I. Isolation of Cell Walls

II. Cellulose Determination; Total Sugar and Uronic Acids

III. Fractionation of Cell Wall Polysaccharides

IV. Methyl Esterification of Uronic Acids

V. Monosaccharide Composition

VI. Linkage (methylation) Analysis

VII. Fourier Transform Infrared (FTIR)

VIII. Protocol for the screening of the UniformMu maize population with near infrared reflectance spectroscopy

VI. Linkage (Methylation) Analysis

METHYLATION OF POLYSACCHARIDES AND OLIGOSACCHARIDES

Two days before methylation:

All carbohydrates used should be lyophilized and "fluffy"

1. One mg or less of compound to be methylated should have been freeze-dried directly in a 15-ml Corex—if not, add it now and then add a dry "flea" stir-bar. Cover beaker with a Kimwipe and fix with a rubber band. Place the tubes in vacuum desiccator over fresh P205 (along with a separate supply of fresh, unpunctured serum sleeve stoppers in a separate, Kimwipe-covered beaker). Leave desiccated under vacuum for at least 24 h. About 8 samples (4 per beaker) is the limit per methylation run. More than that becomes unmanageable because of the importance of expedience during parts of the procedure.

Day of Methylation:

2. Release vacuum on dessicator through tube filled with Drierite. Seal sample tubes
immediately with the serum stoppers while they rest in the dessicator.

3. Turn on vacuum pump. Pierce stopples with an 18-gauge syringe needle connected to
vacuum pump by rubber tubing, and evacuate the sample tubes 30 sec each.

4. (Under hood). Let dry Argon run through small Tygon tubing fitted with a 20-gauge needle (see Appendix) for a few minutes then insert into serum stopper of dry, distilled DMSO (Pierce). Now put on those dispo gloves. With a 1-mL gas-tight syringe and extra long needle (precleaned sequentially with with RBS, water, methanol/or/acetone, and then oven-dry needle and barrel separated--Don't put piston in the oven because the Teflon seal will melt), add 1mL DMSO to each sample tube. Because the sample tube is under vacuum, the DMSO will be sucked into it. Try to aim the stream so most of it goes right to the bottom rather than running down the sides. Now go wash out the syringe (Water then acetone is enough from here on) and put it back in the oven—you'll need it two more times.

5. Place tubes in beaker containing enough water to reach liquid level in tube. Place beaker in sonic bath, also containing enough water to reach upper level of liquid in tubes.

6. Sonicate about 1.5 to 2 h (for oligosaccharides, about 1h is enough) until the bath has warmed to about 50° C and any globs of material have all broken up. If there are any pesky globs, you can stir them intermittently, but do NOT vortex them.

7. Remove tubes from sonication, pour out water, and dry outside of the tubes with Kimwipes. Tape the beaker to the center of a large mag. stirrer. You should have chosen a beaker so that the tubes (up to four in a 150-mL beaker or six in a 250-mL beaker) fit snuggly and stand, more or less, upright. Stuff in some wipes to take up any extra space. You may stir gently. As before, run some Ar through the Tygon tubings fitted with needles for a few minutes to ensure that all is dry. Flow should be strong enough to sense on your lips at a distance of three inches. Insert needle with Ar flow into each tube and then add a second needle as escape valve. This will keep a steady Ar purge during the reactions.

8. Insert another needle with Ar flow into the septum of a fresh bottle of 2.5 M n-butyllithium in hexanes. Don't shake up the precipitate on the bottom of the bottle. It's dirt. Wear those gloves (there's a bottle of talc near the middle sink to make them easier to re-don). Work carefully but quickly as you don't want to let water or O2 contaminate the reagent in the syringe. Now turn up the speed on the stir bars so the DMSO solns are whipped but not spattered. With a clean, dry syringe (see 4) withdraw 1 mL of butyllithium, and then pull up the needle slightly into the atmosphere in the bottle and pull up a little Ar until you see the bubble in the syringe barrel (to get the reagent out of the needle). This will ensure that the butyllithium will not drip out of the syringe onto the septum as well as protect it from atmosphere. Make sure you hold the syringe at the needle connector. If they come apart who knows what might happen. Pull up and insert into a sample tube all the way to about one inch above the DMSO. Do not insert into the DMSO or let it spatter on the syringe. Take a deep breath and add the reagent drop by drop until a milky white cap starts to form, then stop until it evaporates. Don't let that "white cap" form or it could spark (not good). A good whipping stir is essential to keep the cap from forming. Repeat until you have added 0.5 mL of the reagent. Pull up a little Ar from the purge flow, and go to the second tube and repeat. (Taking up 1 mL at a time reduces the number of times you must puncture the septum of the butyllithium reagent. Also punch a new hole each time.). Once you have finished all the tubes, turn down the speed of the stirrer to a gentle but rapid rate. Breath a sigh of relief. Now go wash that syringe before the reagent gels and you end up breaking the glass syringe tip trying to get the damn needle off. The traces of butyllithium will fizz and smoke in water so flush profusely. Cram the tip of the barrel into the delivery tubing of the house dH2O and let water run through the barrel for a few minutes.

