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

The overall goal of this experiment is to characterize three compartments of the root microbiome, soil, rhizosphere, and root endosphere, using community profiling of the 16S ribosomal RNA gene. This method can help answer key questions in microbial ecology, such as the biotic and abiotic factors that are the significant drivers of microbiome structure. The main advantage of this technique is that it standardizes each step, from the separation of root and rhizosphere to DNA's extraction to preparing a sequencing library.

Demonstrating this procedure will be Tuesday Simmons, a graduate student in the Coleman-Derr Laboratory. To begin the protocol, collect bulk soil samples with an ethanol-sterilized soil core collector to obtain soil that is free of plant roots. Collect a core approximately 23 to 30 centimeters from the base of the plant.

Next, transfer the soil to a plastic bag, homogenize the soil by gentle shaking the bag, transfer a 600-milligram aliquot of the soil sample into one two-milliliter tube, and immediately place the two-milliliter tube on dry ice until DNA extraction. 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. Gently shake off excess soil from the roots until there is approximately two millimeters of soil adhering to the root surface.

For large plants, use sterile scissors to cut a representative subsection of roots, and place a minimum of 500 milligrams of root tissue into a 50-milliliter conical vial. Add enough epiphyte removal buffer to cover the roots, then immediately place the sample on dry ice. To separate the rhizosphere from the roots, thaw the root sample on ice, then sonicate the root samples.

Transfer the roots into a chilled, clean 50-milliliter tube using sterile forceps. Do not dispose of the original tube with buffer and soil. This contains the rhizosphere fraction.

Next, centrifuge the tube containing the buffer and rhizosphere. Decant the supernatant, and place the rhizosphere fraction in dry ice. To wash the roots, add approximately 20 milliliters of four degrees Celsius sterile water to the root fraction.

Wash the root by shaking vigorously for 15 to 30 seconds, then drain the water. Use a sterile spatula to quickly transfer 250 milligrams of soil and rhizosphere into separate collection tubes provided in a commercial DNA isolation kit for soil extraction, then proceed using the kit supplier's protocol. After elution, store the DNA at minus 20 degrees Celsius until ready to proceed to the amplicon library preparation, then extract DNA from the root samples.

Chill a sterilized mortar and pestle using liquid nitrogen. Measure out 600 to 700 milligrams of root tissue, and place the tissue into the mortar and carefully add enough liquid nitrogen to cover the roots. Grind the roots into small pieces.

Continue the process of adding liquid nitrogen and grinding at least two times, being consistent between samples, until the roots are a fine powder. Quickly, before the root powder begins to thaw, use a sterile spatula to transfer the root powder into pre-weighed 1.5-milliliter tubes on ice. Record the weight of the tube and powder.

Use a sterile spatula to quickly transfer 150 milligrams of root powder to the collection tube provided in the commercial DNA isolation kit designed for extraction from soil, then proceed with DNA isolation using the kit supplier's protocol. Prepare sufficient PCR master mix to amplify each DNA sample in triplicate. Pour the master mix into a sterile, 25-milliliter multichannel pipette reservoir, and distribute 66 microliters of master mix into each well of a new 96-well PCR plate.

Next, add six microliters of five nanograms per microliter DNA from the normalized DNA plate to the master mix plate. Then, add to the master plate 1.5 microliters of 10 micromolar forward primer, such that each column has a different forward barcode, and 1.5 microliters of 10 micromolar reverse primer, such that each row has a different reverse barcode. Cover the plate with film, and spin down the plate briefly at 3, 000 RCF.

Use a multichannel pipette to gently mix the contents, then divide them into three plates with 25 microliters of reaction mixture, and cover the plates with PCR film. Amplify the DNA in each plate using a thermocycler. After the amplification, pool the three replicate plates into a single 96-well plate.

Using a computer spreadsheet, calculate the volume of 100 nanograms of each sample, as well as the average volume for all samples. For each blank PCR product, add the calculated average volume into a single 1.5-milliliter tube. For the successfully amplified samples, pool 100 nanograms of each sample into the same tube, then measure the concentration of the pooled product using a benchtop fluorometer, and elute 600 nanograms of DNA in molecular-grade water to a final volume of 100 microliters in a 1.5-milliliter tube.

Store the remaining pooled product at minus 20 degrees Celsius. Prior to purifying the 600-nanogram DNA aliquot, prepare 600 microliters of fresh 70%ethanol. Follow the established PCR purification process using paramagnetic beads.

Shake the tube of magnetic beads to resuspend the beads that settle to the bottom. Add 1x volume, 100 microliters, of bead solution to the 600-nanogram aliquot of DNA. Mix the solution and beads thoroughly by pipetting 10 times.

After thorough mixing, incubate the mixture for five minutes at room temperature. Place the tube onto the magnetic stand for two minutes or until the solution is clear to separate the beads from the solution. Keep the tube in the stand, aspirate the clear supernatant carefully without touching the magnetic beads, and discard it.

Leave the tube in the stand, add 300 microliters of 70%ethanol to the tube, and incubate the beads at room temperature for 30 seconds. Aspirate out the ethanol, and discard it. Repeat this process, and remove all ethanol after the second wash.

Remove the tube from the magnetic stand, and air-dry the contents for five minutes. Add 30 microliters of molecular-grade water to the dried beads, and mix by pipetting 10 times. Incubate at room temperature for two minutes.

Return the tube to the magnetic stand for one minute to separate the beads from solution. Transfer the eluate to a new tube. Measure the final concentration of the cleaned, pooled DNA using a benchtop fluorometer.

Dilute an aliquot to 10 nanomolar in a final volume of 30 microliters or to the concentration and the volume preferred by the sequencing facility. Following the amplification step, sequencing was performed to determine the bacterial community composition of each sample. An alignment of the PNA sequence to each chloroplast and mitochondrial 16S ribosomal RNA gene for the plant host investigated should not reveal any mismatches.

A single mismatch to the 13 base pair PNA sequence can drastically reduce the effectiveness, as in the case of the provided chloroplast PNA sequence and the chloroplast 16S ribosomal RNA gene of Lactuca sativa, or lettuce. Since an equal amount of amplified DNA was pooled per sample, an even number of reads, which match to bacterial taxa, were obtained per sample after sequencing, sorted based on their barcoded index. Once mastered, this technique can be done for eight plants in approximately eight hours.

While attempting this procedure, it is important to be consistent between experiments. Small details such as changes in DNA extraction method and PCR master mix can introduce bias. Following this procedure, other methods such as metatranscriptomics can be used to answer additional questions, such as which microbes are active, and what genes are they expressing?

Don't forget that working with liquid nitrogen can be extremely hazardous and precautions such as wearing appropriate PPE should always be taken while performing this procedure.