VI. Protocols

A. TIPS FOR LIPID RESEARCH (parts adapted from P.K. Stumpf)

Suggested readings

· Chapter 10: Lipids. In “Biochemistry and Molecular Biology of Plants” (Buchanan, B. and Gruissem, W., eds.), American Society of Plant Physiologists, Rockville, MD

· Christie, W.W. (1982). Lipid Analysis. Pergammon Press, Oxford.

· Christie, W.W. (1995). In “Advances in Lipid Methodology. Volume 2.” p195-213.

· http://www.lipid.co.uk/

· Kates, M. (1972) Techniques of Lipidology: Isolation, Analysis and Identification of Lipids North Holland/American Elsevier.

General Tips: Logistical

1. Solvents splashed into your eyes can cause severe damage to your vision. The wearing of safely glasses is recommended, even for small volume manipulations.

2. Generally avoid plastics, as they are often incompatible with organic solvents (especially for prolonged storage of organic solutions), and because a large amount of plasticizers, etc. can often be extracted, sometimes interfering with analysis, especially GC analysis. However, if you are only going to analyze radioactivity from extracts, use of polypropylene screw cap tubes (e.g. Falcon tubes) can be convenient for lipid extractions.

1. Never store any alkaline solutions in glass bottles with glass stoppers. The stoppers will invariably freeze in the neck of the bottle.

2. For small scale organic extractions we have a large number of glass screw-capped 13, 16 and 20 mm diameter tubes which are ideal for this purpose. Check tube lip for chips. Make sure you use the caps with teflon-liners when using organic solvents.

3. Never use a rubber stopper or parafilm to cap a tube containing an organic solvent. The solvent will leach out hydrocarbons. Use a Teflon stopper or Teflon lined screw cap or an aluminum foil coated cork stopper. Glass stoppers that fit tightly are also OK.

4. We recycle the screw-capped glass tubes and teflon-lined caps used for lipid extractions, derivations and sample storage. The quality of our work depends on the proper cleaning of these tubes, etc. When you have completed your work, the tubes should be rinsed two or more times with a water-miscible organic solvent (ethanol, acetone) to remove almost all of the lipid or fatty acid residues. Then place them in the correct wash basin for cleaning. If you put tubes in the wash with large amounts of lipid residues then the detergent simply spreads the lipid throughout the wash glassware, and it is not completely rinsed away! NOTE: The disposable glass screw cap centrifuge tubes (5 ml and 10 ml) with conical tips are extremely difficult to clean. Therefore, rather than recycling or washing, these should be disposed to avoid the risk of cross-contamination.

5. For repeated organic extractions it is useful to have at your bench your own small solvent bottles, (with solvent dispensed from the 4 liter bottles kept under the fume hood). Once you have poured some solvent into your own container, never pour unused solvent back into the main bottle. Label your own bottles correctly: full chemical name, ratios for solvent mixtures, name and date; and check that you have a teflon-lined cap.

6. Waste solvents. We have two solvent waste containers in the fume hood for organic wastes (one general purpose, one for solvents containing halogenated organic solvents). There is also a waste container for organic solvents with radioactivity.

7. Keep all tubes and solvents tightly capped except when in actual use. Note that solvent mixtures (e.g. chloroform:methanol) can change composition with long time storage if not tightly capped.

8. We have limited fume hood space in the lab (See ‘Use of the Fume Hood’ protocol.) Thus, working at the bench with solvents may be acceptable in some cases if certain precautions and conditions apply:

* where the lab air flow is good.

* when only small volumes of solvent are being used.

* when there are no naked flames on nearby benches.

* in this context, hexane-isopropanol is a much less toxic solvent mixture than chloroform-methanol, so the former is preferred if bench-work seems necessary.

General Tips: Chemical

1. Never assume that a solution of Na₂ ATP is neutral; it is instead highly acidic. Bring to pH 7.0 before using.

2. NADPH and NADH should not be stored in solution over any period of time. Maintain solutions at about pH 8.0, never below 7.0; otherwise rapid auto-oxidation will occur.

3. Do not neutralize CoA stock solutions. You will encourage disulfide formation. Maintain CoA solutions at their own pH (<7) and allow your reaction buffer to neutralize.

4. Never store acyl CoAs at pHs above 6.5. Rapid hydrolysis will occur. Store at pH 6.0, at the highest. Neutralizing with conc. NH₃ with result in formation of acylamides.

5. If you want to analyze for lipid classes i.e. phospholipid, etc., keep your reaction acidic. Above pH 8.0 at room temperature rapid deacylation of PL  free fatty acids will occur.

6. Store methyl esters of fatty acids in toluene. If polyunsaturated fatty acids are present, flush tube with N₂. Never use diethyl ether for storage which will cause auto-oxidation by ether peroxides. Petroleum ether is a poor solvent for polar methyl esters of hydroxy fatty acids.

7. Never store fatty acids in absolute ethanol or methanol. Esters will form. 80% alcohol with pH >7 is OK to provide a solution which is miscible with water. Ammonium salts of fatty acids are more soluble in water than free fatty acids or other salts. These can be prepared by adding 5-10 μl NH₄OH to fatty acid in a tube and evaporating the excess NH₄OH

8. Extracts of whole tissues will usually contain adequate endogenous antioxidants to protect unsaturated fatty acids in lipids from oxidation. However, once fatty acids or lipid classes are isolated, and thus separated away from the endogenous antioxidants, it is preferable to add a trace amount of BHT (50-200 μg) as an antioxidant for storage of solutions. (Note: BHT will give a peak on GC near 14:0). To minimize oxidation and photo-oxidation store lipid and fatty acid samples in the freezer in screw-capped tubes or vials. Do not use diethyl ether or other solvents that are prone to accumulate peroxides. Typically use heptane or toluene instead. Use small containers to minimize dead-space. For long term storage of UFA and for added protection against oxidation you can flush tube with nitrogen before storage.

9. When performing a lipid extraction from tissues the important point is to use enough extracting solvent to form one phase. To ensure extraction of free fatty acids, the pH must be <4.

USE OF THE FUME HOOD

The lab is busy and we only have one fume hood, so additional cleanliness and courtesy are required.

General notes

1. Because of space limitations manipulations that do not require a fume hood should NOT be done in the fume hood.

2. When working, do not hog the center - consider that there are two side-by-side spaces, and take only one.

3. For continual use of a large area, post a notice beforehand.

4. One set of glass measuring cylinders, marked with tape, will be kept for volatile organic solvents only (not water, glacial acetic acid, buffers etc.). Glassware used ONLY for organic solvents – allow the solvent to evaporate and return the glassware to the drying rack.

5. As soon as is practical, move your stuff out of hood. Label any stuff left in that you are not immediately attending, otherwise it may be tossed.

6. The hood is labeled with radioactive marker tape. However, when using high amounts of radioactivity, or particularly dangerous chemicals, additional labeling is required.

7. Nothing is to be “perched” on the metal lip of the fume hood.

8. Obviously, clean up any spills and contamination promptly. This includes contamination at the ends of the nitrogen lines.

When not in use there should only be the following in the fumehood:

1. Nitrogen evaporator

2. Two 4 liter glass bottles for solvents, one general, and one for solvent mixtures containing halogenated hydrocarbons (See below).

3. On the shelves to the right hand side:

Lower shelf: bottles for chloroform-phenol wastes from plant tissue extractions, and for ethidium bromide, and a container for “general wastes.”

Upper shelf: reagents.

4. There may be TLC tanks and (named) wastes evaporating at the rear, but these should not be left for long: certainly no longer than overnight.

TLC.

1. TLC should always be run in the fume hood.

2. Empty tanks are now kept in the cupboard to the right of the fume hood and hydrogen tank, under the GC.

3. When in use the TLC tank should be labeled with your name, date and with the solvent mixture.

4. When done, pour the solvent mixture into the correct waste solvent bottle, allow the tank (and any blotting paper therein) to dry, remove your label, and return the tank to its proper storage place.

5. TLC tanks should always be placed parallel to the airflow, towards the back of the hood.

6. Generally, if you wish to keep the solvent (a practice I don’t particularly recommend with volatile solvent mixtures), then pour it into a labeled bottle with a proper solvent-resistant cap, and keep the mixture on your bench, not in the fume hood.

Organic Solvents.

1. We keep two 4 liter waste solvent bottles in the fume hood. One is for general organic solvents or solvent mixtures, one is for solvents or solvent mixtures containing anything more than traces of halogenated organics (chloroform is the most common one we use, but this includes dichloromethane, carbon tetrachloride, alkyl bromides etc.).

2. Each bottle will have its original label defaced, and space on the bottle to write down what you add to the mixture. Phenol-water mixtures and methylpyrrolide mixtures can be added to the general wastes bottle; TLC tank mixtures containing chloroform should be added to the chlorinated solvent bottle....etc.

3. Chloroform-phenol wastes containing plant debris - use the bottle on the lower right hand shelf. Likewise there is a special container for ethidium bromide wastes.