9. The "white cap" evaporates quickly but a faint milky solution is left which takes about 15 min, depending on how good the flow of Ar is, to clear. Once clear, the solution will turn a faint yellow (done) to strong yellow (overdone) to bluish-green (way overdone). For oligomers, shorter is better than longer (30 min) whereas for polysaccharides, yields can improve if left for 1 to 1.5 h (starting to turn bluish).

10. Let the CH3I (Mel) from the cold room warm to room temp. Some applications may require CD3I or even CH3CH2I Pull the sash of the fume hood down as this stuff is mutagenic (but it mixes with water to form relatively inocuous methanol and iodine). How come you don't have your gloves on! Withdraw 1 mL of Mel with the washed, dry syringe and insert needle into the first tube. Turn up the stirring speed once again and add the Mel drop by ever so slow drop into the center of the whirring vortex until the solution turns colorless. Then add the rest of 0.5 mL. Withdraw some Ar from the tube atmosphere and repeat with remaining 0.5 mL in tube 2. Repeat until all are done. Recap (but don't tighten—just rest the cap on the bottle) the Mel each time as it is quite volatile, separate needle and syringe as it will tend to stick to each other when the Mel evaporates. Leave the disassembled parts in the hood to evaporate. Remove the Ar and escape needle from each tube as the Mel will tend to evaporate in the Ar purge as well. Turn off Ar cylinder now—it's too easy to forget about it. You'll notice that some gel, etc.; from spattering is still on the walls of the tube. Grasp all 4 (or 6) tubes in one hand and shake them, or "hand-vortex," to now spatter the DMSO-MeI solution on the walls of the tube to wash the crust down into the mix.

11. Let them stir about 1 h until the solution begin to yellow. How come you still don't
have your gloves on! With the sash still way down, pop the serum tops off gently and
throw them into some waste water to quench the Mel. Add about 7 mL of water via squirt
bottle into the mixture. Reaction dead. Stir vigorously for a few seconds; and I mean top
speed, and then add 1.5 mL of chloroform (Pasteur pipet-load) and stir additional 10 min. Spin at 3500 rpm for 5 min to separate phases. Pipet off the chloroform and put it in fresh Corex tube. Really try to avoid transfer of the still nasty aqueous phase along with the chloroform. Try no to let any flocculent interface from breaking up into the chloroform phase either. You want to avoid the interface material too. Repeat extraction with another 1.5 mL of chloroform. Combine chloroform phases; and wash with 7 to 10 mL of squirt-bottle water (whatever it takes to even them up for centrifugation) by vigorous stirring for 10 min and spin. Aspirate off water. Actually; if you are of average deft; you can pull off the last droplet of water without sucking off too much chloroform. Do this water wash a total of five times. Transfer the chloroform phase by Pasteur pipet into fresh 1-dram screw-cap vial and dry down under Ar in N-Evap. Really try hard to avoid any water again as this takes forever to dry. Cap with Teflon-lined cap. You now simply go to the ALDITOL ACETATE procedure and continue. Note: for normal applications, you'll want to reduce with NaBD4.

APPENDIX
All operations with Ar assume that you use an "N-Evap" (Organomation); a Ar delivery
system with 12 needle-valve-controlled ports with Luer-lock fittings. To make a flexible
delivery system for Ar; we pull out the needle shaft of a 20-gauge disposible needle with a pair of pliers and then insert the blunt end snuggly into some 1/16'1 (ID) Tygon tubing; exposing the needle of course. Give yourself about 12 in or so of tubing and slip the other end of the tubing over the needle end of another 20-gauge dispo needle. Cut the sharp tip off with a tube cutter and file smooth so that the needle won't puncture the tubing. We use the fitting to attach the tubing to the N-Evap. We make about 12 at a time.

A. Overview

This method has been around since the time of Fischer, but refinements in methods and the advent of GLC-MS have made the technique one of the most useful methods to determine the linkage structure of oligo- and polysaccharides over the past 30 years. If you can believe it, people used to crystallize individual derivatives first separated by paper chromatography, and then chemically determinal the positions of methyl ethers down the chain. The gist is to make methyl ethers at the free hydroxyl residues, hydrolyze the oligomer or polymer, reduce the anomeric carbon with sodium borodeuteride so that fragments from the top of the derivatives are distinguished from fragments from the bottom, and all remaining hydroxyls are acetylated. After separation of these derivatives by GLC, EIMS easily determines whether each carbon has a methyl and acetyl group. The anomeric and ring carbon always have acetyl groups, but if another sugar was attached to any of the remaining hydroxyl groups it will bear a third acetyl in place of a methyl group.

B. Preparation of Derivatives

C. Separations by GLC

D. Electron-Impact Mass Spectrometry


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