Midi-RNA Extraction from OCT Embedded Tissue and/or Snap Frozen Tissue

Collect and record Sample(s)
1. Fill a Little Playmate Cooler ¼ of the way with Dry Ice.
2. Keep all samples on dry-ice until needed, if more than ½ an hour is needed for set-up; place the samples in the -80 freezer in the Molecular laboratory.
3. Record each sample identification number as appropriate on a sample tracking/extraction worksheet. Assign each sample a different extraction number.

Set-up/Preparation:1. Label two (2) sterile, 15mL conical tubes for every sample to be extracted. Set the tubes up in order on two separate tube racks.
2. Remove the Glycogen from the -20 freezer and allow it to thaw completely on wet ice.
3. Turn on the Sorvall T21 centrifuge and the Sorvall Biofuge Pico. Allow both centrifuges to cool to 4 degrees Celsius before using.
4. If not already done, prepare a stock of 3M Sodium Acetate, pH 5.2.
5. If not already done, prepare a stock of 70% ACS Grade ethanol.
6. 64% Ethanol:  Add 64 ml of ACS Grade 100% (200 proof) ethanol to the bottle containing 36 ml RNase-free water in the RNAqueous kit. Mix well. Place a check in the empty box on the label to indicate that the ethanol has been added.
7. Wash Solution #2/3:  Add four volumes (201.6 ml) of ACS Grade 100% (200 proof) ethanol to the 50.4 ml of Wash Solution #2/3 Concentrate. Mix well. Place a check mark in the empty box on the label to indicate that the ethanol has been added. A precipitate will usually form in the bottle upon storage; this is excess EDTA falling out of solution. Leave these crystals in the bottle when removing wash solution for use.
8. Thoroughly clean the BSL-2 hood and place down fresh absorbent pads in the workspace where the extraction will take place.

Cell Lysis and RNA Extraction
1. Homogenize and lyse the cells using the Polytron Homogenizer.
2. Upon the completion homogenize and suspend in 5 ml of TRIzol Solution in a 30ml, round-bottom polypropylene tube. Shake all tubes thoroughly to ensure the homogenization of cell lysis and TRIzol solution.
3. Place the 30 ml tubes in a rack in the hood and allow them to incubate at room temp. for 10 minutes.
4. While the samples are incubating, set up the Promega Vac-Man vacuum manifold. Make sure that all ports have a new, clean stop-cock. Obtain a clean syringe filter (included with the kit) for each RNA sample and label one for each sample extraction number. Fasten each syringe filter to a stop-cock. Now obtain a clean, sterile 10ml syringe for each RNA sample. Remove each syringe from its individual packaging and attach them to the syringe filters on the manifold, leave the plungers in the syringes for the time being. Place a rubber stopper in the drainage port of the manifold. Put all of the stop-cocks that have a filter and syringe in the open position. Those with no filter and syringe need to remain shut throughout the procedure.
5. At the end of the 10 minute incubation period, add 1000ul of molecular grade Chloroform to each tube containing the cell lysis homogenate. Shake each tube vigorously for 15-20 seconds to ensure complete mixing of the chloroform and the homogenate. Caution: Chloroform is a known carcinogen. Always handle it in a hood with proper ventilation and dispose of it by the proper laboratory means.
6. Transfer the chloroform/homogenate from the 30 ml tubes to the clean pre-labeled 15ml conical tubes.
7. Spin the tubes at 12,000 x g (or 10,000 rpm) for 30 minutes at 4 degrees Celsius.
8. While the samples are spinning, set-up the custom-made sample elution vessel. Label a clean, sterile 15ml round-bottom tube for each RNA sample being extracted. Place the tubes in consecutive order in the interior rack of the elution vessel. Place the top on the vessel. The top of the vessel contains ports that the syringe filters will fit into. Any port that has no collection tube directly below it must be sealed with tape so that it is air tight.
9. Fill a 250ml glass beaker halfway with deionized water. Place the beaker on a hot plate. Obtain the Elution Solution from the RNAqueous kit and fill a 50ml conical with enough solution so that each sample extraction can receive 3ml of the Elution Solution (ex: if there are 10 samples, fill the tube with at least 30 ml of solution). Cap the tube and place it in the glass beaker. Turn the hot plate on high heat and bring the water in the beaker to a boil.
10. When the samples have finished spinning, remove the tubes from the centrifuge. Each tube should now contain 3 layers, an upper clear layer, a middle solid layer, and a lower pink layer. Bring the tubes back to the hood.

11. Remove the upper clear aqueous layer from each tube and pipette it into a clean, sterile, pre-labeled 15 ml conical tube. Be very careful to only pipette up the clear layer and none of the two lower layers. Place each tube with the aqueous layer in wet ice until this step is completed for all samples.