4. If you are concerned about chemical compatability, or any other chemical disposal question, see Mike.

5. No large amounts of strong acids or bases should go in these bottles, nor radioactive organic wastes, nor heavy metal wastes! These should be disposed of separately.

6. When you dispose of organic solvents please list the chemical on the bottle if it is not already marked down, and put some indication of amount you add (in ml). Include water in the list. Use common sense about writing down tiny amounts, traces of buffer or other solutes, or about disposing of small amounts of water soluble solvents or water-saturated aqueous phases.

Quino is currently in charge of chemical waste disposal (11/00). He will see that nearly full bottles get picked up by ORCBS.

B. LIPID EXTRACTIONS

General Principles

Extractions of total lipids from tissues or enzyme extracts is accomplished by extraction with organic solvents, most commonly hexane-isopropanol or chloroform-methanol mixtures. Intact plant tissue often needs to be “quenched” (short heat treatment with isopropanol) to inactivate any lipases prior to extraction. The tissue is then homogenized and solvent added to make a monophasic solvent mixture containing any residual water from the tissue. Upon extraction, a salt solution is added to the monophasic mixture to cause a phase separation, with the lipids partitioning into the organic phase. The solvent is then evaporated to dryness. Wash and re-extraction steps are often included in protocols.

Notes

· Large scale extraction of dried oilseeds is usually undertaken with warm hexane using a Soxhlet extractor. This will extract neutral lipids and most polar lipids.

· There are many solvent extraction protocols in the literature. Very polar lipids (some sphingolipids, lysophospholipids) may require forcing conditions to extract quantitatively. Christie (1982, 1995) covers some of these, and a detailed discussion of extraction artifacts etc.

· Acyl-CoAs partition into aqueous phases but are notorious for sticking to proteinaceous organic-aqueous interfaces, or in emulsified particles suspended in the organic phase. For a protocol which recovers acyl-CoA and acyl-ACP along with lipid extractions see Mancha et al. (1975).

· When performing a lipid extraction from tissues the important point is to use enough extracting solvent to form one phase. To ensure extraction of free fatty acids, the pH must be <4.

Literature

Bligh, E.G. and Dyer, W.J. (1959). Can. J. Biochem. Physiol. 37: 911-917.

Christie, W.W. (1982). Lipid Analysis. Pergammon Press, Oxford.

Christie, W.W. (1995). In “Advances in Lipid Methodology. Volume 2.” p195-213.

Folch, J., Lees, M. and Sloane-Stanley, G.H. (1957). J. Biol. Chem. 226: 497-509.

Hara, A. and Radin, N.S. (1978). Anal. Biochem. 90: 420-426. Or “Methods in Enzymology” (1981) 72: 5-9.

Mancha, M., Stokes, G.B. and Stumpf, P.K. (1975). Anal. Biochem. 68: 600-608.

Hexane-Isopropanol Method

See Hara and Radin (1978).

Heat the tissue (up to 400 mg) in a small volume (3 ml) of isopropanol at 80-85C for 5-10 minutes to inactivate lipases. Plant tissues are notorious for containing lipases that remain active in aqueous-organic mixtures, so this step is sometimes crucial if you wish to conduct lipid class analyses. Grind the tissue using a mortar and pestle, or glass grinder, or Polytron. Then wash the grinder with the remainder of the isopropanol (1 ml) and hexane (6 ml) to give the correct volumes of water:isopropanol:hexane 1:4:6 for a monophasic system. If anything, err on the side of too much isopropanol to ensure mixture is monophasic. The grinding step is omitted (and usually also the quenching step) when extracting assays with cell free extracts.

After shaking and allowing to stand for a few minutes*, add 5 ml aqueous sodium sulphate solution (6.6 g anhydrous sodium sulphate in 100 ml water), shake vigorously and allow to phase separate. Pipet the uppermost phase to a clean tube. If there are three layers, after removing the uppermost layer add another 1-2 ml of hexane, shake, and after phase separation pipet the new uppermost layer and combine with the first. Then add 2:7 isopropanol:hexane (4 ml), shake, allow to phase separate, and combine this upper phase with the organic phases. Evaporate to dryness.

* At this point, if highly accurate determinations of lipid content are required, the organic phase may be pipetted away, and the tissue re-extracted with more 3:2 hexane:isopropanol.

Note: This method has the advantages of: less toxic solvents (than chloroform); faster evaporation because of more volatile organic phase; upper organic phase is more easily removed than removing chloroform from the lower phase. Disadvantages are that larger volumes of organic solvent are needed to achieve monophasic state and that polar lipids are less well extracted (lyso-phospholipids and sometimes polar lipids like PC).

Chloroform-Methanol Methods

A. See Bligh, E.G. and Dyer, W.J. (1959).

Adjust quenched tissue to 1 ml and 0.15 M acetic acid.

Add 3.75 ml of methanol:chloroform (2:1).

Use polytron, or vortex or other method to homogenize.

Add 1.25 ml chloroform.

Add 1 ml water.

Vortex to mix.

Centrifuge to separate phases.

Remove lower phase with glass pipette to suitable vial.

Extract remaining aqueous phase again with 2-3 ml hexane. Remove upper phase and combine with chloroform phase.

B. See Folch et al. (1975).

The quenched tissue is homogenized in methanol (3 ml) then the homogenizer washed with chloroform (6 ml) which is added to methanol. The mixture is vortexed for a minute or two. If there is much debris, spin and transfer the solvent to a clean tube. Rinse the residual debris with 2:1 chloroform-methanol (3 ml) and transfer to the original chloroform-methanol extract. Add 4 ml of 0.88% aqueous KCl, shake, and allow to phase separate. If there is no debris at the interface*, remove the upper aqueous layer. Wash the lower chloroform layer with 1:1 methanol:water (2 ml). Pipet out the aqueous methanol wash phase, and evaporate the chloroform layer to dryness. * If there is debris at the interface, transfer the lower chloroform layer to a clean tube, together with a chloroform wash of the debris. Then wash the combined organic phase with 1:1 water:methanol etc.

USE OF EVAPORATOR/NITROGEN GAS TANKS

The 12-line evaporator/water bath is set up in the fume hood with the nitrogen tank. There is a T-piece in the nitrogen line, to give an additional nitrogen line independent of the evaporator. Usually, this is switched off (with a screw clamp), but be aware if this is in use. If it is in use, cooperate with the person using the additional line, since changes in gas pressure/flow rates will result if you also use the evaporator.

It is not necessary to have very high flow rates of nitrogen blasting through each tube. If the solvent surface is “faintly dimpled” by the gas flow that is more than sufficient. Move the needles down as the level of solvent diminishes in the tube. Also, in most circumstances use the water bath since the sample will be cooled by evaporation.

When finished:

* clean the ends of the needles by bubbling through ethanol.

* close all the needle valves.

* make sure the water bath is switched off.

* make sure the nitrogen tank is turned off at the main valve.

Nitrogen gas tank replacement: the last user is responsible for changing the gas tank, and reordering new tanks if necessary. Reorder tanks if, after you have changed a tank, there is only one full nitrogen tank left in the racks. Indicate on the reorder sheet the number of empties to be picked up. Order at least two full tanks, since we can go through nitrogen gas quickly. When replacing the empty tank in the racks, make sure there is an EMPTY sign on it. If you use the evaporator and there is no pressure on the main nitrogen tank valve, even if there is still gas flow and pressure in the secondary valve, change the tank.

C. PREPARATION OF FATTY ACID METHYL ESTERS (FAMEs)

General notes

· Samples that are predominantly TAG do not dissolve well in methanol; therefore 30% toluene can be added to aid solubility.

· Normally, an internal standard (e.g. 17:0) is added as free fatty acid before methylation, or as methyl ester for transesterification.

1. Acid catalyzed esterification and transesterification

A. Boron Trifluoride Methanol

Background and notes

· This is the most commonly used method for preparing fatty acid methyl esters. The boron trifluoride methanol method can convert both free fatty acids and fatty acid esters to fatty acid methyl esters. This procedure uses BF₃ as a Lewis acid catalyst. Excess BF₃ is removed as a gas. BF₃ reacts with H₂O to form Boric acid which in turn reacts with methanol to form methyl borate, a volatile ester.

· Free fatty acids react much faster than esters. Five min. at 70-90 °C is enough.

· Use the hood: BF₃ is not good for living tissue.

· Because of high temperature involved, this method is not suitable for preparing fatty acid methyl esters if the acyl chains contain any function groups that may be sensitive to high temperature.

· If only small amount of tissue is used, lipid extraction can be omitted; fatty acid methyl esters can be prepared directly from tissue with this method. Browse et al. (Analytical Biochemistry 152:141-145, 1986)

· Methyl esters of short and medium chain fatty acids are volatile and can be lost during evaporation process. To avoid losses, keep cool and do not over evaporate. If you are trying to measure 14:0 or shorter chain lengths, add 13:0 as an additional internal standard to monitor losses.

Procedure:

1. Transfer lipid extract to a screw cap tube with Teflon lining. Be sure the edges of the tube are not chipped since a tight seal will not be possible with a defective tube. Evaporate off the solvent with N₂.

2. (Optional) Add 100μl of toluene and vortex briefly.

