Potter Group Protocols
Preparing OCT blocks of kidney tissue
- Dissect out kidneys rapidly and store briefly (up to 20 min or so) in ice cold PBS.
- Process through OCT only as many kidneys as you plan to freeze in one block at a time. The OCT is hyperosmotic and tends to suck the water out of the tissue it touches, ruining the edges of the kidney, where a lot of interesting stuff is happening. So faster is better. But, at the same time, you need to totally rinse the PBS away, because it can interfere with the sectioning.
- Place kidneys in OCT in a 60 mm plate cooled with ice. Use OCT that has been pre-cooled on ice for at least ten minutes.
- Quickly mix kidneys in with OCT, then transfer carefully to a new 60 mm plate with cold OCT, quickly mix again, and place in a tinfoil mold with cold OCT covering the bottom.
- Cover kidneys with additional OCT and position kidneys near each other in mold, in central position, not too near the top.
- Immediately freeze in 2-methylbutane isopentan that is in a pyrex beaker resting in liquid nitrogen. The isopentan should be frozen. Hold the tinfoil mold with forceps to keep vertical and gently move to keep from freezing in the isopentan and to improve thermal contact.
- When the OCT is completely frozen place the mold in dry ice, and then store long term in liquid nitrogen freezer.
- Throughout this procedure be very careful not to cut yourself on the sharp blades used in the cryostat.
- Wear gloves throughout to reduce RNase contamination.
- Use the cryostat in Prasad’s lab. Place mold in the chamber for a few minutes to temperature equilibrate. Remove tinfoil. Place chuck that has been at room temp in chamber and let cool a minute, but not too much. Place OCT on chuck and let cool a minute, but not freeze, and then place tissue OCT block on chuck, and let freeze in position. Can place additional OCT around the base and spread with gloved finger to help hold in place.
- The tissue is too brittle to section properly if the chamber is too cold. I typically use a setting for the chamber and arm of –12 to –14. We have been using –14 mostly lately, with success.
- Use the trim setting of 40-60 to remove most of excess OCT, til see tissue. When getting close to tissue can drop back from trim to regular setting, of 9 microns for Veritas, with UV cutting, or 7 microns for old Pixcell II machine. Use a regular glass slide to check for presence of tissue in sections. When you hit a good chunk of tissue start saving on the membrane slides (see below).
- Membrane slides are prepared as follows to allow good sticking of the sections to the slides. Dilute the Sigma poly-lysine solution one to ten, as recommended by Sigma. Dip slides and dry in vertical position.
- Need to carve the OCT block with a razor blade, so the tissue sections are not too large. Be very, very careful here not to cut yourself on the blade in the cryostat, which is very sharp.
- Collect sections on the slides. Do not want the tissue sections to be arranged so that they end up under the strut supports of the caps, as this would give cap crap. So the sections should be spaced so that the cap can rest with one tissue section centered, and with no other tissue section under the edges. So space them out, and try to get 5-10 sections per membrane slide. Try to work fairly fast, as the RNA can go bad sitting in one section at room temp while other sections are being cut on the slide.
- Try to get the best sections possible. Sometimes having the cut part go more slowly helps. Sometimes warming a degree or two helps. Sometimes changing the way you catch the section with the brushes as it comes off of the blade helps.
- Place the slides with sections in box with dry ice against the slide, and then store at –80ºC.
Processing slides for LCM
- Remove slides from –80°C freezer, store temporarily on dry ice, air dry on metal slide warmer turned off or set to about 30 to 35°C, for 3 min.
- Fix two minutes. 1:1 mix of acetone 75% ethanol gives a nice compromise, with good section sticking and reasonable histology, and good lectin staining.
- Rinse 2 min in 1/10X PBS, ice cold, to dissolve off some OCT. The 1/10 PBS is made by diluting 1X PBS ten fold with sterile autoclaved super water.
- Stain about 6 min in lectin, on ice. For PNA use 5 µl per ml of 1/10 PBS. Stain on ice.
- Rinse 1/10 PBS, ice cold. Two gentle 10 sec dips and then 3 min.
- Dehydrate, in ethanol, 75%, 95%, 100%, 10-15 sec each, with 2-3 dips, and then another100% one minute.
