub&gt;</td><td>Any</td><td>NA</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Liquid N&lt;sub&gt;2&lt;/sub&gt; dewar</td><td>Thermo Fisher Scientific</td><td>4150-9000</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Milli-Q ultrapure water purification system</td><td>Millipore Sigma</td><td>SYNS0R0WW</td><td></td></tr><tr><td>Mini-centrifuge</td><td>Eppendorf</td><td>5404000014</td><td></td></tr><tr><td>Molecular grade water</td><td>Thermo Fisher Scientific</td><td>4387937</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Mortars</td><td>VWR International LLC</td><td>89038-150</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Nitrile gloves</td><td>Thermo Fisher Scientific</td><td>19167032B</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Paper towels</td><td>VWR International LLC</td><td>BWK6212</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>PCR plate sealing film</td><td>Thermo Fisher Scientific</td><td>NC9684493</td><td></td></tr><tr><td>PCR strip tubes</td><td>USA Scientific</td><td>1402-2700</td><td></td></tr><tr><td>Pestles</td><td>VWR International LLC</td><td>89038-166</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Plastic spatulas</td><td>LevGo, Inc.</td><td>17211</td><td></td></tr><tr><td>Platinum Hot Start PCR Master Mix (2x)</td><td>Thermo Fisher Scientific</td><td>13000014</td><td></td></tr><tr><td>PNAs - chloroplast and mitochondrial</td><td>PNA Bio</td><td>NA</td><td>Make sure to verify sequence bioinformatically</td></tr><tr><td>Protective eyewear</td><td>Millipore Sigma</td><td>Z759015</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Qubit 3.0 Fluorometer</td><td>Thermo Fisher Scientific</td><td>Q33216</td><td></td></tr><tr><td>Qubit dsDNA HS assay kit</td><td>Thermo Fisher Scientific</td><td>Q32854</td><td></td></tr><tr><td>Rubber mallet (optional)</td><td>Ace Hardware</td><td>2258622</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Shears or scissors</td><td>VWR International LLC</td><td>89259-936</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Shovel</td><td>Home Depot</td><td>2597400</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Soil core collector (small diameter: &amp;lt;1 inch)</td><td>Ben Meadows</td><td>221700</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Spray bottles</td><td>Santa Cruz Biotechnology</td><td>sc-395278</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Standard desalted barcoded primers (10 &amp;micro;M) (Table 1)</td><td>IDT</td><td>NA</td><td>4 nmole Ultramer DNA Oligo with standard desalting. NGS adapter and sequencing primer (Table 1) are designed for use with Illumina MiSeq using v3 chemistry.</td></tr><tr><td>Thermocycler</td><td>Thermo Fisher Scientific</td><td>E950040015</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100</td><td>Can substitute with equivalent from other suppliers.</td></tr><tr><td>Weigh boats</td><td>Spectrum Chemicals</td><td>B6001W</td><td>Can substitute with equivalent from other suppliers.</td></tr></tbody></table>

exploring the root microbiome: extracting bacterial community data from the soil, rhizosphere, and root endosphere

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to abiotic stresses and diseases. As the plant microbiome can be highly complex, low-cost, high-throughput methods such as amplicon-based sequencing of the 16S rRNA gene are often preferred for characterizing its microbial composition and diversity. However, the selection of appropriate methodology when conducting such experiments is critical for reducing biases that can make analysis and comparisons between samples and studies difficult. This protocol describes in detail a standardized methodology for the collection and extraction of DNA from soil, rhizosphere, and root samples. Additionally, we highlight a well-established 16S rRNA amplicon sequencing pipeline that allows for the exploration of the composition of bacterial communities in these samples, and can easily be adapted for other marker genes. This pipeline has been validated for a variety of plant species, including sorghum, maize, wheat, strawberry, and agave, and can help overcome issues associated with the contamination from plant organelles.