3. Add 1 ml of Boron Trifluoride Methanol (10% boron trifluoride in methanol, Sigma, Cat# B-1002, which should be stored at 4°C).

4. Cap tightly and place the tube at 90⁰C for 40 minutes. **All oxygen esters and free acids will be transesterfied or esterfied to methyl esters under these conditions. If you have lost considerable volume during the procedure, you probably had a chipped tube. Discard tube and sample and start over.

5. After the reaction is completed, cool the tube, at least, to room temperature.

6. Remove the cap and add 1 ml of water.

7. Extract 3X with 3 ml of hexane by shaking the capped tube vigorously. Allow separation into two phases and remove top phase with a pasteur pipette to a new tube. **If you are working with hydroxy fatty acids, use diethyl ether as the extracting solvent.

8. Evaporate the solvent with a stream of N₂ to dryness, but not over dry.

9. Dissolve the fatty acid methyl esters into desired volume of hexane for GC or any other analysis.

B. Esterification/transesterification with Methanolic:HCl

Free fatty acids are esterified and O-acyl lipids transesterified by heating them with a large excess of anhydrous methanol in the presence of an acidic catalyst. If water is present it may prevent the reaction going to completion.

Procedure:

1. Transfer lipid extract (up to 50 mg) to a screw cap tube with Teflon lining. Be sure the edges of the tube are not chipped. Evaporate off the solvent with N₂.

2. Add 1 ml of the methanolic HCl (1 M) reagent*, purge the tube with N₂ briefly, and seal with a teflonlined cap.

3. Heat at 90°C for 60 minutes.

4. After the reaction is completed, cool the tube, at least, to room temperature.

5. Add 1 ml of hexane and 1 ml of 0.9% NaCl, and vortex vigorously.

6. Centrifuge briefly and transfer the organic phase to a new tube.

7. Extract the mixture two more times with 1 ml of hexane each.

8. Combine the organic phases and evaporate the solvent with a stream of N₂ to dryness

9. Dissolve the fatty acid methyl esters into desired volume of hexane for GC or any other analysis.

* METHANOLIC-HCL is produced by dissolving HCl gas in methanol and can be purchased from Supelco and diluted to 1 N with methanol:

33355 METHANOLIC-HCl (3N) KIT,20X1ML

33051 METHANOLIC-HCl (3N) KIT, 10X3ML

33050U METHANOLIC-HCl (3N) 400ML

Note: This method can be applied to fresh tissue. For detailed information, please refer to the paper by John Browse et al. (Analytical Biochemistry 152 141-145, 1986

2. Base-Catalyzed Transesterification

O-acyl lipids are transesterified very rapidly in anhydrous methanol in the presence of a basic catalyst. Free fatty acids are not esterified, however, and care must be taken to exclude water formation as a result of hydrolysis of lipids.

Procedure:

1. Transfer lipid extract (up to 50 mg) to a screw cap tube with Teflon lining. Be sure the edges of the tube are not chipped. Evaporate off the solvent with N₂.

2. Add 2.5 ml of sodium methoxide in anhydrous methanol, and 2.5 ml of heptane.

3. Vortex at room temperature for 2-5 minutes.

4. Extract the mixture three times with 2.5 ml of hexane each.

5. Wash the combine the organic phases twice with 3 ml of water each.

6. Remove residue water by passing the organic phase through a anhydrous sodium sulphate column packed in a pasteur pipette (optional).

7. Evaporate the solvent with a stream of N₂ to dryness, but not over dry.

8. Dissolve the fatty acid methyl esters into desired volume of hexane for GC or any other analysis.

D. THIN LAYER CHROMATOGRAPHY (TLC) OF LIPIDS

Ref.: Christie, W.W. (1982) Lipid Analysis, 2^nd edition Pergammon Press.

Introduction

Thin layer chromatography (TLC) is used to achieve a wide range of analytical and preparative separations of fatty acids and lipids. The most commonly used stationary phases are silica gel, for “normal-phase” separations based on analyte polarity, and C18-coated silica, for “reversed-phase” (RP) separations based on analyte hydrophobicity. The adsorbent layer may also include an inorganic fluorescence indicator.

General notes

· The amount of lipid loaded (μg/cm) will depend on the type of separation, thickness of the adsorbent layer, and capacity of the adsorbent layer. Lower loadings are required for polar lipids on silica and for reversed-phase TLC plates. Examples are given later. High loading can sometimes be mitigated by multiple solvent developments.

· Load lipid samples as dilute solutions (typically 10-50 mg/ml). It is important to use a solvent that will dissolve your material but evaporate quickly so that the loading zone with remain as narrow as possible. In addition, when loading onto silica, use the least polar solvent possible to minimize band spreading. Usually you can dissolve your neutral lipid samples in hexanes, and polar lipid samples in hexane or acetone. A micro-syringe or a glass micropipette are preferred over a pipettor for maximum control while loading. Additionally, use of organic solvents with pipettor tips could cause leaching of the plastic inside the tip.

· Unsaturated lipids are fairly stable on TLC plates for several days if kept out of light and air. However, we have found that on the reversed-phase TLC plates this is not so. Care should be taken to remove these lipids within 24 hr of being loaded on the plate, especially if polyunsaturated lipids or fatty acids are to be recovered.

Types of TLC plates

Brand: Whatman (or equivalent)

http://www.whatman.plc.uk/products/analytical/chromatography/a_pd_chrom_0**.html

Code Size Thickness Minimum amount*

Analytical

K6 silica gel 60 Å 5 x 20 cm 250 μm 1 full box of 75

K6 silica gel 60 Å 10 x 20 cm 250 μm 1 full box of 50

K6 silica gel 60 Å 20 x 20 cm 250 μm 2 full boxes of 25

K6F silica gel 60 Å 20 x 20 cm 250 μm 1 full box of 25

LK6F silica gel 60 Å 20 x 20 cm 250 μm 1 full box of 25

HPK silica gel 60 Å 10 x 20 cm 250 μm 1 full box of 50

Preparative

PK6F silica gel 60 Å 20 x 20 cm 1000 μm 1 full box of 20

Reversed-phase

KC18F 20 x 20 cm 200 μm 1 full box of 25

*Ordering is the last user’s responsibility. See section I-L, ‘How to order the supplies you need.’

Analytical K6 silica gel 60 Å adsorption TLC plates. K6 60 Å (pore size) and K6F plates provide high-purity silica gels and polarity for normal-phase separations. The F stands for the fluorescence indicator being bound to the silica; L stands for inert loading zone. Moderate layer hardness makes possible convenient spot recovery by scraping the silica off the plate. Silica gel thickness plays a critical role in governing the loading capabilities of a TLC plate. Analytical plates should be used for small loadings, typically less than of 500 μg/cm, and ususally 10-100 μg/cm. The commercial plates have excellent reproducibility, and negligible moisture uptake, so generally they do not needed to be activated by heating.

· Chemically and optically inert organic binder.

· Quality separation of nonpolar to strongly polar compounds.

· Wide applicability, neutral lipids, glycolipids and phospholipids (see solvent systems, below)

Preparative silica gel TLC plates. P stands for preparative. For small scale preparation of lipids use the analytical plates. They can take up to 10 mg neutral lipid/20x20 plate. Samples can be spread over 1 or several plates. Preparative plates take increased loading in proportion to increased thickness. Preparative plates (20x20) are ~$ 10.00 each whereas analytical plates (20x20) are ~$4.

High performance TLC plates. Whatman HPTLC plates can be used for your most challenging separations. These plates consist of a 4.5 um particle size silica gel plus an inert binder in a uniform 200 um layer on glass. They have narrow particle size distribution, homogeneity and overall uniformity. The results are good performance and reproducibility, giving your TLC high resolution and sensitivity. The HPTLC plates are (10x20) are ~$ 8.00

· Dense, uniform layer provides stable baseline in densitometry.

· Low band diffusion provides very compact sample bands.

· Microsample’s nanograms can be analyzed..

· Have a very precise loading zone so loading can take longer.

· Low mass capacity.

Reversed-phase TLC plates KC18 and KC18F. Whatman provides alkyl (C2, C8, C18) and phenyl bonded reversed-phase plates. The chain length of the bonded hydrocarbon functional groups primarily affects retention and the ability to accommodate the water content of solvent systems. The shorter carbon chain is used for increased polarity and affinity for aqueous solutions while the longer chains give greater retention and hydrophobicity. We generally use the C18 plates. KC18 (20x20) plates are ~$ 14.00 each. These plates also have a much lower mass capacity than the normal silica plates, roughly 50-100ug/cm.

