Extraction and Analysis of Microbial Phospholipid Fatty Acids in Soils

The overall goal of this laboratory protocol is to provide information about the structure of soil microbial communities, through identification of phospholipid fatty acids. This method can help answer key questions in science, such as characterizing soil microbial response to land management changes or to natural disturbance. The main advantage of this technique is that it is an easy-to-follow, robust protocol that can applied to a wide range of soil types.

So, demonstrating this protocol will be Katelyn Pretzlaff, a technician in our research school. To begin the extraction, prepare 5.0 molar potassium hydroxide and 0.15 molar citrate buffer, as described in the text protocol. Prepare the 19:0 nonadeconoate surrogate standard daily by diluting 250 microliters of the stock solution in 25 milliliters of chloroform.

This provides enough surrogate standard for a batch of 23 samples. Add 0.5 milliliters of the 19:0 surrogate standard solution to the sample centrifuge tube. To perform the Bligh and Dyer extraction on the soil sample, add 2.0 milliliters of citrate buffer followed by 2.5 milliliters of chloroform and then 5.0 milliliters of methanol.

Cap the sample with a PTFE-lined cap and vortex for 30 seconds. Then, place it in an end-over-end shaker for two hours. After two hours, centrifuge the sample at 226 times G for 15 minutes with the cap on.

Draw the supernatant off with a Pasteur pipette, and transfer it to a labeled 45-milliliter glass vial. Add a second round of Bligh and Dyer extractant to each sample and repeat the shaking and centrifugation steps. Draw the supernatant off with a Pasteur pipette and transfer it to the same labeled 45-milliliter glass vial.

Next, add 5.0 milliliters of chloroform and 5.0 milliliters of citrate buffer to the labeled 45-milliliter glass vial containing the supernatant. Cap it with a white PTFE-lined cap and vortex the glass vial for 30 seconds. Carefully vacuum off the upper aqueous phase of the 45-milliliter vial.

Pipette the lower organic phase into a labeled 15-milliliter vial. Place a batch of 15-milliliter vials under compressed nitrogen at room temperature to avoid oxidation. Evaporate the chloroform off slowly, setting the nitrogen flow to ruffle the surface of the liquid in the vial but not climb the sides of the vial.

Screw on a PTFE-lined cap and store the samples in the freezer at minus 20 degrees Celsius, wrapped in aluminum foil. Label new solid-phase extraction or SPE columns as necessary. Place an SPE column holder with spigots on the glass tank, and insert the labeled columns into the spigots.

Condition each column by adding 5 milliliters of acetone and letting it drain through. Then, add two additions of 5 milliliters of chloroform. Allow the second chloroform wash to flow out until it's approximately 1 milliliter above the and then close the spigot.

Re-dissolve the sample by adding 0.5 milliliters of chloroform to the vial and vortexing gently. Transfer the re-dissolved sample to the SPE column using a Pastuer pipette. Lay the lipid sample directly onto the center of the column and allow the solvent to drain completely into the tank.

Elute the neutral lipids by adding 5 milliliters of chloroform to each column. Allow the solvent to drain completely into the tank. Elute glycolipids by adding 5 milliliters of acetone to each column, allowing the solvent to drain completely into the collection tank.

Remove the column stand from the tank and drain the tank with a vacuum apparatus. Insert the rack with clean and labeled centrifuge tubes into the tank, and replace the column stand on the tank. The labeled column above should line up with the labeled centrifuge tube below.

Elute phospholipids into centrifuge tubes by adding 5 milliliters of methanol to each column. Wait for the SPE columns to dry out in the fume hood before disposing of them. Dry down the phospholipid fractions at room temperature under compressed nitrogen.

After purging the tubes with nitrogen, screw on PTFE-lined caps and store the samples in the freezer at minus 20 degrees Celsius, wrapped in aluminum foil. Turn on the hot water bath, set to 37 degrees Celsius. Remove the samples from the freezer and allow the samples to come to room temperature.

Add 0.5 milliliters of chloroform and 0.5 milliliters of methanol to each sample, followed by 1.0 milliliters of mathanolic potassium hydroxide. Cap the tubes tightly with PTFE-lined caps and swirl to mix. Place the sealed samples in a 37-degrees Celsius bath for 30 minutes.

Ensure that the water level is a minimum of one to two millimeters above the level of the sample liquid. While the samples are in the water bath, label small glass vials with sample IDs. After 30 minutes, remove the samples and allow them to cool.

Next, add 2.0 milliliters of hexane to each sample and swirl to mix. Then, add 0.2 milliliters of 1.0 molar acetic acid to each sample and swirl again. Phase separation should become visible.

Add 2.0 milliliters of distilled water to each sample to break the phase. After vortexing the samples for 30 seconds, centrifuge the samples at 226 times G for two minutes. Using a short Pasteur pipette, transfer the top phase to clean, labeled, 10-milliliter vials.

Be careful not to transfer any of the lower aqueous phase. Add 2.0 milliliters of hexane to each sample centrifuge tube and swirl. Vortex the samples for 30 seconds and then centrifuge as before.

Using a short Pasteur pipette, again add the top phase to the labeled 10-milliliter glass vials. Evaporate the solvent in the labeled 10-milliliter glass vials at room temperature under nitrogen. Screw on PTFE-lined caps and wrap the samples in aluminum foil.

Store the samples in the freezer at minus 20 degrees Celsius until ready to proceed with gas chromatography analysis as described in the text protocol. A representative chromatogram, obtained from subjecting a sample to capillary gas chromatography using a flame ionization detector is shown here. Retention times in minutes are indicated for the PLFAs in parentheses.

Responses from all PLFAs can be summed to obtain the total PLFA biomass in nanomoles per gram of soil. The relative distribution of different PLFA groups can also be calculated and compared among soils. Results indicate greater total PLFAs for the soil at site one, which is located under older trees, followed by the soil under the intermediate-aged trees at site two.

And lastly, the soil under the youngest trees, at site three. Relatively more saturated PLFAs are present at the youngest site, while more unsaturated PLFAs are present at the oldest site. PLFA results can be further analyzed using multivariate ordination techniques such as a non-metric, multidimensional scaling ordination as illustrated here.

The youngest site separates very clearly from the older two sites, which show overlap in their PLFA microbial community composition. The amount of variation in the PLFA community data explained by each axis is included in parentheses. Once this technique is mastered, it is possible for one person to process 20 samples over four days.

While attempting this protocol, it is important to remember that lipids are particularly susceptible to oxidation. Hence, it is important to protect samples from air and light exposure, such as by storing them in the dark and under nitrogen. After watching this video, you should have a good understanding of how to quantify the structure of soil microbial communities by identifying phospholipid fatty acids through a three-step process, lipid extraction, lipid fractionation, and lipid methylation.