C. elegans RNA prep

互联网2013-09-06

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Introduction:
C. elegans RNA preps that have been widely used previously involved fairly slow lysis of the worms, potentially leading to degradation of the RNA. In addition, they failed to separate RNA from genomic DNA, leading to difficulties in determining the yield and purity of the RNA. Also, the contaminating AT-rich genomic DNA may interfere with poly A selection of the RNA. This protocol is based on standard methods that have been used for many years to isolate RNA from mammalian tissues and Drosophila; it involves very rapid lysis of the worms and subsequent purification of the RNA to homogeneity.

Precautions against RNase contamination:
Because RNase is incredibly stable and is widely used in labs during DNA preps, precautions must be taken when preparing solutions, plasticware, and glassware for use with RNA to avoid contamination with this enzyme. Glassware (but not the plastic caps!) stir bars, and spatulas for weighing reagents out should be baked overnight in a 180o C oven (Horvitz lab: this is in the autoclave room). Plasticware (pipette tips, centrifuge tubes, 10 ml disposable pipettes) should be from an unopened package that is labelled RNase free and then kept in a special drawer after opening. Most solutions, as detailed below, can be treated with diethyl pyrocarbonate (DEPC) to inactivate RNases, and then autoclaved to eliminate the DEPC. DEPC reacts with amino groups, and therefore cannot be used, for example, on Tris buffered solutions. Always wear clean gloves when handling RNase free materials. If at all possible, use unopened clean containers of reagents to make up all solutions, label the bottles " RNase free" , and store in a special drawer. When weighing out reagents, clean the balance first and use weigh boats from an unopened container.

Solutions:

Caution: Wear gloves and avoid getting DEPC on your skin.

RNase free water (make several 100 ml bottles)

1.0 M Tris pH7.5
Make using clean technique and DEPC treated water
autoclave

TE (10 mM Tris pH7.5, 1 mM EDTA)
Tris solutions cannot be DEPC treated.

TE + 0.1% SDS
Tris solutions cannot be DEPC treated.

Homogenization buffer

Cesium cushion solution

3M sodium acetate pH 5.2

RNase free 100% EtOH

Use a fresh bottle, then label RNase free and keep clean

RNase free 70% EtOH

Prepare using the clean 100% EtOH and DEPC treated water

1. Start with a pellet of purified worms grown in liquid culture. It is best to have 1-5 mls of worm suspension (~50% worms in 0.1 M NaCl) in a 50 ml disposable centrifuge tube. If you have more than 5 mls of suspension, the tube will overflow when you homogenize. It's probably better to use freshly prepared worms, since these can be lysed most quickly, but a flash frozen pellet that was stored at -80o is fine.

2. Using a motorized tissue homogenizer is the fastest method to lyse the worms. By lysing quickly in a denaturing solution that inactivates RNases, degradation of the RNA can be minimized. We've been using the Brinkmann model 10/35 with a PTA10S tip (this brand of machine is popularly known as a " polytron" ). Be extremely careful when using this machine; it is very powerful and a mishap could be disastrous. The most important safety precaution is to make sure the tip is fastened on the the motor very tightly. During operation the assembly (a big silver nut) that fastens the tip on can start to become unscrewed due to the vibrations; keep your eye glued on the nut and immediately shut the machine off if you see it begin to unscrew! Note: we're not sure that the polytron will lyse eggs or dauers; this may require sonication.

3. After adding BME to the homogenization buffer, add 5 volumes of homogenization buffer to the worm suspension. If using a frozen pellet just pour the buffer on the frozen pellet. Immediately begin homogenizing. With a frozen pellet, start the machine a low speed and push the tip into the pellet to break it up and release it from the bottom of the tube. As soon as the frozen chunks are gone, turn the machine up to full speed, and homogenize for 2 minutes. (The old polytron from the Whitehead seems to turn the solution blackish by releasing small metal? chunks; don't worry, you can spin these out.)

4. Add sarcosyl to 0.5 % (using the RNase free 20% solution) and mix.

5. Pour the mixture into a 28 ml polycarbonate Oak Ridge tube. Spin in the 70Ti ultracentrifuge rotor for 20 minutes at 30K RPM at 20o to pellet debris (and any metallic stuff from the polytron). Pour off the supernatant into a clean 50 ml disposable tube. It should be brownish yellow.

6. You will need to prepare one cesium gradient for each 6 mls of homogenate. Use RNase free 16X102 mm polyallomar tubes, which fit the SW28.1 buckets (which in the Horvitz lab we spin on an SW28 rotor). Don't use " ultraclear" tubes; these crack during the spin. Measure out 11.5 mls of the cesium solution (use a disposable plastic pipette) into each tube. Tap the tubes to eliminate any air bubbles that might be attached to the tube walls within the solution.

7. Draw the worm homogenate into a 5 ml hypodermic syringe fitted with a 23-gauge needle. Layer the sample over the cesium pad by slowly drizzling it down the side of the tube. Fill the tube up to 1-2 mm below the top: this should be ~6 mls of homogenate per tube. If you run out of homogenate with a tube only partly filled, you can fill it up to the top using homogenization buffer + 0.5% sarcosyl. With a felt tip pen make a mark at the interface between the cesium pad and the homogenate.

