Een methode voor de extractie en analyse van lipiden voor het karakteriseren van bodemmicrobollen in experimenten met veel monsters

The goal of this video is to describe a method that increases throughput while balancing effort and accuracy for the extraction of lipids from cell membranes of microorganisms for the use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure and studies with many samples. In the field, you should account for soil heterogeneity collecting soil samples in a way that will represent your site. To preserve the microbial community at the time of sampling, samples should be transported from the field on ice.

Once back in the lab, remove roots and stones and break up clods by coarse sieving. This also serves to homogenize your sample. Prepare for freeze drying by putting sub samples in appropriate containers and freeze dry as soon as possible.

Once soils are freeze dried, store in a sealed container with desiccant until extraction. It is best to store freezer dried soils at 80 degrees C.As a prep for extraction remove freeze dried soils from storage and grind. Methods for grinding include ball mill, ball meter, or mortar and pestle.

After grinding the soil, again, homogenize your soil samples thoroughly and store in the freezer. All labware must be scrupulously clean. Any residual detergent, grease, or dirt can impact the results by appearing as a peak in the final GC chromatogram.

The extractions are carried out in 30 milliliter Teflon centrifuge tubes, which must be solvent rinsed. Add two to three milliliters of hexane to tubes and vortex for a few seconds. Decant the hexane to another tube and vortex.

Two to three milliliters of hexane can be used to serially rinse six tubes. Store hexane rinsed tubes upside down in the fume hood and dispose of the used hexane in an appropriate waste container. Glassware can be muffled or rinsed with solvent immediately prior to use.

Muffling ensures the glassware is clean by oxidation of residues at high temperature. Wrap the glassware in two to three layers of aluminum foil and place in the muffle furnace. Set to 450 degrees C, and once the furnace has reached the set point bake for a minimum of 4 1/2 hours.

Allow at least one hour for cooling before removing from the furnace. Label and weigh hexane rinsed Teflon tubes. Add soil into Teflon centrifuged tubes and reweigh for soil mass.

Always make at least two blanks per batch and include a check standard, preferably a soil that has been previously extracted, if you want to verify that the extraction worked correctly. Lipid extractions. Prepare three five to 10 milliliter re-pipettes for phosphate buffer, chloroform, and methanol.

Chloroform is a very dense liquid having a low surface tension. Be careful that the amount you dispense is accurate and consistent. Keep the bottle at least half full of liquid.

In the fume hood, add the reagents to the soil in the Teflon tube in the following order, phosphate buffer, chloroform, and methanol. This is the first physical extraction of lipids from your sample material. The recipes for reagent solutions are included in the written protocol.

The solvent proportions and the order of addition are important for proper separation of the organic and aqueous phases. Allow the soils time to wet after the buffer addition before adding the chloroform. If convenient, the extractants can be combined and added as a single aliquot for the second and any subsequent extractions.

Cap the Teflon tubes tightly and cover to protect from light. Place them on the shaker horizontally making sure they are well secured. With a speed setting of high, shake for when hour.

While the tubes are shaking, prepare two long glass tubes for each sample as follows. Label the tube, add the same volume of chloroform that was added to the soil and an equal volume of phosphate buffer. After removing the Teflon tubes from the shaker at 25 degrees C centrifuge the tubes for 10 minutes at 2, 500 RPM.

Then, in the fume hood with the lights turned off decant the supernatant from the Teflon tube into one of the long tubes. Phase separation should be visible in the glass tube. The lower layer contains the organic solvent, primarily chloroform, and the lipids.

This extraction is repeated. Pour the supernatant into the second tube prepared earlier. Now you have physically extracted the lipids from the soil twice.

For most soils, this is adequate. Securely cap all the long class tubes with Teflon lined caps and invert 10 times to mix. Separate phases by gravity overnight.

Allow the samples to stand undisturbed overnight to complete separation of the two phases. To do this, keep the samples in a dark cabinet or covered in aluminum foil at room temperature. It's okay to allow the extracts to separate over the weekend.