12. Add 1 volume of chilled 64% ethanol to the aqueous layer in the new tubes. Shake each tube vigorously for 15-20 seconds to ensure complete homogenization of the aqueous layer and the ethanol.

13. Remove the sample tubes to the bench top and turn on the vacuum supply. Remove the plungers from the syringes on the manifold and pour each sample into its corresponding syringe. Gently tough the vacuum hose to the vacuum port of the manifold and allow the sample solutions to be pulled slowly through the syringe filters. Be careful not to apply too much vacuum pressure to the manifold or the filters may rip, causing the RNA to be lost. When a sample has been pulled completely through the filter, close off its stop-cock. Continue pulling the samples through until all stop-cocks are closed.
14. Once all the samples have been pulled through their filters, open up all the stopcocks and apply additional vacuum pressure to the manifold. Watch the neck of the stop-cocks and you will see a foamy substance being pulled down. Continue applying gentle pressure until all of this foamy substance is pulled out.
15. Remove a 10ml serological pipette from its wrapper and add 6ml of Wash Solution # 1 to each syringe/filter set-up. Gently pull the wash solution through using the vacuum pressure and closing off the stop-cocks as you go. When all of the Wash Solution #1 has been pulled through each filter, open the stop-cocks up and apply gentle pressure to remove the foam.

16. Remove a 5ml serological pipette from its wrapper and add 4.20ml of Wash Solution #2/3 to each syringe/filter set-up. Gently pull the wash solution through using the vacuum pressure and closing off the stop-cocks as you go. When all of the Wash Solution #2/3 has been pulled through each filter, open the stop-cocks up and apply gentle pressure to remove the foam.

Now repeat step 16 a second time!

RNA Elution and Precipitation

1. Upon completing all of the wash steps, dispose of the syringes properly. Remove each syringe filter from the manifold and snap them firmly in place in the lid of the elution apparatus so that its corresponding collection tube will be directly below it when the lid is properly in place.

2. Drain the pull-through solution inside of the vacuum manifold into the flammable waste container in the chemical storage hood. Spray Envirocide inside of the manifold, fill it with water, and then leave it tipped up in the sink to drain. When the manifold is dry put it away.

3. Remove a clean, sterile 5ml syringe from its wrapper and attach one to each of the syringe filters now snapped into place on top of the sample elution apparatus. Remove the plungers from each syringe.

4. Make sure that the gasket of the elution apparatus is in place and that the seal between the lid and the chamber is air tight.
5. By now, the water in the glass beaker should be boiling and the elution solution in the 50 ml tube in the water should be at the proper temperature. Remove the tube from the water and add 1ml of Elution Solution to each of the 5ml syringes attached to the syringe filters.

6. Open the stop cock to the chamber and apply gentle vacuum pressure to the chamber so that the elution solution is pulled completely through each filter and into the corresponding collection tubes.

Now repeat step 6 2 more times so that each filter receives a total of three milliliters of Elution Solution.7. Remove the lid from the chamber and dispose of the syringe filters. At this point, the RNA should be in solution in the collection tubes.

8. Add 300ul of 3M Sodium Acetate, pH 5.2 to each collection tube.

9. Add 3ml of 100% Isopropanol to each collection tube.

10. Add 1ul of molecular grade Glycogen to each collection tube.

11. Cap each collection tube and invert them several times. As you invert the tubes, ripples should appear on the sides of the tubes. Continue inverting the tubes until these ripples have disappeared.
12. Place each collection tube in a rack and incubate the samples at -20 degrees Celsius for at least one hour. This incubation stage allows for the precipitation of the RNA.
13. After the hour incubation period, remove the samples from -20 and spin them at 12,000 x g (or 10,000 rpm) for 30 minutes at 4 degrees Celsius.

14. When the samples have finished spinning, a white pellet should be noticeable at the bottom of each tube.

15. Pour off the solution in each tube into a waste beaker being careful not to dislodge and lose the pellet. Dab the opening of each tube on a stack of Kim wipes or paper towels to remove any residual solution.

16. Allow the tubes to air dry for a minute and while this is happening, label a 1.5 ml Eppendorf tube for each sample.

17. Wash the samples by resuspending the pellets in the 15ml tubes with chilled 70% Ethanol. Pipette the mixture up and down several times to ensure that the pellet becomes fully resuspended.

18. Fill the tube holes in a heat block with RNase-free water. Set the heat block to 65 degrees Celsius and allow it to heat up.

19. Spin the samples in the Eppendorf tubes at 12,000 x g or (10,000 rpm) for 5 minutes at 4 degrees Celsius.

20. When the spin is complete, pour off the ethanol making sure not to dislodge and lose the pellet. Dab the mouths of the Eppendorf tubes on a stack of Kim wipes to remove any excess ethanol.