8. Carefully transfer the tubes to SW28.1 buckets, screw on the caps firmly, carefully hang the buckets and mount the rotor in the ultracentrifuge, always making sure not to disturb the gradients. Spin at 20o C for 24 hours at 27K RPM. Use program #4 (on the Beckmann L8-80 centrifuge) for both acceleration and deceleration so as to disturb the gradient as little as possible before and after the spin. It's OK to do the spin for a few more or less hours if necessary.

9. After the spin, carefully remove the tubes from the buckets into a rack, taking care not the disturb the gradient. The brownish proteins should still be above the felt tip pen mark; you may see a whitish doublet band within this brown material. Further down in the clear part of the cesium gradient there should be another white band; this is the DNA. The RNA is a crystal clear gelatinous pellet LOOSELY attached to the bottom of the tube; it will remain invisible until almost all the liquid is removed. The pellet looks about like a wispy flattened chunk of low melt agarose, with a volume of ~30 ml.

10. Removing the liquid requires great care, both to avoid contaminating the RNA with material from higher up in the gradient, and to avoid losing the difficult-to-see RNA pellet. Using a 10 ml disposable plastic pipette attached to a rubber bulb, carefully suck off the supernatant down to the felt tip pen mark. Do this by holding the pipette tip at the very surface of the liquid, so that both liquid and air are sucked up; this ensures that the least dense (and most contaminated) material at the top of the gradient will be removed first, and won't just be lowered down in the tube as the solution below is sucked out. Change to a fresh pipette, and suck off more liquid, in the same manner, down to just below the whitish DNA band. Change pipettes, and suck most of the rest of the liquid, leaving ~2 mls, so that liquid just fills the curved bottom part of the tube. The remaining liquid will have to be removed extremely carefully to avoid losing the RNA pellet.

11. Hold a new razor blade in a bunsen burner (using forceps or a hemostat) until it is red hot, and use the blade to cut (really melt) through the tube just above the level of the remaining liquid. It is convenient to leave a small attachment between the upper part of the tube and the bottom so that the bottom of the tube can be carried around using the upper part as a handle. The point of removing the top of the tube is to separate the contaminated walls of the tube from the clean bottom portion, and to help you to see the RNA pellet in the bottom.

12. Using an RNase free 200 ml pipette tip, VERY CAREFULLY remove the remaining liquid. The clear gelatinous RNA may be floating or in several pieces at this point; sometimes it helps to tilt the tube bottom around to try to separate the liquid from the pellet so that the liquid can be safely sucked off.
Sometimes it is truly impossible to remove the last ~30 ml of liquid without sucking up chunks of RNA. In this case, just leave the last bit of liquid and add about 2 volumes of room temperature 100% ethanol (RNase free). This will cause the remaining CsCl to form a white precipitate which, along with the RNA, will stick to the bottom of the tube better. Then just go ahead with the next step (filling the tube with 70% EtOH). Eventually all this cesium will be eliminated in the next ethanol precipitation.

13. Fill the bottom of the tube with room temperature 70% EtOH (this soaks out the remaining CsCl). The RNA will turn white over the next couple of minutes, and will attach to the tube much more firmly.

14. Pour or suck off the liquid, and dry the pellet briefly. When the drying RNA starts to turn clear, add 175 ml of TE + 0.1% SDS to each tube, and suspend the RNA by pipetting up and down. This should take about 1 minute. Transfer the RNA to a fresh RNase free screw cap eppendorf tube. Wash out the bottom of the ultracentrifuge tube with another 25 ml of TE + 0.1% SDS, and combine with the rest of the RNA. If the RNA is not completely in solution (sometimes a few chunks remain) freeze it in liquid nitrogen and thaw in a 50o water bath once or twice.

15. To the 200 ml of RNA solution, add 150 ml TE (without SDS), 30 ml 3M sodium acetate pH 5.2, and 900 ml EtOH. Should see a fluffy precipitate. Chill at -20o for 30 minutes, and microfuge in the cold room for 10 minutes. Discard the supernatant, wash the pellet with 70% ethanol, dry, and dissolve in a small volume of water, (perhaps ~80 ml, depending on how big the pellet looks).

16. Measure the concentration and purity of the RNA by making an appropriate dilution (~1000 fold) in water and taking the OD260 and OD280. RNA prepared by this method should be comletely pure, with an OD260/OD280 of 2.0. For calculating the RNA concentration, a solution of OD260=1.0 is 40 mg/ml.

16. Store the RNA at -80o. Diluting all samples to a standardized concentration (say 5 mg/ml) and aliquoting it is a good idea.

17. Yield: 0.8 to 1.1 mg of pure total RNA per ml of packed worms. These preps will be contaminated to some extent with RNA from E. coli that were in the starting material. The proportion of E. coli RNA in the prep can be assessed by running the RNA out on a gel, blotting, and staining the RNA on the blot with methylene blue; the C. elegans rRNAs, which run at 3.5 and 1.7 kb, can be resolved from the E. coli rRNAs, which run at 3.0 and 1.5 kb (see our Northern blot protocol for details on these methods). We found that if the worms were prepared using our liquid culture protocol, the amount of E. coli RNA in the final prep varied from undetectable to about 40% of the RNA.