Day two, isolation of lipids. On day two, the aqueous layer is aspirated and the lipids concentrated by removing the solvent in a vacuum evaporation system. The samples may also be dried by placing them in a water or sand bath and applying a gentle stream of nitrogen.

The liquid in the tubes should now be well separated and largely clear. If not, or if the interface between the two layers is especially thick, allow the separation to continue for another day. Set up a vacuum aspirator in the hood.

This is a side arm flask connected to a vacuum pump with a leaf Tygon tubing and a pasture pipette. With the pump running, using the pipette to draw off the aqueous phase into the flask. Aspirate the top layer and the interface.

This will be approximately 2/3 of the way down. Do this to the two to three sets of glass tubes. The top layer may contain a few particles of soil.

Remove these if possible. Combine the extract from the second and or third set of tubes with that in the first two by carefully decanting. Try to exclude from pouring any solid material and rinse the walls of the tube by rotating.

Use a clean pipette for each sample and repeat this process for the remaining samples. Once the chloroform extracts have been combined and have been sitting for a few minutes, inspect the surface of the liquid. Often, a thin layer of residual water will form.

If this is present, aspirate before proceeding. Residual water may attack fatty acid double bonds, but a very small amount should co-evaporate with the solvents as the samples are dried. Dry down all the samples using the vacuum evaporation apparatus.

Cap the tubes tightly and store in a freezer at 80 degrees C.Day three, saponification and methylation. First, turn on the water baths. Check the water level and set bath one to 95 degrees C and bath two to 80 degrees C.Using a re-pipette, add one milliliter of the saponification reagent, Reagent 1, to the dried lipids.

Cap tightly, vortex briefly, and place on a rack. Once down with this step, place the rack of tubes into the 95 degree C water bath and wait five minutes. Remove the rack of tubes from the bath and check the tubes for leaks.

This will be indicated by bubbles rising in the tube like a foam. Retighten or replace the caps of the leaking tubes. Continue heating the tubes in the water bath for another 10 minutes.

Reduce the temperature setting on the water bath to 80 degrees C and continue the incubation for another 15 minutes. Remove the tubes and cool by placing the rack into a pan of tap water. Do not use ice water.

Once the samples have cooled, add two milliliter of the methylation reagent, Reagent 2, to each sample. Again, cap tightly and vortex for five to 10 seconds. Granular salts may become visible as a precipitant in the solution, which happens sometimes because of excess reagents.

Place the rack into the 80 degree C water bath and incubate for 10 minutes. Remove the rack of tubes from the water bath and put it into a pan of tap water for cooling. Agitate the rack to accelerate the cooling process.

Using a re-pipette, add one 1.25 milliliters of hexane and methyl tertiary butyl ether, Reagent 3, to each tube to extract the phase. Seal the caps tightly and put the tubes on a shaker for 10 minutes. After shaking, allow the rack of tubes to sit for 10 minutes for the phases to separate.

Transfer the organic phase, which is now the top layer, to a short glass tube using a pasture pipette. It is best to recover most of the liquid. Very small amounts of the aqueous phase will not compromise the extraction.

Repeat aqueous phase extraction by adding Reagent 3, shaking, allowing phases to separate, and transferring the top phase. Depending on the condition of the bottom phase, you can repeat this one more time for a total of three transfers. Add 3 milliliter of the base wash, Reagent 4, which is a dilute solution of sodium hydroxide, to the extracts in the short tubes.

Cap the test tubes tightly and vortex for 20 to 30 seconds and then centrifuge for three minutes at 2, 000 RPM. Once centrifuged, using a clean pasture pipette aspirate the top organic phase and transfer to a four milliliter amber vial. Be very careful not to aspirate any of the aqueous phase.

The next step is to evaporate the solvent to dryness in a vacuum evaporation apparatus. Day four, prepping the extracts for GC analysis. Using the pipetter, add 100 microliters of Reagent 3 to each of the 4 milliliter vials containing the dried phase.

