PROTOCOL FOR PLFA (+NLFA)

Transcription

1 1 of 7 PROTOCOL FOR PLFA (+NLFA) Kuzyakov Lab Michaela Dippold Callum Banfield Michaela Dippold Yakov Kuzyakov Rev. Date Prep. Modify. Check. Approv.

2 2 of 7 Introduction: To minimize cross contamination, use gloves and use tin foil to put your lids, etc on. Do not use paper towels. Never touch the tin foil on the mat side, try to pinch the foil on the other side. If you touch glassware which might get in contact with liquids, use acetone (100%) to rinse it. Always label glassware. Pasteur pipettes are for one-time use only. Required chemicals for rinsing (small bottles): acetone 1+2, MeOH 1+2; Tol 1+2, EtOH for cleaning glassware. First day: What to do Hints Chemicals and glassware Preparation of the extraction buffer: Preparation of a one-phase-solution with chloroform, methanol, citrate buffer (v:v:v = 1:2:0,8) - therefore first produce the citrate buffer: produce 100 ml of a 0,15 M citric acid solution (this means 2.88 g per 100 ml flask) you can directly produce the solution in a glass beaker but a bit less than 100 ml solution should be added: then put a magnetic stir bar into it, mix it permanently and check the ph with a calibrated ph-meter during the addition of KOH until a ph of 4.0 (± 0.1) will be reached - for 8 samples: mix 60 ml of chloroform with 120 ml of methanol and 48 ml of the citrate buffer with ph 4: check that the solution has only one phase (no emulsion!) Preparation of the soil sample: - Put an initial weight of 6 g (± 0.2 g) of fresh, wet soil per 50 ml glass centrifuge tube (the soil should be 2 mm sieved and must be directly frozen after sampling) - add 25 µl of the first internal standard (IS 1) = PLFA 19:0 (Dinonadecanoylglycerol-phosphatidylcholin, 1 µg/µl), which is solved in MeOH - add 18 ml of extraction buffer to each centrifuge tube Citrate: 3,145 g if using monohydrate KOH ~ 3 pellets = ph4 Pour a few pellets into the lid for easier handling. Rinse stir bar, first with acetone, then water. Fill up with Millipore water. Order of ph solutions: 7 4 4,66. Calibrate device: 7 and 4, only check with 4,66. Always rinse the ph electrode afterwards with deion. water and store in reservoir. Produce new solution every day. It may help to vortex the centrifuge tube shortly after adding extraction buffer to disperse the soil. Be careful with the electronic pipette, as it rereleases some of the volume. Wait for the beep. Normal syringe: Rinse 3x with acetone (1,2) and MeOH (1,2), then discard IS1 once. Make one extra hole into the lid for air flow (no vacuum). If you soil it, rinse it again (see above). Remove lid and re-crimp GC vial lid. Put syringe back, after you rinsed it first with MeOH, then with acetone. 100ml 0,15M Citric acid 60ml Chloroform 120ml MeOH KOH, Millipore water Stir bar, volumetric flasks*, beakers*. * = glassware wrapped in tin foil 18ml extraction buffer 25µl IS1 (12:0 NLFA standard) Centrifuge tubes*, lids, measuring cylinder* (20ml). Decapper, crimper First extraction: - put the centrifuge tubes horizontal on a horizontal shaker for 2 h with 225 rpm - centrifuge the samples for 15 min with 2000 rpm - transfer the supernatant with a glass pipette into a 250 ml separating funnel (glass funnel with teflon taps a bracket for 8 separating funnel should be installed in the fumehood) Second extraction: - add additional 6 ml of extraction buffer to each centrifuge tube - shake it again for 30 min on the horizontal shaker - centrifuge for 15 min with 2000 rpm If proceeding with NLFA, also add 12:0 NLFA standard (15µl) First extraction: 99% You can use a cardboard box; cushion the tubes with paper towels. Make sure to align the tube that the longest axis of the tube matches the movement of the shaker. 160rpm is enough. Make sure to counterweight the centrifuge tubes. Make sure that the rubber parts in the centrifuge are properly installed and match the tubes; Unscrew cap somewhat (otherwise it could be really hard to do it later). Check if the tubes cases are able to move. Wait for it to reach final rpm. Easy with the tubes after centrifugation, you do not want to disturb them. Put them in beakers, upright position. Separating funnel: Rinse Teflon tap with acetone. Optionally, add 6ml of Chloroform to check for leakiness. If possible, choose the same type of funnel. Make sure not to aspirate centrifugate. Second extraction: 99,9% Use a box for carrying the funnels. Check that screw is still tightly installed. 2h 160rpm 15min 2000rpm Teflon taps, separating funnels*, lids*, bracket. Beakers. (6ml Chloroform) + 6 ml extraction buffer 30min 160rpm 15min 2000rpm