Solvents and solvent systems

TLC should always be run in the fume hood. Lining the tank with filter paper will saturate the atmosphere inside with solvent vapor, which speeds up analysis time especially with polar solvents and can improve resolution. Empty TLC tanks are now kept in the cupboard to the right of the fume hood and hydrogen tank, under the GC. When in use the TLC tank should be labeled with your name, date and with the solvent mixture. Use forceps to place the TLC plate in tank as quickly as possible. The longer the lid of the tank is off the more your solvent system evaporates. Sometimes it might be useful to place weights over the solvent chambers to ensure solvent vapor does not escape. For the large TLC tanks 150-200 ml of solvent is adequate. There are also smaller cylindrical chambers for the 5x10 cm plates. When done, pour the solvent mixture into the correct waste solvent bottle, allow the tank (and any blotting paper therein) to dry, remove your label, and return the tank to its proper storage place. TLC tanks should always be placed parallel to the airflow, towards the back of the hood. Generally, if you wish to keep the solvent (a practice not recommended with volatile solvent mixtures), then pour it into a labeled bottle with a proper solvent-resistant cap, and keep the mixture on your bench, not in the fume hood.(See section VI, ‘Use of the fume hood’.)

Literature

There are a large number of solvent systems to use with a variety of absorbents for a large number of separations. Many of these can be found in:

Christie, W.W. (1982) Lipid Analysis, 2^nd edition Pergamon Press.

Kates, M. (1972) Techniques of Lipidology: Isolation, Analysis and Identification of Lipids North Holland/American Elsevier.

Neutral lipids

· To separate fatty acid methyl esters from free fatty acids:

K6 plates, half then full development with 90/10 hexane/diethyl ether.

· To separate triacylglycerols, diacylglycerols and free fatty acids:

K6 plates, develop half then fully with 70/30/1 hexane/diethyl ether/glacial acetic acid.

Polar lipids

For separation of all major chloroplast phospho- and galactosyl- glycerol-lipids use ammonium sulphate impregnated plates. Dip Whatman K6 TLC plate in 0.15 M ammonium sulphate for 30 s let air dry for 3 h. To activate place plate in 120 °C oven for 3 h then cool, load and run the plate in 91/30/8 (v/v/v) acetone/toluene/water on the same day. Do not use K6F plates by mistake, as the inorganic fluoro messes up the separation.

Figure 1 shows a typical separation of MGDG, PG, DGDG, SQDG, PS + PI, PE and PC.

Isolation of PC and MGDG. Load K6 or K6F silica plates and run in chloroform/methanol/ glacial acetic acid/water 85/15/5/2 (v/v/v/v). First develop until the solvent front has reached 1/3 of the height of the plate then let dry for 30 to 60 min; develop once more in the same solvent system this time until the solvent front has reached 2/3 the height of the plate. Once again let dry for 30 to 60 min and finally develop to within 1 cm of the top of the plate in 200/2/1 (v/v/v) acetone/glacial acetic acid/water.

Fatty acid methyl esters (FAME) by reversed-phase TLC

For fractionation of fatty acid methyl esters by chain length/number of double bonds. A cis-double bond has approximately the same effect on the partition coefficient of the fatty acid or lipid as removing two methylene groups. Therefore, you get critical sets such as 16:0/18:1 or 14:0/18:2/16:1 that cannot be resolved by TLC (though this can be achieved by HPLC).

For a standard separation of fatty acid methyl esters, develop KC18F reversed-phase TLC plates half way, air dry then re-develop fully in acetonitrile/methanol/water, 130/70/1 (v/v/v).

Fatty acid methyl esters (FAME) by silver nitrate (argentation) TLC

In argentation TLC, silver ions form polar complexes reversibly with double bonds. This property is used to isolate fatty acid esters or lipids based on number, geometric isomer (cis or trans) and position of double bonds. In the Ohlrogge/Pollard research group, argentation TLC is often used for separation of positional isomers of monoenoic fatty acids. However, this technique can be used to separate many types of lipids based upon the number, type and position of double bonds. Silver concentrations above 10% are typically only used for monoene separation. Lower silver concentrations are used for most other applications. For more in depth information see Morris (1966, J. Lipid Research 7:717-732) and Gunstone et al. (1967, Chem. Phys. Lipids 1:376-385).

Monoene separation. Prepare solution of AgNO₃ (15% in acetonitrile, w/v). Solution can be reused if stored properly: dark container in a dark cabinet. Dip plate (K6) for 10 minutes (keep container covered while dipping to prevent evaporation of solvent). Briefly dry plates in the hood, then store overnight in a dark cabinet to ensure complete drying. Set up covered TLC tank containing fresh toluene in a –20°C freezer the night before running your samples. Spot samples in a small line. Develop the plate 3 times (1/3, 2/3, and full) drying completely between developments.

Detection of lipids on TLC plates

Direct Visualization Under UV Light . If the silica plates used for TLC have been impregnated with fluorescence indicator and you are running derivatives that quench fluorescence, such as phenacyl esters, then when viewing under UV light your compounds will appear as dark spots in the yellow-green background.

Staining with iodine vapor . Iodine complexes with any unsaturated lipids and fatty acids through interactions with double bonds; therefore, it will not detect saturated fatty acids.. Put plate into iodine chamber wait for band to become visible, remove plate and enclose in Saran wrap and make a photocopy. The longer the plate is in the chamber the lower the detection limit; with careful analysis you can detect down to 1-2 μg. The iodine will sublime over time once the plate is removed and the bands will disappear. Iodine staining often reveals very faint bands that can disappear rapidly. This method of lipid detection is potentially destructive to the lipids, often causing cis-trans isomer scrambling as well as double bond oxidation, and should not generally be used for preparative work. It is also possible to stain only part of the TLC plate with iodine. A lane of standards or sample can be exposed to iodine by putting iodine crystals into a dispo pipette with glass wool. Then blow a N₂ stream through the pipette only onto the lane to be stained while covering the other parts of TLC plate with glass. Iodine vapors are toxic: do not breath

Phosphomolybdic acid . Phosphomolybdic acid is a general, destructive charring detection method. Spray plate with a 1 % phosphomolybdic acid solution in ethanol, then heat in an oven at 80-100 °C. Lipids should appear as blue or gray spots on a yellow background in a few minutes. Saturated lipids require a more prolonged intense heating.

Dichlorofluorescein . A non-specific reagent that renders all lipids visible under UV light, dichlorofluorescein requires spraying of a 0.1% (w/v) solution in 95% methanol (against the spray board in the back of the fume hood). Allow the plate to dry and the lipids will appear as yellow spots under UV light. Aerosol spray and solution are found under the fume hood in the top shelf. This is not a sensitive lipid detection method but it is useful for nondestructive location of lipid bands in preparative TLC.

α-Naphthol for glycolipids . Spray with 2.4% (w/v) α-Napthol in 10% H₂SO₄, 80% EtOH (v/v). Heat to 120 C in oven. Glycolipids appear as purple-blue spots; other lipids as yellow spots.

Radioactivity . See section on Instant Imager. Radioactive ink can be used on preparative TLC plates as marker spots to locate bands for isolation.

E. SIMPLIFIED ARABIDOPSIS TRANSFORMATION PROTOCOL

http://plantpath.wisc.edu/~afb/protocol.html

(Brief version for those who are familiar with the method)

Steve Clough and Andrew Bent, University of Illinois at Urbana-Champaign.

Our present protocol (Clough and Bent, 1998; modified from Bechtold et al. 1993) is extremely simple. We have found that the MS salts, hormone, etc. make no difference, that OD of bacteria doesn't make much of a difference, that vacuum doesn't even make much of a difference as long as you have a decent amount of surfactant present. Plant health is still a major factor - healthy fecund plants make a big difference! With this method you should be able to achieve transformation rates above 1% (one transformant for every 100 seed harvested from Agrobacterium-treated plants).

1. Grow healthy Arabidopsis plants until they are flowering. Grow under long days in pots in soil covered with bridal veil, window screen or cheesecloth.

2. (optional) Clip first bolts to encourage proliferation of many secondary bolts. Plants will be ready roughly 4-6 days after clipping. Clipping can be repeated to delay plants. Optimal plants have many immature flower clusters and not many fertilized siliques, although a range of plant stages can be successfully transformed.

3. Prepare Agrobacterium tumefaciens strain carrying gene of interest on a binary vector. Grow a large liquid culture @ 28°C in LB with antibiotics to select for the binary plasmid, or grow in other media. You can use mid-log cells or a recently stationary culture.

4. Spin down Agrobacterium, resuspend to OD600 = 0.8 (can be higher or lower) in 5% Sucrose solution (if made fresh, no need to autoclave). You will need 100-200 ml for each two or three small pots to be dipped, or 400-500 ml for each two or three 3.5" (9cm) pots.

5. Before dipping, add Silwet L-77 to a concentration of 0.05% (500 ul/L) and mix well. If there are problems with L-77 toxicity, use 0.02% or as low as 0.005%.

6. Dip above-ground parts of plant in Agrobacterium solution for 2 to 3 seconds, with gentle agitation. You should then see a film of liquid coating plant. Some investigators dip inflorescence only, while others also dip rosette to hit the shorter axillary inflorescences.

7. Place dipped plants under a dome or cover for 16 to 24 hours to maintain high humidity (plants can be laid on their sides if necessary). Do not expose to excessive sunlight (air under dome can get hot).

8. Water and grow plants normally, tying up loose bolts with wax paper, tape, stakes, twist-ties, or other means. Stop watering as seeds become mature.