Vortex the sample and then allow it to stand for 10 minutes. Using the second pipetter, carefully transfer the suspended FAMEs to a GC vial. Add another aliquot of the solvent to the four milliliter vial and vortex briefly.

Roll the vial to ensure that the residual FAMEs on the walls of the vial are dissolved. Using the second pipetter again, transfer the solvent to the GC vial. Complete the transfer by adding a third aliquot of solvent to the vial, vortexing, and transferring to the GC vial.

Cap the GC vial and store in the freezer. Store sealed GC vials in freezer prior to analysis. GC analysis.

To use this system, the analysis must be carried out using a specific GC column. The gas chromatograph is equipped with a split splitless inlet fitted with a four millimeter ID glass inlet liner with a deactivated glass wool plug. The inlet is set to 250 degrees C and is operated in a constant pressure mode.

The carrier gas is hydrogen. Nitrogen and air are needed as detector support gasses. An Agilent Ultra 2 column is used.

The column is 25 meters long with a 2 millimeter inside diameter and a 33 micrometer stationary phase film thickness. The stationary phase in this column is 5%phenyl, 95%methylpolysiloxane, also known as a DB-5 type column. To analyze the lipid extracts, a two microliter aliquot is injected at a 100:1 split ratio with the oven temperature at 170 degrees C.Post injection, the oven is programmed to increase at five degrees per minute up to 300 degrees C and then hold for 12 minutes.

A series of injections of the MIDI standard are made and the results are used to make adjustments according to the instructions in the MIDI manual. Once calibrated in this way, the system may need minor adjustments on occasion, but should generally achieve good results using these parameters. The MIDI system produces a report for each sample containing a table with one line per realized peak.

The software reports the peak retention time, peak area, peak identification, along with ECL or estimated chain length, a parameter used for peak identification, and a response factor, a parameter used to normalize variations and detector response with respect to retention time. The ECL expresses where among a series of straight chain FAMEs each unknown FAME elutes. So for example, if the retention time of an unknown elutes halfway between those of a 12 and 13 carbon chain, the ECL is reported as a 12.5 carbon chain.

The software compares the ECL of each peak to those of FAMEs in a database, and where matches occur assigns the corresponding name to the unknown. For cases where two FAMEs in the database have ECLs that are very close, the software reports both names listing the closest first. The data tables from the reports can be collated into a spreadsheet or database.

After adjusting for response factor, peak areas can then be compared with the peak area of an external or internal standard to arrive at a concentration of the extract. By dividing through the mass of soil extracted, the data can be expressed as mass of FAME per gram of soil or, by also using the molecular weight of each FAME, as nanomoles per gram of soil. Biomarker FAMEs can then be summed to yield biomass of microbial guilds, and these guilds can be further analyzed.

For example, here we see unfertilized prairies with more FAME biomass than fertilized prairies. Both have more biomass than nearby corn fields. Also, FAMEs are associated with particular functional groups like fungi or bacteria.

These associations are ecosystem specific, so it is important not to overgeneralize them. This type of analysis shows whether certain groups are more abundant in certain environments. Here, fungi are more abundant in the unfertilized prairie than in the corn field.

And finally, another way to look at the overall microbial community composition is to compare by looking at the relative abundance of all FAMEs at once using ordination methods like nonmetric multidimensional scaling, NMDS, or principal components analysis, PCA. In an ordination, microbial communities that are more similar will be closer together. So, from our example data, corn and unfertilized prairie are very far apart, while some fertilized prairie samples have microbial communities that resemble those from corn and others resemble the unfertilized prairie.

Microbial communities are often highly variable even within an environment, so they won't always separate neatly. After viewing this video, one should have a good understanding of the process by which lipid biomarkers from the cell membranes of microorganisms are extracted from soil. Extraction of phospholipid fatty acids is an effective, rapid, and inexpensive way to assess microbial guilds and total microbial biomass through the identification of unique microbial biomarkers.