3 3 of 7 - take the supernatant with a Pasteur pipette of and combine it with the previous extract in the separating funnel: Production of the two-phase solution: - add to the separating funnel 6 ml chloroform and 6 ml citrate buffer - shake the funnel for 1 min intensively by hand - then put the 8 funnels on the horizontal shaker and shake with 225 rpm for 15 min - put the separating funnels back to the brackets and wait until a phase boundary has evolved Separation of the two phases: - open each separating funnel and run the downer phase into 25 ml copped bottom flasks - add additional 6 ml of chloroform, shake 1 min by hand, then put again for 15 min on the horizontal shaker - put the funnels back to the brackets and wait overnight, until a complete separation of the two phases has occurred - next day in the morning: run again the downer phase and combine with the first time in the 25 ml copped bottom flask Reduce the volume of the downer chloroform phase in the flask with the rotation evaporator to less than 0.5 ml (40 C, pressure mbar) sample can now be stored (frozen) or directly continued with the purification Shake by hand, carefully open tap to eject gas. Use other horizontal shaker, secure funnels with rubber band and paper towel. Check if they re installed properly after switching the machine on. Again, carefully eject gas to prevent lid from coming off. The degassing will inevitably push some liquid out of the funnel; allow it to flow back into the funnel. Rinse lower part, if you touched it. Keep dirt/particulate matter in the funnel. When running the downer phase (i.e. lowest), make sure to remove the lid. The copped bottom flasks can remain in the fumehood during the night. Switch on heating of rotation evaporator. First, open valve for cooling water, then switch on pump. Reverse, when switching off. Make sure that the start value is 600mbar, sometimes people forget to reset this. Use brownish flask with chloroform to clean inside of device (~300mbar). Reset pressure. Install sample flask, secure it, set to 600mbar. Be patient, you do not want to get boiling retardation. Decrease pressure in 25mbar increments. If you use two rotation evaporators, one of them receives slightly warmer cooling water (serial connection), so this one needs a slightly lower pressure setting. Set to medium rpm ( calm fluid level) and submerge flask 0,5cm. In between samples, clean using the brown flask ( mbar, see above). Always check if the sample is boiling. Always check the flow in the waste container. Final mbar value: ~ 350. If content of flask runs dry, simply add 1-2ml of Chloroform and ultrasonicate. Measuring cylinder* (10ml) 6ml Chloroform +, if not added earlier!! 6ml Citrate buffer Chloroform Lids*, copped bottom flasks*, put them in cork rings. Chloroform for cleaning Second day Preparation of the purification columns: - a baker column with an inner diameter of 10 mm has to be installed on a baker system with Luer taps (stainless steel or Teflon, both possible) - a glass fiber filter has to be installed on the bottom of the column (alternatively a dense mat of annealed (300 C) glass wool can be placed on the bottom of the columns) - the column should be washed with one pipette (~ 1 ml) of methanol and one pipette of chloroform - activated (that means heated for 120 C overnight) silica (silica gel Merck, silica 60, particle size 0,063-0,200 mm should be filled into the columns: the silica in chloroform should have a length of 1,5 cm in the column (activated silica can be stored in chloroform (in an desiccator), if the chloroform is added directly after cooling down: then it can be transferred into the column with a glass pipette with a broken tip: Luer taps should be closed during transferring the silica into the column) - add once again glass wool or a glass fiber filter on top of the silica: take care that during preparation, sample addition and elution the silica never gets dry: it will lose its separating properties for PLFAs if it gets deactivated by air Produce activated silica gel by taking half or twothird filled bottle, put it into the furnace overnight and then add/soak it in chloroform. Store in desiccator. Make sure, you have annealed glass wool. There are two types of columns; smaller ones are for amino sugars! Install column and apply loads of chloroform, then check Luer taps for leaks. Open tap to rinse. Then rinse with MeOH, then again with chloroform. Apply glass wool, rinse glass stick with acetone, make sure it s not too dense/thick to prevent flow obstruction. Rinse with chloroform. Mark 1,5cm level. Be patient when adding silica gel, as it takes a moment to settle. Never ever should the silica gel be dry. If necessary, clear silica gel off the column wall with chloroform. Add top glass wool and chloroform, then open tap to adjust the fluid level to the marking. Rinsed sample flasks, 3x3 times with chloroform. When adding acetone, be careful and pour it along the wall. When eluting with MeOH, make sure to do it slowly. Stop all taps at the same Glass wool, silica gel; Chloroform, acetone, methanol. Waste containers, coppedbottom flasks, ruler, pen. Luer taps, columns, table. Broken Pasteur pipette.