9. Harvest dry seed. Transformants are usually all independent, but are guaranteed to be independent if they come off of separate plants.

10.Select for transformants using antibiotic or herbicide selectable marker. For example, vapor-phase sterilize and plate 40 mg = 2000 seed (resuspended in 4 ml 0.1% agarose) on 0.5X MS/0.8% tissue culture Agar plates with 50 ug/ml Kanamycin, cold treat for 2 days, and grow under continuous light (50-100 microEinsteins) for 7-10 days.

11.Transplant putative transformants to soil. Grow, test, and use!

Note

For higher rates of transformation, plants may be dipped two or three times at seven day intervals. We suggest one dip two days after clipping, and a second dip one week later. Do not dip less than 6 days apart.

References:

Bechtold, N., Ellis, J., and Pelletier, G. (1993). In planta Agrobacterium-mediated gene transfer by infiltration of adult Arabidopsis thaliana plants. C. R. Acad. Sci. Paris, Life Sciences 316:1194-1199.

Clough SJ and Bent AF, 1998. Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J 16:735-43.

Additional commentary can be found by searching the Arabidopsis newsgroup archives:

http://genome-www.stanford.edu/cgi-bin/biosci_arabidopsis

Note: Obtain proper approval for transformation work from institutional authorities. Autoclave and properly dispose of all materials.

F. TRANSFORMATION OF ARABIDOPSIS BY VACUUM FILTRATION (Green Lab protocol)

http://www.bch.msu.edu/pamgreen/vac.htm a step-by-step picture demonstration is available online.

The Green Lab protocol is adapted from protocols by Nicole Bechtold (Bechtold et al., 1993), Andrew Bent (Bent et al., 1994) and Takashi Araki. No claims are made that any of the steps are necessary or ideal; these experiments have not been done. However, this protocol gives us very good results, with at least 95% of all infiltrated plants giving rise to transformants, and a transformant rate of 1-4% of seed.

1. Sow seeds of ecotype Columbia in lightweight plastic pots prepared in the following way: mound Arabidopsis soil mixture into pots (We use 3 1/2 inch to 4 inch square pots), saturate soil with Arabidopsis fertilizer, add more soil so that it is rounded about 0.5 to 1 inch above the top, dust with fine vermiculite, cover soil with a square of window screen mesh (Circle Glass Co., Detroit, MI) and secure mesh with a rubber band.

2. Grow plants under conditions of 16 hours light/ 8 hours dark at 20 °C to 22 °C, fertilizing from below with Arabidopsis fertilizer once a week , adding approximately 0.5" to each flat. Thin the plants to one per square inch or fewer per pot. After 4-6 weeks, depending on your conditions, plants will be ready to infiltrate when they are at this stage: the primary inflorescence is 5-15 cm tall and the secondary inflorescences are appearing at the rosette. No clipping of bolts is necessary before infiltration.

3. In the meantime, transform your construct into Agrobacterium tumefaciens strain GV3101 (C58C1 Rifr) pMP90 (Gmr) (Koncz and Schell, 1986). When plants are ready to transform, inoculate a 500 ml culture of YEP medium containing 50 mg/l rifampicin, 25 mg/l gentamycin and the appropriate antibiotic for your construct with a 1 ml overnight starter culture. Be sure to water your plants well the day before infiltration so that the stomata will be open that day.

4. Grow culture overnight at 28 °C with shaking, until culture OD600 is > 2.0. Spin down the culture and resuspend it in 1 liter of infiltration medium.

Infiltration medium (l liter)

2.2 g MS salts

1X B5 vitamins

50 g sucrose

0.5 g MES

pH to 5.7 with KOH

0.044 μM benzylaminopurine (be sure the final concentration is micro molar). 200 μl Silwet L-77 ( OSi Specialties request that purchases be made at Lehle Seeds, fax # (512) 388-3974 catalog # vis-01)

5. Place resuspended culture in a Rubbermaid container inside a vacuum desiccator. Invert pots containing plants to be infiltrated into the solution so that the entire plant is covered, including rosette, but not too much of the soil is submerged. One good way to do this is to place the corners of the pots on rubber stoppers sitting in the culture. Make sure no large bubbles are trapped under the plant.

6. Draw a vacuum of 400 mm Hg (about 17 inches). Once this level has been obtained, close the suction (i.e., so that the vacuum chamber is still under 17 inches of mercury but the vacuum is not still being directly pulled) and let the plants stay under vacuum for five minutes. Quickly release the vacuum. Briefly drain the pots, place them on their sides in a tray, cover the tray with plastic wrap to maintain humidity, and place the flats back in a growth chamber. The next day, uncover the pots and set them upright. Keep plants infiltrated with different constructs in separate trays from this stage on.

7. Allow plants to grow under the same conditions as before (see step 2). Stake plants individually as the bolts grow. The leaves that were infiltrated will degenerate, but plants continue growing until they finish flowering. Gradually reduce water and then stop watering to let them dry out. Harvest seeds from each plant individually.

8. Prepare large selection plates:

4.3 g/l MS salts

1X B5 vitamins (optional)

1% sucrose

0.5 g/l MES

pH to 5.7 with KOH

0.8% phytagar

Autoclave. Add antibiotics (30 g/ml works well for kanamycin). Pour into 150 X 15 mm plates. We also add vancomycin at 500mg/l to control bacterial growth.

9. Dry plates well in the sterile hood before plating. Twenty minutes to half an hour with the lids open is usually sufficient.

10. For each plant sterilize up to 100 μl of seeds (approximately 2500 seeds) and plate out individually. Sterilize seeds (7 minutes rocking in 50% bleach/0.02% Triton X-100, 3 rinses in sterile distilled water). Resuspend seeds in approximately 8 ml sterile 0.1% agarose and pour onto large selection plates as if plating phage. Tilt plate so seeds are evenly distributed, and let sit 10-15 minutes. After a while the liquid should soak into the medium; if evaporation is too slow, open the plate in the hood and let dry until the excess liquid is gone. Seal plates with Parafilm or paper surgical tape and place in a growth room.

11. After 5 to 7 days, transformants will be visible as green plants. Transfer these onto "hard selection" plates (100 x 20 mm plates with same recipe as selection plates but with 1.5% phytagar) this allows roots to elongate and eliminates any false positives. Place in growth room.

12. After 6-10 days, plants will have at least one set of true leaves. Transfer normal conditions. Keep covered for several days. Note: We usually move just one transformant to soil from any one plant that was infiltrated, to ensure independent transformants.

References

Bechtold N, Ellis J, Pelletier G (1993) C. R. Acad. Sci. Paris 316:1194-1199

Bent A, Kunkel BN, Dahlbeck D, Brown KL, Schmidt R, Giraudat J, Leung J,

Staskawicz BJ (1994) Science 265:1856-1860

Koncz C, Schell J (1986) Mol. Gen. Genet. 204:383-396

Solutions

YEP medium (1 liter)

10 g Bacto peptone

10 g yeast extract

5 g NaCl

1000X B5 vitamins (10 ml)

1000 mg myo-inositol

100 mg thiamine-HCl

10 mg nicotinic acid

10 mg pyridoxine-HCl

Dissolve in ddH₂O and store at -20C.

Arabidopsis fertilizer (20 liters)

100 ml 1M KNO₃ (5ml/L)

50 ml 1M KPO₄ (pH 5.5) (2.5ml/L)

40 ml 1M MgSO₄ (2ml/L)

40 ml 1M Ca(NO₃)₂ (2ml/L)

10 ml 0.1M Fe.EDTA (.5ml/L)

20 ml micronutrients (see below) (1ml/L)

Dissolve in H₂O and store at room temperature

Arabidopsis micronutrients (500 ml)

70 ml 0.5M boric acid

14 ml 0.5M MnCl₂

2.5 ml 1M CuSO₄

1 ml 0.5M ZnSO₄

1 ml 0.1M NaMoO₄

1 ml 5M NaCl

0.05 ml 0.1M CoCl₂

Dissolve in ddH₂O and store at room temperature

Commonly asked questions about this vacuum infiltration method

Q. Why add vancomycin to the seed selection medium?

A. We find it decreases Agrobacterium growth that may occur as the seedlings germinate. Although the seed coat is surface sterilized, in our experience some seeds may contain Agrobacterium inside the seed coat. We have found an inexpensive source of vancomycin at our university pharmacy. You may want to check at your pharmacy, indicating it is for research purposes only, and ask for it in injectable vials.

Q. Why thin plants to just a few per pot?

A. We found that it increased the transformation rate of primary plants to at least 95%. It also increases the number of seeds generated per plant and it facilitates harvesting

individual plants. We harvest plants individually and choose one transformant per primary plate to assure that transformants are independent.

Q. If the leaves die shortly after infiltration, is something wrong?

A. No, we always see rapid degeneration of leaves after infiltration. Keep watering and

growing plants until they finish flowering.

Q. What method does the Green lab use to transform Agrobacterium tumefaciens ?

A. We prefer electroporation. Refer to the manual of your electroporation unit for method.

G. DEVELOPMENT OF WESTERN (PROTEIN) BLOTS

Blot development reagents, buffers and stock solutions are kept in room 366, on the shelf next to the filing cabinet or in the refrigerator door compartment (fridge/freezer G).

