Isolatie en analyse van microbiële gemeenschappen in bodem, rhizosfeer en wortels in de overblijvende gras experimenten

This method can help answer key questions in the soil, rhizosphere, and root endosphere microbiome field, such as how microbial community diversity changes in response to sample type, treatment, and plant genotype. The main advantage os this technique is that it is well-suited for field experiments that require large sample numbers and replication. Demonstrating the procedure will be myself and Stephanie Futrell, a grad student and research technician from my laboratory.

To begin the protocol, label a wash pan and bucket with a sticky note with plant sample details. Carry the labeled bucket to the plot and leave the wash pan at the established workstation in the field. Randomly choose and collect two plants per plot from different areas within the plot.

Next, pierce the soil with a shovel to a depth of 30 centimeters to cut any of the lateral roots holding the plant in the soil. Excavate the plant roots by leveraging the shovel and place the root ball in the labeled bucket. Bring the bucket back to the workstation in the field.

Shake the roots and use a spade or handheld tiller to remove soil from the roots. Wear gloves and place the roots near the processing station. After shaking the roots, mix the soil in the wash pan and break up any soil clods with a handheld tiller.

Place a sample of soil that is free of debris into a labeled, 17.7 by 19.5-centimeter zipper storage bag and place it on ice. Sterilize pruning scissors in 70%ethanol. Use the sterile scissors to excise a variety of roots, approximately four to six roots per plant and each root about nine to 12 centimeters in length.

Place the excised roots in a labeled 50-milliliter tube containing 35 milliliters of autoclaved phosphate buffer with a surfactant such as Silwet L-77. Shake the tubes for two minutes to release the rhizosphere from the surface of the roots. Improve shaking with a generator and vortexer.

With forceps sterilized in 70%ethanol, remove the roots from the tube, blot them briefly on paper towels, and place them in a new, labeled, 50-milliliter tube. Place both the tube containing the rhizosphere and the one containing roots on ice. Add approximately 35 milliliters of 50%bleach plus 0.01%Tween 20 to the 50-milliliter tubes with roots collected in the field.

Vortex or shake the tubes for 30 to 60 seconds. Pour off the bleach, add 35 milliliters of 70%ethanol, and vortex or shake the roots for another 30 to 60 seconds. After shaking, pour off the 70%ethanol and add 35 milliliters of sterile, ultrapure water and vortex or shake the sample for one minute.

Repeat the water-rinse step two more times so the root receives a total of three rinses with sterile, ultrapure water. Blot the roots on dry, clean paper towels and use a clean paper towel for each sample. Using sterile forceps and pruning scissors, cut the roots into approximately five-millimeter pieces and place cut roots in a clean, labeled, 15-milliliter conical tube, then store the samples at minus 80 degrees Celsius until further processed.

Shake the 50-milliliter tubes containing rhizosphere samples from the field for 20 to 30 seconds to re-suspend the entire sample. Using a sterile, 100-micron mesh cell strainer, filter the re-suspended sample into a new 50-milliliter tube. Then, centrifuge the tubes at 3, 000 times g for five minutes at room temperature.

Immediately pour off and discard the supernatant. Place the rhizosphere pellets in the 50-milliliter tubes on ice. Next, add 1.5 milliliters of sterile phosphate buffer without surfactant to the rhizosphere pellets and vortex them to re-suspend the sample.

Pipette the suspended liquid into a clean, labeled, two-milliliter microfuge tube. Spin the tubes at 15, 871 times g for two minutes at room temperature. Immediately pour off the supernatant and drain the tubes on clean paper towels.

Store the pellets at minus 20 degrees Celsius until further processed. Using a sterile metal spatula, fill a clean, labeled, two-milliliter tube with approximately three grams of soil for DNA extraction and avoid any small root pieces and debris. Rinse the metal spatula in 70%ethanol between each sample.

Store this soil sample at minus 20 degrees Celsius. In a clean wash pan, empty the bag of soil into stacked sieves with the larger sieve on top of the smaller sieve and manually sieve the soil through both sieves. Use a brush to carefully clean the sieves between samples.

Set aside 100 to 125 grams of sieved soil in a 17.7 by 19.5-centimeter zippered bag for future soil physicochemical and texture analysis. Place the bags of soil at four degrees Celsius for short-term storage. Next, pour liquid nitrogen into a plastic beaker with clean spatulas and in a clean mortar.

Place the frozen tissue in the mortar and grind it with the pestle into a fine powder. Continually add liquid nitrogen throughout grinding to keep samples frozen. Use a spatula to transfer the ground tissue in a clean, labeled, two-milliliter tube and then store the tissue at minus 80 degrees Celsius.

Wipe down the work area with 70%ethanol and 1%household bleach. Remove the soil samples from the minus-20-degrees-Celsius storage and thaw them in an ice bucket. Remove the sealing mat cover from a 96-well extraction plate and place the cover between two paper wipes to keep it clean while not in use.

Cover the 12 columns of the plate with adhesive 8-well PCR strips to avoid contamination. Tare a sterile, size small weigh-funnel on a scale and weigh out 200 to 250 milligrams of soil. Carefully lift the adhesive strip up, place the neck of the filled weigh-funnel into the appropriate well, and gently guide the soil sample into the appropriate well.

Replace the adhesive strip to cover the well. Repeat this process for every well, using a new sterile funnel for each sample. Replace the sealing mat cover on the extraction plate and store the plate at minus 20-degrees Celsius until ready for DNA extraction.

Remove the rhizosphere samples from the minus 20-degrees-Celsius storage and thaw in an ice bucket. Prepare the DNA extraction plate as done in the previous section. Place a clean paper wipe on a scale, then tare a sterile metal spatula on the scale.

Use the spatula to carefully scoop out some of the rhizosphere pellet from a sample tube. Weigh between 200 and 250 milligrams of the rhizosphere sample. Carefully lift the adhesive strip to uncover the first well of the extraction plate.

Angle the filled spatula into the well and scrape off the rhizosphere material into the appropriate well with a sterile toothpick. Rinse the metal spatula in water followed by 70%ethanol between samples. Finally, repeat this process for every well of the plate until the plate is filled, leaving one well empty as an extraction blank control.

In this representative data set, there were highly significant differences between sample types, and the rhizosphere and soil appear to be more similar in composition to each other than the root. Further analysis shows that there were also highly significant differences in the microbial community composition between rhizosphere and soil. Alpha diversity analysis shows that the microbial communities in the endosphere were lower than communities in the soil and rhizosphere.

The only significant difference in diversity between the grass species was between the endosphere samples of big bluestem and switchgrass. The relative abundance analysis highlights the dominance of Proteobacteria followed by Actinobacteria in all sample types. Soil and rhizosphere are dominated by Acidobacteria and Chloroflexi, whereas the roots had a larger relative abundance of Bacteriodetes.

PERMANOVA analysis of all sample types elucidated a highly significant difference in microbial community composition due to plant species. After watching this video, you should have a good understanding of how to perform a series of steps to use in the study of soil, endosphere, and rhizosphere microbiomes. We would like to thank the Nebraska EPSCoR and the National Science Foundation EPSCoR Research Infrastructure Program Track 1 for partial funding of this educational video.