4 4 of 7 Separation of the Lipids: - transfer the sample from the copped bottom flask onto the column with a pasteur pipette - rinse the flask 3 times with ~ 200 µl of chloroform and transfer this chloroform also onto the columns time when refilling. Note down, how often you have added MeOH. Keep elution rate constant. (Elution of neutral lipid fraction into pointed bottom flask, transfer using chloroform) Chloroform, acetone, MeOH if NLFA are to be analysed: Instead of waste devices, but a pointed bottom flask under the columns. Open tap carefully, so that the PLFA will have time to bind to the column and (possibly present early ) NLFA are collected. As soon as the chloroform level is just above the silica gel level, close the tap. Elution of the neutral lipids: now add 5 ml of chloroform and elute into flask the very same flask for NLFA. Waste containers, copped bottom flasks see above Proceed with glycolipids. See below if NLFA are not interesting to you: - put the waste devices under the columns and open the Luer taps slowly: chloroform should elute slowly, so that the PLFA will have time to bind to the column; close the Luer tap, if the last sample has disappeared in the glass wool or filter - Elution of the neutral lipids: add 5 ml of chloroform to the column and elute to waste - Elution of the glycolipids: add 20 ml of acetone to the column and elute this acetone to the waste - Elution of the phospholipids: Replace the waste by 50 ml copped bottom flask, than add 4 times 5 ml of methanol to the column and elute slowly (not faster than two drop per second) into the flasks (always close the Luer taps between the elution of the next 5 ml) Preparation of the final PLFA sample - Reduce the volume of the sample in the flask with the rotation evaporator to less than 0.5 ml (40 C, pressure mbar) - transfer the sample to 5 ml Reactivials rinse the flask 3 times with ~ 100 µl of methanol and combine with the sample in the Reactivial - dry the sample under a gentle nitrogen stream - close the Reactivial with a Teflon septa and store the PLFA sample frozen until derivatization (storage for 2 months without loss tested) Start value: 400mbar, Final value: 300mbar. Rinse with MeOH in between with 200mbar. 3x3 times rinse, Teflon septa towards sample; Teflon: thinner layer. Freeze. Careful/easy with the gas flow. MeOH Reactivials, lids, teflon inlets. Optional: Tray for reactivials. N2 apparatus with Pasteur pipettes. (repeat first and second day for additional 8 samples 16 frozen PLFA-samples are prepared in Reactivial for derivatization) Third day Preparation of the external mixed standards: - see attached table of the concentrations of stock solutions (~ 1 mg/ml) if there s no mixed standard prepared first prepare a mixed standard of a concentration of 30 ng / µl by mixing 600 µl of each substance (including the IS1) into a 20 ml flask and filling up with dried MeOH (this standard should be stored at -20 C in closely crimped GC Vials - now prepare your six external standards by filling the following volumes with a microliter pipette into 5 ml Reactivials : Std1: 50 µl ( 1.5 µg Std), Std2: 150 µl 4.5 µg Std), Std3: 300 µl ( 9 µg Std), Std4: 600 µl ( 18 µg Std), Std5: 800 µl ( 24 µg Std), Std 6: 1000 µl ( 30 µg Std); if sample peak areas are more than 10% above that line add an additional higher standard (or replace the lowest by an higher standard) - dry the standards under a gentle nitrogen stream (can be stored like the samples) Hydrolyzation of the PLFA: - Produce the water-free hydrolyzation reagent: 0.5 M NaOH in MeOH: therefore first put a certain amount of dried MeOH into a volumetric flask, add the needed amount of NaOH-pellets (stored dry!), fill up with dried MeOH (and store on annealed molecular sieve in an exiccator) - add 0.5 ml of the 0.5 M NaOH to each of the 16 samples and 6 external standards - dissolve the sample/standard in the methanolic NaOH by ultra-sonification for ~10 min - put the properly closed vials into the pre-heated (constant temperature) Check if there is an external standard prepared. Use 100ml flask, 2g NaOH and MeOH for HPLC (= dried MeOH). Use special displacement pipette (set to 0-5-0). Check that temperature = const. Dispose of pipette tips toxic waste Change temperature to 80 C Standards. MeOH N2 apparatus with Pasteur pipettes. GC vials, crimping equipment, decapper NaOH and MeOH for HPLC displacement pipette