Replenish stocks as needed, especially if you have been using them often. Recipes for buffers and stocks can be found at the end of the protocol.

Blot preparation/fixing

· Rinse blot briefly to remove bits of gel. (If blot is dry, wet in distilled water for nitrocellulose, methanol for PVDF.)

· Fix blot by incubating in 5% formaldehyde 5 min followed by several rinses in distilled water.

- or -

· Stain blot with reversible stain, Ponceau Red S, several minutes, rinse briefly in distilled water to destain and visualize bands, then destain completely with several washes in TBST. (It is not necessary to fix blots that have been stained)

Suggested incubation volumes: 25 ml for large (11x14 cm) gel, 10 ml for mini (8x5 cm) gel.

Blot development

1. Blocking: 1 hr in TBST + 1 mg/ml BSA or 5% nonfat dry milk in TBST (aka “Blotto”)

2. Primary antibody: Dilute in TBST, incubate 1-2 hrs.

Proper dilutions can range from 1:250 to 1:5000 so be sure to check for each different antibody used. Primary antibody dilutions should be saved and reused several times. To conserve stocks of valuable antibodies, keep using until you notice loss of sensitivity. To freeze, add glycerol to 5%, or to keep at 4^oC, add antimicrobial agent such as Na-azide.

3. Wash 2X 5 min in TBST.

4. Secondary antibody: Dilute secondary antibody, KPL alkaline phosphatase goat anti-rabbit, 1:2000 in TBST, incubate 1 hr.

5. Wash 2X 5 min in TBST, pour off excess buffer.

6. Rinse 1 X with alkaline phosphatase buffer.

7. Development. Just before use, dilute BCIP and NBT stock solutions, together, 10 ul/ml in alkaline phosphatase buffer. Incubate blot in this solution to desired band intensity but before purple background is too high. This may take less than 1 min or as long as 15 min.

8. Stop development with several rinses of distilled water. Store developed blots dry, away from light.

Buffer and stock solutions

Ponceau Red S staining solution:

0.1% Ponceau Red S

1% acetic acid

1X TBST:

10 mM Tris/HCl, pH 8.0

0.15 M NaCl

0.3% Tween 20

(make 10X stock)

BCIP (5-bromo-4-chloro-3-indoyl phosphate, aka X-Phos-p-tol) stock solution:

17 mg/ml in DMSO

NBT (p-Nitro blue tetrazolium chloride) stock solution:

33 mg/ml in 70% DMSO, 30% water

Alkaline phosphatase buffer:

100 mM Tris/HCl, pH 9.5

100 mM NaCl

5 mM MgCl

H. PROTEIN ASSAY

General Principles

A simple protein assay method used in most labs is based upon the dye-binding assay of Bradford, 1976, Anal. Biochem 72:248-254. Binding of Coomassie Brilliant Blue G-250 dye by basic and aromatic amino acids results in a shift in absorbance from 465 nm to 595 nm. The extinction coefficient of a dye-albumin complex solution is constant and linear over a 10-fold concentration range. The relative concentration of a complex or purified protein solution can be determined by constructing a standard curve using purified protein such as bovine serum albumin (BSA) or thyroglobulin for comparison.

Method

Protein standard solution preparation

Accurately prepare a 1 mg ml^-1 bovine serum albumin (or thyroglobulin) in a 0.05% sodium azide working solution. Note: NaN₃ (sodium azide) is functionally equivalent to cyanide, a potent inhibitor of cytochrome oxidase, so be careful when using this reagent. When stored with azide preservative, this working protein standard solution can be used for months.

Protein dye-reagent preparation

For a working solution, dilute one part Bio-Rad protein assay dye reagent concentrate (kept at 4° C; catalog number 500-0006) with four parts deionized water. Remove particulate matter by filtering through a Whatman #1 filter or by centrifugation (5 min, 5 K g).

Protein quantitation

1. Add one ml of protein dye working solution to clean glass test tubes.

2. To dye reagent add 20 ul of standards or samples. Mix by vortexing and keep at room temperature for at least five minutes but no more than one hour.

3. Measure absorbance at 595 nm.

4. Create standard curve using graph paper or calculate the line equation using a calculator or computer software. Determine the concentration of unknown sample(s) using the line equation or interpolation from a standard curve

Additional notes:

· Standard curves should contain at least five points (including blank) and must be generated each time the assay is performed.

· The absorbance for protein samples must be within the range of the standard curve.

· Detergents at concentrations greater than 0.1% should be avoided since they interfere with protein dye-binding. For a complete list of compatible reagents consult Bio-Rad manual available online at www.biorad.com

· Sample quantitation is usually performed in triplicate.

I. ACP ASSAY USING ACYL-ACP SYNTHETASE REACTION

Make chart for each tube of synthetase reaction. 15 μl and 10 ug maximum of ACP. 50 μl total for each reaction. Make up difference with water.

	Tube	ACP (ul)	H20 (ul)	Rx Mix (ul)	Do ‘0 ACP’ control Use a known ACP for standard
0.5 ml microfuge tubes	1	0	15	35
	2	1	14	
	3	3	12	
	4	10	5	
	5	15	0	.

Prepare Rx mix from stock solutions as show below.

		per 50 μl rxn	per 40 μl	per 25 μl
	Tris/Mg/Li 4X buffer	12.5	10	6.25
	*16:0 (3H or 14C)	10.0	8	5
Keep on ice! Esp. enzyme	ATP (100mM)	2.5	2	1.25
	DTT (100mM)	1.0	0.8	0.5
	H2O	5.0	4	--
	Enz (E. coli) synthetase))	4.0	3.2	2
	Total volume (ul):	35.0	28	15

1. Add in order: H₂O, ACP Rx Mix, and finally enzyme; cap tightly.

2. Mix gently.

3. Incubate at 37°C for 1 hour.

4. Mix gently.

5. Add 25 μl from each tube onto numbered (w/pencil) ½ DE-81 filter papers (25 mm circles).

6. Dry for a few minutes - not overnight.

7. Wash 4X for 2-3 min each with 80% isopropanol and 20% 10mM KPO₄ pH 6.0 + 0.1M NaCl.

8. If using 3H-16:0, put filters into scintillation vials with1 ml of 0.1N NaOH and heat for 15 min at 65°C. Cool. This hydrolyzes acyl-ACP and releases 3H into scintillation fluid for more efficient counting.

9. Add 10 ml of scintillation fluid and count. If using 14C-16:0, omit step 8. Add filters to vials + 10 ml scintillation fluid and count.

10. When counted, use the dpm’s from the 0-ACP control as background and subtract it from your data.

Notes

· Avoid freeze thaw cycles for enzyme.

· Store working aliquots at 4°C.

Preparation of fatty acids for acyl-ACP synthetase reaction

1. Transfer organic solution of radioactive FA to glass vial or tube.

2. Add 5-10 μl NH₄OH. This creates the NH₄-salt which is more soluble in aqueous solutions.

3. Evaporate under N₂ in hood - watch carefully - do not over evaporate.

4. Redissolve in 10% purified Triton-X-100 (Pierce). Regular Triton X-100 contains peroxides.

5. Warm solution to 50-60°Cto help FA dissolve. (Triton may become cloudy when warm).

6. Check to make sure FA is in solution by counting 2 + 4 μl aliquots.

7. Store at -20°C.

Acyl-ACP Synthetase Stock Solutions

100 mM ATP

Dissolve 61.5 mg Sigma #A6144 in 850 μl H₂O.

Add 110 μl 1 N NaOH.

Adjust pH to 7 with 1 N NaOH using pH paper (6-8 range).

Store at -20°C, stable for several months

100 mM DTT

Dissolve 15.6 mg DTT in 1 ml H₂0.

(DTT is unstable above pH 7.)

Store at -20°C, stable for 1-2 months

4X Buffer - pH 8.0

0.4 M Tris 4.8 g

40 mM MgCl₂6H₂O 0.81 g

1.6 M LiCl₂ 6.7 g

100.00 ml

Adjust pH to 8.0, adjust vol. to 100 ml, store at 4°C

J. PROTOCOL FOR ISOLATING CHLOROPLASTS FROM SPINACH, PEA AND AMARANTHUS (Grattan Roughan, author)

Another reference for chloroplast isolation, also has many other chloroplast biochemical procedures: Perry, S.E. et al. 1991. In Vitro Reconstitution of Protein Transport into Chloroplasts. Methods in Cell Biology. 34:327-344.

