PLFA extraction protocols

Transcription

1 2015 Fyffe Ct, Columbus, OH Parker Food Science & Technology Building, Room 070 PLFA extraction protocols page 1. Laboratory safety and general guidelines 1 2. PLFA-Extraction (Frostegård et al. 1993, Bardgett et al. 1996) 2 3. References 7

2 1. Laboratory safety and general guidelines, FAME extract and soil sample storage Phospholipid fatty acids are extracted and fractionated by using solvents that are potentially hazardous for working personnel in the laboratory. Please study Material Safety Data Sheets (MSDS) carefully prior analysis. Avoid spills, inhalation and skin contact with solvents. Collect all waste in labeled containers (please use yellow hazardous waste stickes) under the fume hood. Solvent waste needs to be picked up for recycling or disposal, please contact the OSU Hazardous waste pickup service at Please make sure that you are wearing appropriate protection (safety glasses, nitrile or neoprene gloves and lab coat) before handling solvents. Open solvent bottles only under the fume hood. Before removing solvents from the fume hood, make sure that bottles or vials are capped tightly. All glassware used for the extraction needs to be thoroughly washed (4% soap bath (2-4hrs.) and 10% acid bath (overnight)), baked (250ºC for 6hrs.). If you have trouble with contamination (FAME peaks are showing up in the blanks), please rinse glassware prior use with hexane (EL-FAME method) or chloroform (PLFA). The solvent rinse should be done under the fume hood 1 day prior analysis, please rinse glass tubes twice with 2 ml solvent, vortex tubes thoroughly, discard solvent in the labelled PLFA waste bottle. Place the tubes upside down in the racks to dry. Also rinse glass pipettes when required. Phospholipid fatty acids are degradable due to exposure to light, heat, water and oxygen. Avoid sample contact with any of these conditions. Use only glass vials/tubes for storing FAME extracts. We are currently running a test measuring FAMEs after dried extract storage (last step of the protocol before dissolving the sample for measurement) at -20 C for several weeks. We will include the results of the storage test here latest in March In the meantime, storage of extracted PLFAs or methylized FAMEs should be done dissolved in solvent at -20 C and not exceed two weeks (MIS operating Manual 2002, page 2-17). Storage of soil samples should be avoided when analyzing PLFAs. It is better to store the first PLFA extract (PLFA in solvent) rather than to store the soil sample. If however required, store soil samples at -20 C prior analysis (Lee et al. 2007). 2

3 2. PLFA Extraction from soils (Frostegård et al. 1993, Bardgett et al. 1996) RATIONALE Lipids are extracted from soil with a single-phase mixture of chloroform, methanol and aqueous citrate buffer (Bligh and Dyer reagent). The organic, lipid-containing, phase is collected and the lipids are separated into neutral, glyco- and phospholipid fractions using silicic acid columns. The phospholipids are then converted to their methyl-esters by alkaline methanolysis and detected by GC-FID. REAGENTS AND EQUIPMENT (all organic solvents should be HPLC grade) M citrate buffer, ph 4: g citric acid dissolved in 1 l water and adjusted with NaOH pellets to ph 4. - Extraction solution (Bligh and Dyer reagent): Chloroform, methanol and citrate buffer mixed in the proportions 1:2:0,8 (v/v/v). - Acetone - Hexane-chloroform solution (4:1, v/v) - Methanol-Toluene solution (1:1, v/v) M methanolic KOH: 5.61 g KOH pellets dissolved in 500ml methanol. (prepare fresh) - 1 M acetic acid: 5.72 ml glacial (100%) acetic acid make up to 100 ml with deionized H2O. - Internal Standard: Dissolve 0.01 M Methylnonadecanoate [Nonadecanoic acid methyl ester, CH3(CH2)17COOCH3, MW (order through Sigma, # G)] in Hexane/ MTBE (1:1). Prepare the standard in a 25 ml volumetric flask, add g Methylnonadecanoate, fill up to 25 ml with Hexane/MTBE (1:1). Keep in small (5 ml) teflon-lined screw cap vials at -20 C - 35 ml and 10 ml centrifuge tubes with teflon-lined screw caps, 20 ml test tubes - Water bath for methanolysis - Sample concentration apparatus (Techne dri-block) - Apparatus for lipid fractionation: Visiprep Vacuum manifold with common solvent trap and 10ml collection vials (Visiprep Solid Phase Extraction Vacuum manifold, #57030-U, Supelco). - Silicic acid columns for the solid phase separation: 500 mg, 3 ml capacity; Cat.no.: from Supelco (order through Sigma) - Vortex for mixing samples, solvent resistant pump - Centrifuge PREPARATION IN ADVANCE 3