5 5 of 7 derivatization block at 100 C for 10 min (! if you sample evaporates it is dead!) - wait before opening, until vials are cooled down to room temperature! Methylation of the fatty acids: - add 0.75 ml of 12.5 M BF3 in methanol to each sample and external standard (pay attention: toxic and mutagenic: nitrilic gloves are obligatory for that step!) - put the properly closed vials into the pre-heated (constant temperature) derivatization block at 80 C for 15 min - wait before opening, until vials are cooled down to room temperature! Separation of the fatty acid methyl esters (FAME) from the head groups - add 0.5 ml of saturated NaCl solution (simply prepared in water until no more NaCl can be dissolved) to each vial this causes hydrolyzation of the not-used BF3 and precipitation of the formed HF as NaF (so toxicity is strongly reduced) - add 1 ml of Hexan to each vial, close the vial properly and shake each vial intensively for at least 30 sec; If no good phase separation in the vials occurs it helps to centrifuge them for 5 min at 500 rpm (Reactivials will survive that without damage) - carefully remove the upper apolar layer into a new Reactivial without taking any of the polar phase - repeat that hexane-extraction 2 additional times and combine the 3 ml hexane extract in the vial - - dry the hexane extract under a gentle nitrogen stream Preparation for measurement: - add 15 µl of the first internal standard (IS 2) = FAME 13:0 (fatty acid methyl ester 13:0, 1 µg/µl) per each vial (sample and standard), which is solved in toluol - add 185 µl of toluol per vial - dissolve in the sonification bath for 10 min - transfer to a 100 µl inlet for a 1.5 ml GC-Autosampler-Vial - screw or crimp the vial properly vials can be stored at -20 C for one year until measurement BF3 needs to be stored in a desiccator. Use special displacement pipette (set to 0-7-5). Use wet paper towels to cool block down. Use special displacement pipette (set to 0-5-0). Add about 1 Pasteur pipette of Hexan. When transferring the upper phase, always check if there is a phase boundary within your Pasteur pipette! If yes, return and retry. Rotate Reactival to that the liquid inside absorbs all the tiny droplets on the inner wall coatings. Label GC vial with sticker. BF3 Gloves, lab coat. Displacement pipette Hexan, NaCl Displacement pipette, N2 apparatus, new reactivial. IS2, Toluol Syringe, GC vial, inlet for GC vial. Crimper.

6 6 of 7 Appendix Straight chain saturated fatty acids concentration of FA (µg µl) 14:0 1, :0 1, :0 1, :0 0, :0 0, :0 1, Cyclopropyl branched fatty acids Cy19:0 1, Cy17:0 0, Methyl branched saturated fatty acids 10Me16:0 1, Me18:0 1,34 Anteiso branched saturated fatty acids a14:0 1, a15:0 1, a16:0 1, a17:0 1, Iso branched saturated fatty acids i14:o 1, i15:0 1, i16:0 1, i17:0 1, Monounsaturated straight chain fatty acids 14:1w5c 1, :1w5c 1, :1w7c 1, :1w9c 0, :1w7c 1, :1w9c 1,

7 7 of 7 Polyunsaturated straight chain fatty acids 18:2w6,9 1, :3w3,6,9 1, :4w6c 1, IS1 = Dinonadecanoyl-Glycerindiphosphat 1,000 µg/µl