Performing the recommended protocol should result in a dataset of indexed paired-end reads that can be matched back to each sample and assigned to either a bacterial operational taxonomic units (OTU) or exact sequence variant (ESV, also referred to as amplicon sequence variant (ASV) and sub-operational taxonomic unit (sOTU)), depending on downstream analysis. In order to obtain high-quality sequence data, care must be taken at each step to maintain consistency between samples and minimize the introduction of any potential bias during the sample processing or library preparation. After collecting, processing, and extracting DNA from samples (steps 1 and 2), the resulting eluate should appear clear and free of organics that would inhibit amplification. While purity can be verified by measuring each DNA sample via a microvolume spectrophotometer, we have found that the soil DNA extraction kit reliably removes all contaminants. As a result of the predictable DNA quality, quantification methods that rely on fluorescence-based dyes that specifically bind DNA are more appropriate than those based on UV absorbance19,20,21. Prior to PCR amplification, soil and rhizosphere samples average around 10 ng/&#181;L DNA, while root samples typically have a mean concentration of approximately 30 ng/&#181;L (Table 2).
Following the amplification of the environmental DNA (step 3), success or failure can be determined by measuring the concentration of the PCR product via benchtop fluorometer reagents on a plate reader, if available, or manually (Table 2). In our experience, successful amplifications that result in high-quality amplicon data yield greater than 15 ng/&#181;L PCR products. If there are multiple failures on a plate, the positional arrangement within the plant and the sample type of failed samples may help determine the problem. For instance, if they are all adjacent on the plate, it may indicate pipette error, whereas if they are all in the same row or column, it could suggest issues with a specific primer. If they all belong to the same sample type, it might suggest problems with sample processing or DNA extraction.
It is important to check the compatibility of the universal PNAs with your specific plant system bioinformatically during experimental design in order to verify that they will block amplification of chloroplast and mitochondrial 16S genes. Following the amplification step, it is not clear whether the PNAs successfully bound to mitochondrial and chloroplast templates; this is only revealed after sequencing (Figure 3). To help ensure that the PNAs will effectively block contaminant amplification, an alignment of the PNA sequence to each chloroplast and mitochondrial 16S rRNA gene (there may be multiple copies) for the plant host being investigated should not reveal any mismatches. Even a single mismatch to the 13 bp PNA sequence, especially in the middle of the PNA clamp, can drastically reduce the effectiveness, as in the case of the provided chloroplast PNA sequence and the chloroplast 16S rRNA gene of Lactuca sativa (lettuce) (Figure 3).
Since an equal amount of amplified DNA is pooled per sample, there should be an approximately even number of reads obtained per sample after sequencing and sorting reads based on their barcoded index (Figure 4). The majority of these reads should match to bacterial taxa. Any eukaryotic, mitochondrial, or chloroplast matches should be discarded. Depending on the analysis pipeline and taxonomic database chosen, chloroplast and mitochondrial reads can mistakenly be assigned to bacterial lineages, often Cyanobacteria and Rickesttia, respectively (Figure 3). A degree of manual curation is often prudent to check for these common mis-assignments. Specific details will depend on the choice of analysis, but relative abundance profiles should generally be similar (no significant difference) among biological replicates and significantly different between soil, rhizosphere, and root samples (Figure 5). It is important to note, however, that while there may be no significant difference between biological replicates, it is important to collect at least three replicates per.
Methods for interpreting the data obtained in these experiments are hotly debated amongst microbial ecologists. Until recently, amplicon sequence analysis has been dependent upon grouping reads into OTUs. However, these are problematic because: 1) they are based on a somewhat arbitrary threshold of 97% similarity, 2) diversity is often underestimated, and 3) there can be low taxonomic resolution. Recently developed tools such as DADA2, Deblur, and UNOISE222,23,24 are able to sort reads into ESVs, which solves some problems presented when using OTUs. Caveats to using ESVs include: 1) artificial increases in diversity due to the differences in rRNA copies within a species, and 2) increased sensitivity to PCR and sequencing errors25,26.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57561/57561fig1.jpg" /> 