4 Clean all glassware before use as described in chapter 1 and rinse twice with Chloroform before analysis. Use fresh soil samples, if soil used is frozen, it needs to be homogenized. LIPID EXTRACTION Weigh out 0.5 g humus-rich soil or 1.5 to 2 g mineral soil into a 35 ml glass centrifuge tube. Include one or two 35 ml glass centrifuge tubes as blanks (no soil). Note: Recently we had trouble to extract fatty acids from very sandy soils. Make sure to test the extraction efficiency from sandy soils first before you run several batches of samples. Try to increase the amount of soil to 15 g, while extracting 5 times 3 g of soil. Follow the protocol until the end and run the sample on the GC to check the amounts of PLFAs you get. After weighing the soil in the vials, add up to 1.5 ml citrate buffer (reduced to take into account the amount of water contained within each soil sample, you want a total aqueous phase of 1.5 ml at this stage), 1.9 ml chloroform, 3.8 ml methanol and 2 ml Bligh and Dyer reagent to the samples and mix with a vortex (or: in total you might want to add 9.2 ml Bligh and Dyer reagent to dry soil). Please use the black teflon-lined caps to close the vials. Leave the samples for 2 hours in the dark, but vortex every 20 minutes (keep light on for the minimum amount of time light degrades PLFAs). At the end of extraction vortex once more and then centrifuge (Sorvall RC-2B, HS-4 rotor) at 2000 rpm for 15 min. The centrifuge can hold 12 samples in the HS-4 rotor. If you have more samples, leave remaining samples in the fridge in the meantime. Before placing the tubes into the centrifuge, make sure that the tube holders are clean, any debris can cause stress and the glass tubes are likely to break. Please also use cotton wool to line the tube holders to prevent breaking of the glass centrifuge tubes. Place the tubes back under the fume hood and use a Pasteur pipette to transfer the supernatant into a new 35 ml glass centrifuge tube. Re-extract the soil pellet with a 2.5 ml Bligh and Dyer reagent. Remove the supernatant as done before and combine supernatants for each sample. Do not forget to treat the blanks in the same way. PHASE SEPARATION The organic and aqueous phases are separated by adding 3.1 ml chloroform and 3.1 ml citrate buffer to the combined supernatants. The samples are then vortexed and centrifuged at 2000 rpm for 10 min. Using a Gilson-type (Eppendorf) pipette, remove the LOWER (organic) phase and record the volume collected (typically 2 ml, but you can use up to 6 ml). The collected organic phase is transferred to a 10ml glass test tube and dried down under nitrogen. Place the samples in the heating block. Turn on the heater in the back of the block, leave the temperature on the value that is already set (35ºC). The evaporation takes about 20 mins. Make sure that the nitrogen flow is not too high, adjust it carefully until you can see ripples on the surface of the samples. FAMEs are easy degradable under oxygen, make sure 4

