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

Summary

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.

Abstract

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.

Introduction

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' 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 gene^2,3. 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 developed^5,6.

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 kits^8,9 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 studies^10,11. 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 communities^12,13,14. 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 microbiome^16,17 and strawberry microbiome¹⁸.

Protocol

1. Collection and Separation of Root Endosphere, Rhizosphere, and Soil Samples

1. 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 µm pore size.
   1. 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.
2. 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.
3. 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).
   1. 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.
4. 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.
5. 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 °C.
6. To separate the rhizosphere from the roots, thaw the root sample on ice, then sonicate the root samples at 4 °C for 10 min with pulses of 160 W for 30 s, separated by 30 s. Transfer the roots into a chilled (4 °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).
7. Centrifuge the tube containing buffer and rhizosphere for 10 min at 4 °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 °C until ready to proceed with DNA extraction (step 2).
8. To wash the roots, add approximately 20 mL of chilled (4 °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.
9. 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 °C until ready to proceed with DNA extraction.

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.

1. Extract DNA from the soil and rhizosphere samples.
   1. 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's protocol.
   2. After eluting the DNA in the elution buffer supplied by the DNA isolation kit, store the DNA at -20 °C until ready to proceed with step 3.
2. Extract DNA from the root samples.
   1. 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.
   2. 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 °C.
   3. 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.
   4. 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'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.
3. Measure the concentration of all DNA samples using a high-sensitivity benchtop fluorometer.
   1. Add 1 - 20 µL 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 µL.
   2. Prepare two additional tubes containing 10 µL of DNA standard 1 (0 ng/µL DNA) or 10 µL of standard 2 (100 ng/µL), and add 190 µL of fluorometer working solution to each standard.
   3. 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.

3. Amplicon Library Preparation and Submission

1. Set up materials for the amplification reaction.
   1. Thaw DNA samples at 4 °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.)
   2. In a 96-well PCR plate, dilute DNA from each sample in molecular-grade water to 5 ng/µL in a total volume of 20 µL. Add 20 µL of molecular-grade water to the four corner wells as negative controls for amplification (blanks) (Table 2).
   3. Arrange the barcoded primers (10 µM) in either PCR strip tubes or a 96-well plate such that they can be added with a multi-channel pipette (Figure 2).
   4. Prepare sufficient PCR master mix to amplify each DNA sample in triplicate. Prepare 1.5 µL of BSA (20 mg/mL), 37.5 µL of pre-made 2x master mix (composed of PCR buffer, MgCl₂, dNTPs, and Taq DNA polymerase), 0.57 µL of chloroplast PNA (100 µM), 0.57 µL of mitochondrial PNA (100 µM), and 25.86 µL of molecular grade water.
   5. Pour the master mix into a sterile 25 mL of multichannel pipette reservoir and distribute 66 µL 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.
   6. Using a multi-channel pipette, add 6 µL of 5 ng/µL DNA (from the normalized DNA plate) to the master mix plate. Then add to the master plate 1.5 µL of 10 µM forward primer such that each column has a different forward barcode, and 1.5 µL of 10 µM 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.
   7. 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 µL of reaction mixture.
      NOTE: Although three replicates are not strictly necessary, it decreases the impact of technical variability.
2. Amplify the DNA in each plate using a thermocycler set to the following conditions: 180 s at 98 °C, 30 cycles of: 98 °C for 45 s (denaturing), 78 °C for 10 s (PNA annealing), 55 °C for 60 s (primer annealing), and 72 °C for 90 s (extension), then 600 s at 72 °C followed by a 4 °C hold step. After the amplification, pool the three replicate plates into one single 96-well plate.
3. Quantify the DNA using high-sensitivity fluorometer reagents in a 96-well plate reader.
   1. Add 2 µL of each PCR product to a 96-well microplate, along with 98 µL of fluorometer working solution (1:200 dye:buffer). Include 4 wells as standards: 5 µL of DNA standard 1 (0 ng/µL DNA), 1 µL of standard 2 (10 ng/µL), 2 µL of standard 2 (20 ng/µL), and 5 µL of standard 2 (50 ng/µL). Then add fluorometer working solution for a final volume of 100 µL.
      NOTE: Each sample can be measured using a benchtop fluorometer as described in step 2.3 if a plate reader is not available.
   2. Calculate the DNA concentration from the absorbance output by a linear regression of the four standards.
   3. For the successfully amplified barcoded products (those that have a concentration greater than 15 ng/µL), pool 100 ng of each sample into a single 1.5 mL tube (Table 2).
   4. Calculate the average volume of samples added to the pool by using the =AVERAGE() function in a spreadsheet program. Add the volume of the "blank" PCR products to the pooled samples.
      NOTE: Since the "blank" PCR products have their own unique barcode combinations, they can be sequenced to check for any laboratory contaminants.
4. 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 µL in a 1.5 mL tube. Store the remaining pooled product at -20 °C.
5. 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.
   1. Make a fresh 600 µL aliquot of 70% ethanol. Shake the bottle of magnetic beads to re-suspend beads that settle to the bottom.
   2. Add 1x volume (100 µL) of bead solution to the 600 ng aliquot of DNA. Mix thoroughly by pipetting 10 times. Incubate for 5 min at room temperature.
   3. 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.
   4. Leave the tube in the magnetic stand and add 300 µL 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.
   5. Add 30 µL 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.
6. 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 µL, or to the concentration and volume preferred by the sequencing facility.
7. Utilize the services of a sequencing facility to sequence the DNA on a NGS platform, 2 x 300 bp paired-end sequencing.

Results

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...