Figure 1: Separation of root and rhizosphere fractions. Flowchart displaying the steps for separating the rhizosphere from the root samples, followed by washing the roots with sterile water to remove any remaining rhizoplane organisms. <a href="/files/ftp_upload/57561/57561fig1large.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/57561/57561fig2.jpg" /> 
Figure 2: Example of stock primer layout for amplification and distribution within plates. Stock primers (Table 1) can be prepared in strip tubes for optimal distribution within 96-well plates (each strip of primers is represented by a different color; purple for forward primers 1 - 8, orange for forward primers 9 - 16, blue for reverse primers 1 - 12, and green for reverse primers 13 - 16.) In this case, 16 forward and 16 reverse primers can be distributed efficiently with a multi-channel pipette such that each well has a unique barcode combination. <a href="/files/ftp_upload/57561/57561fig2large.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/57561/57561fig3.jpg" /> 
Figure 3: Results that suggest chloroplast PNA is ineffective. Representative result from rhizosphere (&#34;Rhizo&#34;), root, and soil samples from lettuce that were non-treated (NT) or treated (VT) with a biological soil amendment. The PNA sequence used to block chloroplast contamination of most plants is GGCTCAACCCTGGACAG27. However, lettuce contains a mismatch in the chloroplast 16S ribosomal RNA gene (GGCTCAACTCTGGACAG). This renders the PNA ineffective, resulting in a high relative abundance of reads that match to Cyanobacteria in rhizosphere and root samples. <a href="/files/ftp_upload/57561/57561fig3large.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/57561/57561fig4.jpg" /> 
Figure 4: Distribution of read counts among samples in a library. Bar chart showing number of read counts (y-axis) from different samples (bars, x-axis), matched by the barcode combination in the read. The number of reads per sample can vary based on how many samples are in the library; this subset was sequenced in a library of 192 samples. <a href="/files/ftp_upload/57561/57561fig4large.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/57561/57561fig5.jpg" /> 
Figure 5: Relative abundance of the top 12 classes in root, rhizosphere, and soil communities. Stacked bar chart showing relative abundance of classes present in a representative 16S dataset containing 6 replicates for each sample type (bulk soil, rhizosphere, and root endosphere). <a href="/files/ftp_upload/57561/57561fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Primers for amplifying the V3-V4 region of the 16S rRNA gene. Primers are composed of, sequentially: an adapter for a common NGS platform, a unique barcode, the primer for NGS, a spacer region of variable length to shift the frame for sequencing, and a universal PCR primer that amplifies either 341F or 785R of the 16S rRNA gene. The number of primers needed is dependent upon how many samples are sequenced per library; a combination of 16 forward and 16 reverse primers is sufficient for 244 samples (256 primer combinations with 12 used for blank wells during PCR (Figure 2)). <a href="/files/ftp_upload/57561/Table_1.xlsx">Please click here to download this file.</a>
Table 2: Normalization of randomized DNA samples prior to and following amplification. Example worksheet listing samples in a randomized order and indicating their location on a single 96-well plate, which also determines the primer combination assigned to it. Formulas in the bottom row describe calculations for adding 100 ng of each sample to the normalized plate, plus the volume of water to reach 20 &#181;L. Following amplification, the volume of 100 ng of each successful product is calculated and added to a final pool. The volume of &#34;blank&#34; PCR product to add to the final pool is the average of the other samples. <a href="/files/ftp_upload/57561/Table_2.xlsx">Please click here to download this file.</a>
Supplemental Figure 1: Approximation of minimum root biomass during sample collection. When collecting roots, try to collect at least 500 mg of tissue. Here, roots collected from a young sorghum plant (left, in both A and B) and a young rice plant (right) are shown next to (A) and inside (B) 50 mL conical tubes. Both samples weigh approximately 1 g, however, it is important to note that this weight includes rhizosphere and root, and the rhizosphere weight is, in this case, approximately half the total weight. <a href="/files/ftp_upload/57561/Supplemental_Figure_1.tiff">Please click here to download this file.</a>