- Xylene 1.5 min with three dips.
- Xylene 2 min with two dips.
- Air dry about 2-4 min.
Laser Capture Microdissection
- Use the Veritas machine, (or palm zeiss when available), with membrane slides, pretty much as per standard protocols, but with following considerations.
- Too strong a cutting laser might degrade RNA according to Chris Erwin. So cut back on power as much as possible. I’ve had good luck with the setting at 3.0.
- Be careful to avoid cap crap. But if there is a little then can use ablation to remove. But be careful to re-set laser after ablation, so don’t ruin sample.
- Label caps clearly and take photos before, after and of caps to record material captured.
- Place caps in 0.5 ul tubes after capture is complete.
- Store on dry ice, then at –80ºC.
LCM RNA preparation
- Make up RLT by adding 10 ul of pure beta mercaptoethanol per ml of RLT (in the hood),
- VERY IMPORTANT. ADD 1/2 UL OF EPICENTRE POLYINOSINE PER 100 UL OF RLT. THIS CARRIER IS REQUIRED TO PROTECT THE RNA SAMPLE AND PREVENT LOSS ON THE COLUMN.
- Take 0.5 ml tube with cap from –80ºC freezer and let thaw on ice. The tube is brittle if too cold, and will crack on handling, resulting in loss of sample.
- Remove the cap, using a vise grip, and add the 100 ul of RLT WITH CARRIER to the tube.
- Place the cap back on the tube, and invert, and tap the tube to get the RLT to drop onto the cap.
- Vortex vigorously for about a minute. Must vortex very strongly to remove the membrane from the CAP and expose the cells to RLT buffer to isolate RNA. Will get a very low yield if you do not vortex vigorously, and check for membrane removal with a dissecting scope after next step.
- Spin in centrifuge, with adaptor tube, for 30 sec to get sample back to the bottom of the tube. Remove and discard cap.
- Add 100 ul of 70% ethanol to the tube and mix by pipeting, and place on RNeasy minelute column.
- Spin 30 sec 2000 rpm, then 15 sec top speed. Discard flow through.
- Add 700 ul of RW1 buffer, spin top speed room temp 15 sec and place in new collection tube.
- Add 500 ul of complete RPE, spin top speed, and discard flow through.
- Add 500 ul of 80% ethanol and spin top speed 2 min, with cap connection at bottom, and discard flow through and collection tube.
- Place in fresh collection tube, rotate column 180 deg from previous spin, with cap connection now at the top, leave the cap open, and spin 5 min full speed, to dry column and remove any residual fluid.
- Transfer column to a fresh collection tube, add 14 ul of RNase-free water directly to center of the column, wait 1-5 min then spin 1 min top speed to elute RNA.
FACS analysis for E15.5 mouse kidneys
1. Dissect out and isolate E15.5 mouse kidneys in ice-cold PBS.
2. Place four (4) kidney in a 1.5ml eppendorf tube.
3. Add 400ul 0.05% Trypsin-EDTA.
4. Incubate 37ºC, five (5) minutes.
5. Triturate 30X.
6. Add 600ul ice-cold 0.1% BSA/PBS or 2% FBS/PBS.
7. Triturate 30X.
8. Centrifuge tubes at 1500 rpm for five (5) minutes. Keep cells cold.
9. Remove supernatant.
10. Resuspend pellet with 600ul ice-cold 0.1% BSA/PBS or 2% FBS/PBS.
11. Recentrifuge tubes at 1500 rpm for five (5) minutes. Keep cells cold.
12. Remove supernatant.
13. Resuspend pellet with 600ul ice-cold 0.1% BSA/PBS or 2% FBS/PBS.
14. Filter cells using 70um mesh FACS collection tube. Keep cells on ice.
15. Immediately FACS using a high-speed digital BD FACS Aria II Cell Sorter.
Use TargetAmp 2-round aminoallyl amplification kit from Epicentre biotechnologies. Add an additional 3/4 ul of SBI semi-random primer at step one, before speed-vacing to 3 ul. Also add 1 ul of this primer for part F, step one, in addition to the Epicentre primer.