Discussion

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 millimeters^28,<...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.1-10/20 µL filtered micropipette tips	USA Scientific	1120-3810	Can substitute with equivalent from other suppliers.
1.5 mL microcentrifuge tubes	USA Scientific	1615-5510	Can substitute with equivalent from other suppliers.
10 µL multi-channel pipette	Eppendorf	3122000027	Can substitute with equivalent from other suppliers.
10 µL, 100 µL, and 1000 µL micropipettes	Eppendorf	3120000909	Can substitute with equivalent from other suppliers.
100 µL multi-channel pipette	Eppendorf	3122000043	Can substitute with equivalent from other suppliers.
1000 µL filtered micropipette tips	USA Scientific	1122-1830	Can substitute with equivalent from other suppliers.
2 mL microcentrifuge tubes	USA Scientific	1620-2700	Can substitute with equivalent from other suppliers.
2 mm soil sieve	Forestry Suppliers	60141009	Can substitute with equivalent from other suppliers.
200 µL filtered micropipette tips	USA Scientific	1120-8810	Can substitute with equivalent from other suppliers.
25 mL reservoirs	VWR International LLC	89094-664	Can substitute with equivalent from other suppliers.
50 mL conical vials	Thermo Fisher Scientific	352098	Can substitute with equivalent from other suppliers.
500 mL vacuum filters (0.2 µm pore size)	VWR International LLC	156-4020
96-well microplates	USA Scientific	655900
96-well PCR plates	BioRad	HSP9631
Agencourt AMPure XP beads	Thermo Fisher Scientific	NC9933872	Instructions for use: https://www.beckmancoulter.com/wsrportal/ajax/downloadDocument/B37419AA.pdf?autonomyId=TP_DOC_150180 &documentName=B37419AA.pdf
Aluminum foil	Boardwalk	7124	Can substitute with equivalent from other suppliers.
Analytical scale with 0.001 g resolution	Ohaus Pioneer	PA323	Can substitute with equivalent from other suppliers.
Bioruptor Plus ultrasonicator	Diagenode	B01020001
Bovine Serum Albumin (BSA) 20 mg/mL	New England Biolabs	B9000S
Centrifuge	Eppendorf	5811000908	Including 50mL and 96-well plate bucket adapters
Cryogenic gloves	Millipore Sigma	Z183490	Can substitute with equivalent from other suppliers.
DNeasy PowerClean kit (optional)	Qiagen Inc.	12877-50	Previously MoBio
DNeasy PowerSoil kit	Qiagen Inc.	12888-100	Previously MoBio
Dry ice	Any	NA
DynaMag-2 magnet	Thermo Fisher Scientific	12321D	Do not substitute
Ethanol	VWR International LLC	89125-188	Can substitute with equivalent from other suppliers.
Gallon size freezer bags	Ziploc	NA	Can substitute with equivalent from other suppliers.
Gemini EM Microplate Reader	Molecular Devices	EM	Can use another fluorometer that reads 96-well plates from the top.
K2HPO4	Sigma-Aldrich	P3786
KH2PO4	Sigma-Aldrich	P5655
Lab coat	Workrite	J1367	Can substitute with equivalent from other suppliers.
Liquid N2	Any	NA	Can substitute with equivalent from other suppliers.
Liquid N2 dewar	Thermo Fisher Scientific	4150-9000	Can substitute with equivalent from other suppliers.
Milli-Q ultrapure water purification system	Millipore Sigma	SYNS0R0WW
Mini-centrifuge	Eppendorf	5404000014
Molecular grade water	Thermo Fisher Scientific	4387937	Can substitute with equivalent from other suppliers.
Mortars	VWR International LLC	89038-150	Can substitute with equivalent from other suppliers.
Nitrile gloves	Thermo Fisher Scientific	19167032B	Can substitute with equivalent from other suppliers.
Paper towels	VWR International LLC	BWK6212	Can substitute with equivalent from other suppliers.
PCR plate sealing film	Thermo Fisher Scientific	NC9684493
PCR strip tubes	USA Scientific	1402-2700
Pestles	VWR International LLC	89038-166	Can substitute with equivalent from other suppliers.
Plastic spatulas	LevGo, Inc.	17211
Platinum Hot Start PCR Master Mix (2x)	Thermo Fisher Scientific	13000014
PNAs - chloroplast and mitochondrial	PNA Bio	NA	Make sure to verify sequence bioinformatically
Protective eyewear	Millipore Sigma	Z759015	Can substitute with equivalent from other suppliers.
Qubit 3.0 Fluorometer	Thermo Fisher Scientific	Q33216
Qubit dsDNA HS assay kit	Thermo Fisher Scientific	Q32854
Rubber mallet (optional)	Ace Hardware	2258622	Can substitute with equivalent from other suppliers.
Shears or scissors	VWR International LLC	89259-936	Can substitute with equivalent from other suppliers.
Shovel	Home Depot	2597400	Can substitute with equivalent from other suppliers.
Soil core collector (small diameter: <1 inch)	Ben Meadows	221700	Can substitute with equivalent from other suppliers.
Spray bottles	Santa Cruz Biotechnology	sc-395278	Can substitute with equivalent from other suppliers.
Standard desalted barcoded primers (10 µM) (Table 1)	IDT	NA	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.
Thermocycler	Thermo Fisher Scientific	E950040015	Can substitute with equivalent from other suppliers.
Triton X-100	Sigma-Aldrich	X100	Can substitute with equivalent from other suppliers.
Weigh boats	Spectrum Chemicals	B6001W	Can substitute with equivalent from other suppliers.