Watch this Scientific Journal Video about Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere at JoVE.com

Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere and Root Endosphere | Genetics | Video

Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere

Here, we describe a protocol to obtain amplicon sequence data of soil, rhizosphere, and root endosphere microbiomes. This information can be used to investigate the composition and diversity of plant-associated microbial communities, and is suitable for the use with a wide range of plant species.

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to ...

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26.3K Views.  University of California, Berkeley. Hier beschrijven we een protocol om amplicon reeks gegevens van bodem, rhizosfeer en wortel endosphere microbiomes te verkrijgen. Deze informatie kan worden gebruikt voor het onderzoeken van de samenstelling en de diversiteit van de plant-geassocieerde microbiële gemeenschappen, en is geschikt voor het gebruik met een breed scala van plantensoorten.

Video: Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere

Soil Lysimeter Excavation for Coupled Hydrological, Geochemical, and Microbiological Investigations

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled Earth-system processes. Such knowledge is imperative in improving predictions of hydro-biogeochemical cycles, especially under climate change scenarios. We present an experimental method, designed to capture sub-surface heterogeneity of an initially homogeneous soil system. This method is based on destructive sampling of a soil lysimeter designed to simulate a small-scale hillslope. A weighing lysimeter of one cubic meter capacity was divided into sections (voxels) and was excavated layer-by-layer, with sub samples being collected from each voxel. The excavation procedure was aimed at detecting the incipient heterogeneity of the system by focusing on the spatial assessment of hydrological, geochemical, and microbiological properties of the soil. Representative results of a few physicochemical variables tested show the development of heterogeneity. Additional work to test interactions between hydrological, geochemical, and microbiological signatures is planned to interpret the observed patterns. Our study also demonstrates the possibility of carrying out similar excavations in order to observe and quantify different aspects of soil-development under varying environmental conditions and scale.

This study presents an excavation method for investigating subsurface hydrological, geochemical, and microbiological heterogeneity of a soil lysimeter. The lysimeter simulates an artificial hillslope which was initially under homogeneous condition and had been subjected to approximately 5,000 mm of water over eight cycles of irrigation in an 18-month period.

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled ...

Environment

A Lipid Extraction and Analysis Method for Characterizing Soil Microbes in Experiments with Many Samples

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial communities, a relatively precise but effort-intensive technique to assay microbial community composition is phospholipid fatty acid (PLFA) analysis. PLFA was developed to analyze phospholipid biomarkers, which can be used as indicators of microbial biomass and the composition of broad functional groups of fungi and bacteria. It has commonly been used to compare soils under alternative plant communities, ecology, and management regimes. The PLFA method has been shown to be sensitive to detecting shifts in microbial community composition.
An alternative method, fatty acid methyl ester extraction and analysis (MIDI-FA) was developed for rapid extraction of total lipids, without separation of the phospholipid fraction, from pure cultures as a microbial identification technique. This method is rapid but is less suited for soil samples because it lacks an initial step separating soil particles and begins instead with a saponification reaction that likely produces artifacts from the background organic matter in the soil. 
This article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples. The method combines the accuracy achieved through PLFA profiling by extracting and concentrating soil lipids as a first step, and a reduction in effort by saponifying the organic material extracted and processing with the MIDI-FA method as a second step.

The article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples.

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial ...

Isolation and Analysis of Microbial Communities in Soil, Rhizosphere, and Roots in Perennial Grass Experiments

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The purpose of this manuscript is to provide detail on how to rapidly sample soil, rhizosphere, and endosphere of replicated field trials and analyze changes that may occur in the microbial communities due to sample type, treatment, and plant genotype. The experiment used to demonstrate these methods consists of replicated field plots containing two, pure, warm-season grasses (Panicum virgatum and Andropogon gerardii) and a low-diversity grass mixture (A. gerardii, Sorghastrum nutans, and Bouteloua curtipendula). Briefly, plants are excavated, a variety of roots are cut and placed in phosphate buffer, and then shaken to collect the rhizosphere. Roots are brought to the laboratory on ice and surface sterilized with bleach and ethanol (EtOH). The rhizosphere is filtered and concentrated by centrifugation. Excavated soil from around the root ball is placed into plastic bags and brought to the lab where a small amount of soil is taken for DNA extractions. DNA is extracted from roots, soil, and rhizosphere and then amplified with primers for the V4 region of the 16S rRNA gene. Amplicons are sequenced, then analyzed with open access bioinformatics tools. These methods allow researchers to test how the microbial community diversity and composition varies due to sample type, treatment, and plant genotype. Using these methods along with statistical models, the representative results demonstrate there are significant differences in microbial communities of roots, rhizosphere, and soil. Methods presented here provide a complete set of steps for how to collect field samples, isolate, extract, quantify, amplify, and sequence DNA, and analyze microbial community diversity and composition in replicated field trials.

Excavation of plant roots from the field as well as processing of samples into endosphere, rhizosphere, and soil are described in detail, including DNA extraction and data analysis methods. This paper is designed to enable other laboratories to use these techniques for the study of soil, endosphere, and rhizosphere microbiomes.

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The ...

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 ...