5 that the nitrogen flow stays on until the end of the evaporation. After evaporation, turn the nitrogen flow off. The samples can also be stored prior the dry down step (in chloroform) at -20 C until the next step. LIPID FRACTIONATION (Solid phase separation): Activate the columns by adding 2x1 ml chloroform. Reconstitute the dried sample in 1 ml chloroform and apply to the columns with a pasteur pipette. Rinse the sample vial twice with 500 µl chloroform and also apply these washes to the column. Elute the neutral lipids first with 3x2 ml chloroform, then the glycolipids with 3x2 ml acetone and finally the phospholipids with 3x2 ml methanol. The first two eluates (neutral and glycolipids) are discarded. The methanol (phospholipid) eluate is collected in separate 10 ml glass centrifuge tubes and dried down under nitrogen. Place the samples in the heating block. Turn on the heater in the back of the block, leave the temperature on the value that is already set (35ºC). The evaporation takes about 30 mins. Make sure that the nitrogen flow is not too high, adjust it carefully until you can see ripples on the surface of the samples. FAMEs are easy degradable under oxygen, make sure that the nitrogen flow stays on until the end of the evaporation. After evaporation, turn the nitrogen flow off. ALKALINE METHANOLYSIS Add 1 ml methanol-toluene solution (1:1) to each sample. Then add 1 ml 0.2 M methanolic KOH and incubate sample for 15 min. at 37 C in a water bath. After Methanolysis, transfer the samples into 15 ml glass centrifuge tubes using a Pasteur pipette. Rinse each test tube with 2 ml hexane-chloroform (4:1) solution and add this to the corresponding centrifuge tube. Add 0.3 ml 1M acetic acid and 2 ml ultrapure water to your sample. Mix by vortexing, then centrifuge at 2000 rpm for 10 min. Use a pasteur pipette to transfer the upper (organic) phase into a new 10 ml glass test tube. The residual aqueous phase is re-extracted with 2 ml hexane-chloroform solution (4:1), vortexed, centrifuged, and, as before, the upper organic phase is removed and combined in the appropriate glass test tube. Dry down the samples under nitrogen. Therefore, Place the samples in the heating block. Turn on the heater in the pback of the block, leave the temperature on the value that is already set (35ºC). The evaporation takes about 20 mins. Make sure that the nitrogen flow is not too high, adjust it carefully until you can see ripples on the surface of the samples. FAMEs are easy degradable under oxygen, make sure that the nitrogen flow stays on until the end of the evaporation. After evaporation, turn the nitrogen flow off. Dissolve the sample by adding 170 µl Hexante/MTBE (1:1) and 30 µl internal standard (in Hexane/MTBE). Final volume: 200 µl. Transfer the sample as quickly as possible in GC vials (with insert) and cap the vials. 5

6 MEASUREMENT GC system OSU: Midi measurement takes ~30min. per sample Capillar column: HP Ultra 2, length= 25 m Carrier gas: Hydrogen, Nitrogen ( column makeup gas ) Oven Temp. 170 C initial increase: 5 C per min until 270 C Temperature of the injector: 280 C Split: 10.0 Detector: FID (flame ionization detector); T= 300 C Detector gases: Synthetic air Hydrogen 6

7 CALCULATION OF THE RESULTS (ZELLES 1996) FFM: Area of the phospholipid methylester MGFM: Molecular weight of the phospholipid methylester in µg/µmol cis: Concentration of the internal standard in µg FIS: EW: TS: Area of the internal standard Soil in g Dry matter of the soil in % (100/TS= dry matter factor) 2: Variable Factor! Use factor 2 if you take 3ml from 6ml organic Phase in the step of the phase separation after the extraction of PLFAs, use factor 3 if you take 2 ml. 1000: Factor to obtain nmol c: Concentration of the FAME in nmol/g soil 7

8 3. REFERENCES Bardgett R D, Hobbs PJ, Frostagård A (1996) Changes in soil fungal:bacterial biomass following reduction in the intensity of management of an upland grassland. Biology and Fertility of Soils 22, Frostegård A, Bååth E, Tunlid A (1993) Shifts in the structure of soil microbial communities in limed forests as revealed by phospholipid fatty acid analysis. Soil Biology & Biochemistry 25, Lee Y-B, Lorenz N, Kincaid Dick L, Dick RP (2007) Cold storage and pretreatment incubation effects on soil microbial properties. Soil Science Society of America Journal. In press. MIS Operating Manual (2002) Sherlock Microbial Identification System, Vers (we have it in the lab on the shelf opposite of the HP GC 5890). Schutter ME and Dick RP (2000) Comparison of fatty acid methyl ester (FAME) methods for characterizing microbial communities. Soil Science Society of America Journal 64, Zelles (1996) Fatty acid patterns of microbial phospholipids and lipopolysaccharides. In: Schinner F, Öhlinger R, Kandeler E, Margesin R (eds). Methods in soil biology. Springer Pub. Berlin. 8