(With no apology to those so shamelessly plagiarized - for references see Meth. Enzymol. 148, 327-337)

Growing appropriate plants

Highest biosynthetic activities are measured in isolated chloroplasts isolated from plants which have not recently experienced water stress. Therefore, it is worthwhile precaution to invest in a system for growing spinach (and other plants) in water culture. Also, for consistent biosynthetic activities, and to prevent spinach plants from flowering it is advisable to grow the plants in constant environment i.e. in a growth chamber. A day length of 10 h, day temperature of 25^oC and night temperature of 20^oC seems to work very well. It now appears likely that preparation of the most highly-active chloroplasts requires that leaves be harvested from plants grown from relatively fresh seed. Even seed that has been stored at 2-4^oC will not produce suitable plants after 3-5 years. Freshness of the seed is probably more important than the cultivar used for the preparation of chloroplasts, although hybrids may have some advantages. Sow 2 g of fresh spinach seeds into a 8" pot of vermiculite and keep well watered, with nutrient solution when the seedlings begin to show their first true leaves,. After 12-14 days, transfer the seedlings to water culture taking great care to avoid damaging the hypocotyls in particular. Gently wedge the seedling in place with wads of foam rubber. Place two seedlings in half of the holes to use as backups for replacing the plants that do not make it. Be very conscious that the roots need to be in the nutrient solution! Beware of gawkers who will lift the lids to look at the roots and then leave roots of seedlings dangling over the side of the container! Between 2 and 4 weeks following this transfer the plants are producing expanding leaves from which the most active chloroplasts may be isolated. Therefore, to ensure a continuing supply of the very best plants, fresh seed should be sown every 2-3 weeks. The ideal is to have five containers of plants in water culture representing seedlings transferred 1 and 5 weeks previously. These are then rotated so that the oldest plants are discarded, and a fresh lot of seedling is transferred to water culture each week. In this case, containers 3 and 4 provide the bulk of the leaves for chloroplast isolation. The nutrient solution is completely replaced each week and the containers are thoroughly washed out at the same time. Apply sensible hygiene procedures. We have grown spinach, Solanum nigrum, safflower, Chenopodium alba, and amaranthus in this system.

Pea seedlings may be raised with rather less trouble. Evenly space 36 fresh pea seeds onto wet pumice/peat potting mix (soil) in each of three 8" pots. Cover with about 1" potting mix, transfer to growth chamber and water daily, particularly as the seedlings are emerging. After 6-8 days, depending on temperature and daytime length but before any of the leaves are fully expanded, harvest all the shoots.

ISOLATION BUFFERS

For SPINACH

Homogenizing and wash buffer: 9 g. Sorbitol

1.2 ml 0.25 M HEPES-KOH, pH 7.8

60 ul 1M KCl

60 ul 0.1M EDTA

To 150 ml with water

Transfer 100 ml to homogenizing vessel - homogenizing buffer

and 50 ml to screw-capped flash - wash/resuspend buffer.

Note that if there is any doubt about the quality of the spinach plants, or if the spinach are growing indifferently, then use the buffers and follow the procedure as outlined below for peas.

For PEA

Homogenizing buffer: 6.4 g. Sorbitol

0.8 ml 0.25 M HEPES-KOH, pH 7.8

40 ul 1M KCl

40 ul 0.1M EDTA

100 mg BSA (optional)

To 100 ml with water.

X2 Wash/resuspend buffer: 0.4 ml 0.25 M HEPES-KOH, pH 7.8

3 g. Sorbitol

20 ul 1M KCl

20 ul 0.1M EDTA

To 25 ml with water

40% Percoll cushion: 6 ml X2 buffer

6 ml 80% Percoll

Wash/resuspend; dilute the remaining X2 buffer (19 ml) with an equal vol. of water.

For AMARANTHUS

Homogenizing buffer: 6.4 g. Sorbitol

10 ml 0.25 M HEPES-KOH, pH 7.8

2.5 ml 0.2M EDTA

1 ml 0.1M MgCl₂

200 mg BSA

To 100 ml with water.

Immediately before freezing add 70 mg DTT and 40 mg iso-ascorbate.

X2 Wash/resuspend buffer: 6 g. Sorbitol

10 ml 0.25 M HEPES-KOH, pH 7.8

2.5 ml 0.2M EDTA

100 mg BSA

To 50 ml with water

35% Percoll cushion: 6 ml X2 buffer

6 ml 70% Percoll

Wash/resuspend buffer: 10 ml X2 wash buffer

10 ml water

ISOLATION PROCEDURES

Select clear, 50 ml centrifuge tubes (polycarbonate) that are free from scratches and other internal imperfections. Do not wash these tubes in detergent (unless you do it yourself).

Spinach

Slice 9-10 expanding leaves into 100 ml semi-frozen buffer and homogenize with 2-3 bursts of 3-5 s from Polytron. Filter through 2 layers of wetted Miracloth. Distribute filtrate between 2 x 50 ml centrifuge tubes. Spin 4000 rpm in HB-4 rotor for 30 s, then stop by hand. Dump the supernatant by quickly upending the tubes, and allow any sloppy material to escape. Be firm! Rinse the surfaces of each pellet with 5 ml wash buffer before resuspending in 2.5 ml wash buffer, using a 5 ml pipet. Dilute each chloroplast suspension with a further 15 ml of wash buffer. Re-centrifuge 4000 rpm for 30 s. Hand brake. Pour off the supernatants and, keeping the tubes inverted the whole time, use a tissue held in forceps to wipe excess buffer from sides of tubes. Resuspend and combine final pellets in 1 ml wash buffer, then measure volume of chloroplasts using a 2 ml pipet with controller. Add more buffer if necessary so that the pellet is diluted with 9 vol of buffer.

Pea

Transfer 100, 7 to 8-day shoots (about 15g tissue) to semi-frozen buffer and chop with scissors. Homogenize using Polytron and transfer filtrate to centrifuge tubes as above. Layer 5.5 ml 40% Percoll from a syringe under each portion and spin at 4000 rpm in HB-4 rotor for 60 s. Hand brake. Decant supernatant and, keeping the tubes inverted, wipe around insides of tubes below the pellets with a tissue held in forceps. Resuspend each pellet, and dilute suspensions as for spinach, but using a total of 30 ml wash buffer. Centrifuge at 4000 rpm for 30 s. Hand brake. Resuspend final pellets in 1 ml wash buffer as above.

A fairly rapid Percoll density gradient separation of intact from broken chloroplasts may be performed as an alternative to the Percoll cushion procedure. Add 13 ml of wash buffer to one side of the gradient former and 13 ml of Percoll, containing 0.33 M sorbitol, 2 mM Hepes/0.4mM KCl, 0.04 mM EDTA, to the other side. Form the gradient as usual. Chloroplasts (0.5-1 mg chl) prepared as for spinach (above) are resuspended in about 5 ml of wash buffer and layered over the gradient. Centrifuge in HB-4 rotor for 1 min at 13,000xg (9000 rpm) and allow the rotor to stop without the brake. Two well-separated green bands represent broken chloroplasts and thylakoids (upper) and intact plastids (lower). Aspirate the solution from above the intact plastids and recover same. Dilute X4 with wash buffer and centrifuge at 4000 rpm for 30 s. Treat the pellet as above.

Amaranthus

Slice 50 expanding leaves into 100 ml semi-frozen buffer, homogenize and filter as above. Centrifuge filtrate in 2 X 50 ml tubes at 6000 rpm for 30 s. Resuspend each pellet as above but using a total of 18 ml only of wash buffer, and underlay with 5.5 ml of 35% Percoll. Centrifuge at 2000 rpm for 3.5 min. Decant supernatants and use a tissue in forceps to wipe away any broken chloroplasts on sides of tubes above the pellets. Resuspend pellets in 1 ml of wash buffer as above. The starch content of these pellets renders then more difficult to suspend then those from spinach and peas.

The following protocols are for chloroplast incubations to determine rates of fatty acid synthesis and analysis of products of fatty acid synthesis from chloroplasts.

INCUBATION BUFFERS - Stock solutions.

For spinach and pea: 1.2 g sorbitol

2.5 ml 0.25M HEPES/NaOH pH 8.0

0.5 ml 0.5M KHCO₃

0.5 ml 0.1M EDTA

0.25 ml 0.1M MgCl₂

0.25 ml 0.1M MnCl₂

0.25 ml 50 mM K₂HPO₄

To 10 ml with water.

Use 100 ul for a final assay volume of 250 ul.

For amaranthus: 1.2 g sorbitol

2.5 ml 0.25M HEPES/NaOH pH 8.0

0.5 ml 0.5M KHCO₃

To 10 ml with water.

Use 100 ul for a final assay volume of 250 ul.

ASSAYS

Basal medium - 100 ul stock solution, 10 ul [1-¹⁴C] acetate (5 mM, 125 nmole/uCi), 90 ul water and 50 ul chloroplast suspension.

The [1-¹⁴C] acetate supplied by Amersham will be 1 mCi in 5 ml and about 58 Ci/mole. I call this 17.2 nmole/uCi and 3.45 mM. Right? For assays where routine lipid analyses only are to be performed this specific radioactivity should be diluted to 125 nmole/uCi by the judicious addition of cold sodium acetate. However, where analyses of very minor constituents, e.g. acetyl-CoA, acyl-ACPS is contemplated, or where very brief time-courses are to be analysed, the [1-¹⁴C] acetate should be used undilute.