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

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

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

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

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

Immunology and Infection

Divergence of Root Microbiota in Different Habitats based on Weighted Correlation Networks

The root microbiome plays an important role in plant growth and environmental adaptation. Network analysis is an important tool for studying communities, which can effectively explore the interaction relationship or co-occurrence model of different microbial species in different environments. The purpose of this manuscript is to provide details on how to use the weighted correlation network algorithm to analyze different co-occurrence networks that may occur in microbial communities due to different ecological environments. All analysis of the experiment is performed in the WGCNA package. WGCNA is an R package for weighted correlation network analysis. The experimental data used to demonstrate these methods were microbial community data from the NCBI (National Center for Biotechnology Information) database for three niches of the rice (Oryza sativa) root system. We used the weighted correlation network algorithm to construct co-abundance networks of microbial community in each of the three niches. Then, differential co-abundance networks among endosphere, rhizoplane and rhizosphere soil were identified. In addition, the core genera in network were obtained by the "WGCNA" package, which plays an important regulated role in network functions. These methods enable researchers to analyze the response of microbial network to environmental disturbance and verify different microbial ecological response theories. The results of these methods show that the significant differential microbial networks identified in the endosphere, rhizoplane and rhizosphere soil of rice.

Network analysis was applied to evaluate the association of various ecological microbial communities, such as soil, water and rhizosphere. Presented here is a protocol on how to use the WGCNA algorithm to analyze different co-occurrence networks that may occur in the microbial communities due to different ecological environments.

The root microbiome plays an important role in plant growth and environmental adaptation. Network analysis is an important tool for studying communities, ...

Compost Microcosms as Microbially Diverse, Natural-like Environments for Microbiome Research in Caenorhabditis elegans

The nematode Caenorhabditis elegans is emerging as a useful model for studying the molecular mechanisms underlying interactions between hosts and their gut microbiomes. While experiments with well-characterized bacteria or defined bacterial communities can facilitate the analysis of molecular mechanisms, studying nematodes in their natural microbial context is essential for exploring the diversity of such mechanisms. At the same time, the isolation of worms from the wild is not always feasible, and, even when possible, sampling from the wild restricts the use of the genetic toolkit otherwise available for C. elegans research. The following protocol describes a method for microbiome studies utilizing compost microcosms for the in-lab&#160;growth in microbially diverse and natural-like environments.
Locally sourced soil can be enriched with produce to diversify the microbial communities in which worms are raised and from which they are harvested, washed, and surface-sterilized for subsequent analyses. Representative experiments demonstrate the ability to modulate the microbial community in a common soil by enriching it with different produce and further demonstrate that worms raised in these distinct environments assemble similar gut microbiomes distinct from their respective environments, supporting the notion of a species-specific core gut microbiome. Overall, compost microcosms provide natural-like in-lab environments for microbiome research as an alternative to synthetic microbial communities or to the isolation of wild nematodes.

Compost Microcosms as Microbially Diverse, Natural-like Environments for Microbiome Research in Caenorhabditis elegans

Compost microcosms bring the microbial diversity found in nature into the laboratory to facilitate microbiome research in Caenorhabditis elegans. Provided here are protocols for setting up microcosm experiments, with the experiments demonstrating the ability to modulate environmental microbial diversity to explore the relationships between environmental microbial diversity and worm gut microbiome composition.

The nematode Caenorhabditis elegans is emerging as a useful model for studying the molecular mechanisms underlying interactions between hosts and their ...

Isolation, Characterization, and Total DNA Extraction to Identify Endophytic Fungi in Mycoheterotrophic Plants

Mycoheterotrophic plants present one of the most extreme forms of mycorrhizal dependency, having totally lost their autotrophic capacity. As essential as any other vital resource, the fungi with which these plants intimately associate are essential for them. Hence, some of the most relevant techniques in studying mycoheterotrophic species are the ones that enable the investigation of associated fungi, especially those inhabiting roots and subterranean organs. In this context, techniques for identifying culture-dependent and culture-independent endophytic fungi are commonly applied. Isolating fungal endophytes provides a means for morphologically identifying them, analyzing their diversity, and maintaining inocula for applications in the symbiotic germination of orchid seeds. However, it is known that there is a large variety of non-culturable fungi inhabiting plant tissues. Thus, culture-independent molecular identification techniques offer a broader cover of species diversity and abundance. This article aims to provide the methodological support necessary for starting two investigation procedures: a culture-dependent and an independent one. Regarding the culture-dependent protocol, the processes of collecting and maintaining plant samples from collection sites to laboratory facilities are detailed, along with isolating filamentous fungi from subterranean and aerial organs of mycoheterotrophic plants, keeping a collection of isolates, morphologically characterizing hyphae by slide culture methodology, and molecular identification of fungi by total DNA extraction. Encompassing culture-independent methodologies, the detailed procedures include collecting plant samples for metagenomic analyses and total DNA extraction from achlorophyllous plant organs using a commercial kit. Finally, continuity protocols (e.g., polymerase chain reaction [PCR], sequencing) are also suggested for analyses, and techniques are presented here.