Assays are conveniently carried out in 20 x 125 culture tubes (Kimax 45066-A) for up to 0.5 ml reactions. Greater volumes (1-2 ml) should be incubated in 25 ml Erlenmeyers to expose a greater surface area to the incident light. A shaking, illuminated, constant-temperature water bath is essential - this can be adapted from a “photosynthetic Warburg apparatus.” Acetate incorporation is usually linear for at least 15 min but look for a brief lag initially. At very high rates of FAS the substrate concentration may become limiting in less than 15 min.

Additions to the basal medium- 10 ul 0.2% (w/v) Triton X-100

10 ul 10 mM sn-G3P

10 ul 12.5 mM CoA

10 ul 25 mM ATP

10 ul 2.5 mM UDP-gal

10 ul 25 mM CTP

etc. In place of an equivalent amount of water.

Start reactions by adding chloroplasts in the light and stop at suitable times by adding 2.5 ml chloroform/methanol (1:1, v/v). Shake with 0.9 ml of 0.3 M H₃PO₄ in 1M KCl and centrifuge. Recover lower phase using a syringe and re-extract aqueous layer with 2 ml pet ether. Remove solvent under N₂ and dissolve residue in 0.5 ml of chloroform. When reactions have contained both CoA and ATP, add 0.2 ml of 40% (w/v) KOH to remaining aqueous phases and heat to 80^oC for 30 min to hydroyse acyl-CoA. Cool, acidify with 1 ml 1.8 M H₂SO₄, add 25 ug carrier fatty acid (ex. Olive oil) and extract fatty acids into 3 + 2 ml pet ether. Remove solvent under N₂.

Streak 50 ul of the lipid extract and 25 ul of DAG/UFA (about 10 ug of each) reference across 1 cm in 2 cm lanes on thin layers of 5% (w/w) boric acid in silica gel and develope chromatograms with 4% acetone in chloroform. Stain in I₂ vapor to locate 1,2-DAG, UFA and polar lipids (origin). 1,2-DAG is the fastest band (behind chl) followed by UFA, and polar lipids stay on the origin. Scrape bands into scintillation vials and count. Fatty acids recovered from the aqueous methanol phase represent acyl-CoA and are applied as 1 cm streaks in 2 cm lanes on thin layers of silica gel. Chromatograms developed to 10 cm with pet ether/diethyl ether/HAc (70:30:1). Fatty acids are located with I₂ vapor, and the sniffer, and are scraped into vials for scintillation counting.

For analysis of acyl-CoA and acyl-ACP

Reactions (0.5 ml) are stopped with 0.5 ml of 5% TCA (it should be 10% TCA but SHE won’t allow it). 50 ul of the resultant slurry is transferred to 2 ml of CHCl₃/MeOH and partitioned against 0.9 ml water. Recover the lower layer as above for lipid analysis. 900 ul of the remaining slurry is transferred to ufuge tube and spun to the max. 850 ul of the supernatant is recovered, and the precipitate is handed over to herself. The supernatant is extracted x4 with 1 ml diethyl ether to remove TCA, then dried in the Speedvac. Add 200 ul of 0.1M KH₂PO₄ to the residue and store at -20^oC.

HPLC analysis

Stock buffer; Dissolve 68 g KH₂PO₄ in 900 ml water and add 1.5 ml of the 10M KOH on the shelf. This will give 0.5 ml KH₂PO₄, pH 5, when diluted to 1000 ml. Store at 0-4^oC.

Buffer A: Dilute 100 ml of stock to 500 ml with water.

Buffer B: Dilute 100 ml of stock to 300 ml with water and mix in 200 ml acetonitrile. Add more water to 500 ml.

Equilibrate the HPLC column with 97% A, 3% B. Ensure that both pumps are delivering the correct volumes. CoA reference compounds are 1-2 mg/ml in 0.001M HCl (pH 3). Dilute these x50 with 0.1M phosphate, pH 5, for about 25 uM. Confirm concentrations spectrophotometrically using a millimolar extinction coefficient of 15.4 at 260 nm. Program the fraction collector to recover 1 ml fractions from 25 to 45 ml after injection. Load 150 ul of the sample and initiate program #4. Use a Pipetman to transfer the fractions to scintillation vials, and then rinse the collecting tubes with 2 x 2.5 ml of scintillation cocktail.

For CoA compounds in isolated chloroplasts, the plastids should be immediately resuspended in water, sampled for [chl], and brought to 5% with TCA. It is probably best to divide the suspension into equal parts for this. Centrifuge, and recover a measured amount of supernatant so that equivalent chl contents may be calculated. Extract x5 with equal volumes of diethyl ether to remove TCA, transfer to 12 x 75 mm disposable culture tubes and dry in Seedvac. Dissolve residue in KH₂PO₄ so that 100 ul corresponds to 100 ug chl, and transfer to ufuge tubes. Spin solution at top speed before injecting 100 ul. To confirm the identification of CoA esters, treat 150 ul with 25 ul of conc ammonium hydroxide for 10 min at room temp. Dry in Speedvac and dissolve residue in 150 ul water. Centrifuge before injecting 100 ul. Alkali-labile CoA esters will disappear from the trace and be replaced by an equivalent amount of GSH-CoA.

Chlorophyll

In duplicate; 50 ul chloroplast suspension into 0.95 ml water. Add 4 ml acetone. Centrifuge and measure absorbance against 80% acetone at 663, 652 and 645 nm.

[A(663] x 8.02 + A(645) x 20.2] = ug chl/10 ul of suspension.

Or A(652) x 29 = ug chl/10 ul suspension

ADDITIONAL INFORMATION

Nutrient mix for growing plants in water culture (“modified Hoagland’s solutions”)

Stock solution A Ca(NO₃)₂ ^. 4H₂O 295 g/L

Sequestrene 10 g/L

Stock solution B KH₂PO₄ 34 g/L

KNO₃ 126 g/L

MgSO₄ ^. 7H₂O 123 g/L

Boric acid 715 mg/L (23ml of .5M stock)

MnCL ₂ ^. 4H₂O 452 mg/L (4.6ml of .5M stock)

ZnSO₄ ^. 7H₂O 55 mg/L (382 ul of .5M stock)

CuSO₄ ^. 5H₂O 20 mg/L (80 ul of 1M stock)

NaMoO₄ or H₂MoO₄ ^. H₂O 7 or 5 mg/L (290 ul of .1M stock)

KCl or NaCl 1-2 g/L

Stock solutions found with the Arabidopsis nutrient ingredients in the cabinet 104 in room 362

Use 25 ml of each per tub of about 10 liters. Ensure adequate aeration of the solutions.

We have grown Spinacia oleracea, Chenopodium alba, Solanum nigrum and Amaranthus lividus successfully in this mix.

K. USE OF PAT GENE AS SELECTABLE MARKER FOR PLANT TRANSFORMATIONS

A major advantage of the PAT selectable marker is that selection can be done on soil and does not require asceptic growth of plants on agar plates as with kanamycin, etc. However, note that kan has also been used as a spray to select soil grown plants (Plant Mol Biol Reporter).

The herbicide glufosinate ( phosphinothricin or PPT ) inhibits Gln synthetase (GS), a key enzyme in the assimilation of inorganic nitrogen into organic compounds. Inhibition of GS by L-glufosinate, the active isomer, leads to depletion of the amino acid Gln, a concomitant accumulation of ammonia in treated tissues, and glyoxylate accumulation, which inhibits Rubisco and carbon fixation. Glufonsinate can rapidly degrade into water, CO2 and nitrogen and has no residual soil activity. Glufosinate has been used world-wide as a non-selective foliar herbicide under various trade names, such as Basta, Ignite, Finale and Challenge. “LibertyLink” is the agronomic trait for resistance available from AgrEvo.

Resistance to glufosinate has been created through the insertion of the phosphinothricin-N-acetyltransferase (pat) gene, derived from the homologous gene from Streptomyces viridochromogenes. This gene codes for L-phosphinothricin (= glufosinate) acetyl-transferase (PAT), which catalyzes the acetylation of L-glufosinate to N-acetyl-L-glufosinate (see below for structures). Another name for a resistance gene is BAR.

Acetyl-L-glufosinate is not phytotoxic, and constitutive expression of the pat gene confers glufosinate resistance in transformed plants. PAT is stereospecific for the L-isomer of glufosinate; D-glufosinate in the racemic D,L mixture is not acetylated and is not phytotoxic to plants.

Transformants of Arabidopsis are selected by spraying phosphinothricin (PPT) at 100 mg/L 4 to 5 days after germination. Spraying is repeated three days later. The untransformed seedlings will turn to yellow color in three days and die in about one week. The transformants will keep green color. I suggest to repeat spraying 1~2 times more to eliminate possible pseudo-transformants.

“Finale” Concetrate Weed and Grass Killer: Purchase from Planet Natural, Bozeman, MT (www.planetnatural.com) $21.50 for 16 oz of concentrated 5.8% solution of glufosinate-ammonium:butanoic etc. equivalent to 0.5 lb per gallon or 60 gm per liter. Dilute 600 fold (1.7 ml per liter) for use. Pure Glufosinate-ammonium can be purchased from Riedel-de Haen (Cat#45520) but is expensive.