The current article aims to provide detailed and adequate protocols for the isolation of plant-associated endophytic fungi, long-term preservation of isolates, morphological characterization, and total DNA extraction for subsequent molecular identification and metagenomic analyses.

Mycoheterotrophic plants present one of the most extreme forms of mycorrhizal dependency, having totally lost their autotrophic capacity. As essential as ...

Developing Synergistic Multispecies Biofilms from Rhizosphere Soil

An Approach to Constructing Multispecies Biofilm Communities from Rhizosphere Soil

The multispecies biofilm is a naturally occurring and dominant lifestyle of bacteria in nature, including in rhizosphere soil, although the current understanding of it is limited. Here, we provide an approach to rapidly establish synergistic multispecies biofilm communities. The first step is to extract cells from rhizosphere soil using the differential centrifugation method. Afterward, these soil cells are inoculated into the culture medium to form pellicle biofilm. After 36 h of incubation, the bacterial composition of the biofilm and the solution underneath are determined using the 16S rRNA gene amplicon sequencing method. Meanwhile, high-throughput bacterial isolation from pellicle biofilm is conducted using the limiting dilution method. Then, the top 5 bacterial taxa are selected with the highest abundance in the 16S rRNA gene amplicon sequencing data (pellicle biofilm samples) for further use in constructing multispecies biofilm communities. All combinations of the 5 bacterial taxa were quickly established using a 24-well plate, selected for the strongest biofilm formation ability by the crystal violet staining assay, and quantified by qPCR. Finally, the most robust synthetic bacterial multispecies biofilm communities were obtained through the methods above. This methodology provides informative guidance for conducting research on rhizosphere multispecies biofilm and identifying representative communities for studying the principles governing interactions among these species.

Author Spotlight: Developing Synthetic Microbial Communities for Generating Second-Generation Biofertilizers

A rapid and standardized procedure for establishing synergistic multispecies biofilm communities from various rhizosphere soils is presented here. It is a unique protocol designed to probe and simulate the complex rhizosphere soil microbiota.

The multispecies biofilm is a naturally occurring and dominant lifestyle of bacteria in nature, including in rhizosphere soil, although the current ...

 Microscale SEM Analysis of Microbial Communities in Fungus-Growing Ant Cultures

Microbiota of Attine Ants' Gardens: Visualizing a Microbial Landscape by Scanning Electron Microscopy

In macroscale ecosystems, such as rainforests or coral reefs, the spatial localization of organisms is the basis of our understanding of community ecology. In the microbial world, likewise, microscale ecosystems are far from a random and homogeneous mixture of organisms and habitats. Accessing the spatial distribution of microbes is fundamental for understanding the functioning and ecology of the microbiota, as cohabiting species are more likely to interact and influence each other&#39;s physiology.
An interkingdom microbial ecosystem is at the core of fungus-growing ant colonies, which cultivate basidiomycete fungi as a nutritional resource. Attine ants forage for diverse substrates (mostly plant-based), metabolized by the cultivated fungus while forming a spongy structure, a &#34;microbial garden&#34; that acts as an external gut. The garden is an intertwined mesh of fungal hyphae growing by metabolizing the substrate, opening niches for a characteristic and adapted microbiota to establish. The microbiota is thought to be a contributor to substrate degradation and fungal growth, though its spatial organization is yet to be determined.
Here, we describe how we employ Scanning Electron Microscopy (SEM) to investigate, with unprecedented detail, the microbiota and biofilm spatial organization across different fungiculture systems of fungus-growing ants. SEM imaging has provided a description of the microbiota spatial structure and organization. SEM revealed that microbiota commonly assemble in biofilms, a widespread structure of the microbial landscapes in fungiculture. We present the protocols employed to fix, dehydrate, dry, sputter coating, and image such a complex community. These protocols were optimized to deal with delicate and heterogeneous samples, comprising plant and fungal biomass, as well as the microbiota and the biofilm.

We propose an optimized Scanning Electron Microscopy protocol for visualizing highly heterogeneous and delicate samples containing plant and fungal biomass, together with microbiota and biofilm. This protocol allows describing the spatial dimensions of the microbiota organization.

Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere

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