21. Use a clean sterile cotton-tipped applicator to remove any residual ethanol inside each tube. Make sure to use a new applicator for each sample and be careful not to make contact with the RNA pellet.

22. Discretion must now be taken when resuspending the pellets in RNase-free water. If the pellet is small (approximately the size of a pin-head) add 100ul of RNase-DNase-Free water. If the pellets are larger, anywhere from 110 – 200ul of water may be used.

23. Once you have added the water to each sample place the tubes with the caps open in the heat block. Leave the tubes on the heat block for about 1 minute as this will aid in removing any left over ethanol.24. After a minute, resuspend the pellets in the water by gently pipetting up and down several times.
25. Keep all samples on wet ice until needed for Quantitation and QC.

Immunoprecipitation (IP)

1. Solutions and Reagents
   Lysis buffer:
   1. Typically use RIPA buffer (25 mM Tris-HCl pH 7.6, 150mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS). Proteinase inhibitor cocktail should be added fresh before each use.
   2. The usage of SDS depends on the nature of the cells. For some cell lines, 0.1% SDS will release DNA and thus make it hard to extract proteins out. In those cases, omit SDS
2. Preparation of cell lysates
   1. Add ice cold lysis buffer (1ml per 100mm-dish or 107 cells, or adjust based on your specific requirements). Scrape off cells (for adherent cells still on plate) and resuspend cells. Collect cells in a centrifuge tube and agitate for 30 min at 4°C.
   2. Spin cells at 4°C for 20 min at 12000 rpm.
   3. Save the supernatant which is the cell lysates.
3. Pre-clearing
   1. Add normal serum or irrelevant antibody from the same species and isotypes as the IP antibody you will use. The amount should be at least 5-fold more than the amount you will use for IP. Incubate for 1 hr at 4°C.
   2. For 1 ml lysate, add 100 ul of proteins A or protein G beads slurry (50 ul solid bed volume), and incubate at 4°C for 30 min on a rotator.
   3. Spin down beads at 14000g for 5 min at 4°C.
   4. Save the supernatant which is the pre-cleared lysates.
4. D. Immunoprecipitation (IP)
   1. Add IP antibody to the pre-cleared lysates. You will need to determine the best amount of antibody to use. As a starting point, you may use 1 ug antibody for every ml of lysates.
   2. Incubate for a certain amount of time (from 1 hr to overnight, depending on your specific conditions) at 4°C.
   3. Add 100 ul of protein A or protein G slurry (50 ul solid bed volume) to 1 ml lysate and incubate for 3 hr at 4°C on a rotator.
   4. Spin down beads, and remove supernatant.
   5. Wash beads 3 times with lysis buffer.
   6. Add SDS-PAGE sample buffer to beads. Boil and run gel.

FACS Protocols

Flow Cytometry for Intracellular Staining

1. Solutions and Reagents
   1. 1X Phosphate Buffered Saline (PBS): Dissolve 8g NaCl, 0.2g KCl, 1.15g Na2HPO4 and 0.2g KH2PO4 in 800mL distilled water (dH2O). Adjust the pH to 7.4 with HCl and the volume to 1 liter. Store at room temperature.
   2. Fixation buffer: 2% paraformaldehyde in 1xPBS
   3. Permeabilization buffer : 0.1% Triton X-100 in 1xPBS
   4. FACS buffer: 0.5% BSA , 0.05% Azide in 1xPBS
   5. Fluorescent dye conjugated secondary antibody.
2. Fixation
   1. Collect cells by centrifugation and aspirate supernatant.
   2. Fix the cell by 125μl cold fixation buffer, vortex briefly.
   3. Incubate at room temperature for at least 30 min or for 1hr 40C.
   4. Centrifuge for 5min at 300g,remove the supernatant.
3. Permeabilization
   1. Add 1ml permeabilization buffer to each tube.
   2. Centrifuge briefly, and aspirate supernatant.
   3. Resuspend cells in 125μl of permeabilization buffer and incubate at room temperature for 5min.
4. Staining
   1. Aliquot 1-2x106 cells into each tube.
   2. Add 1 ml FACS buffer to each tube, centrifuge to pellet the cells.
   3. Resuspend cell pellet with 125μl FACS buffer containing diluted primary antibody, vortex and incubate on ice for 30min.
   4. Rinse as before in FACS buffer by centrifugation.
   5. Resuspend cells in fluorescent dye conjugated secondary antibody, diluted in FACS buffer per manufacturer’s recommendations.
   6. Incubate for 30 minutes on ice.
   7. Rinse the cells as before in FACS Buffer by centrifugation.
   8. Resuspend cells in 0.5 ml PBS and analyze on flow cytometer

Immunofluorescent Staining Protocol

Immunocytochemistry

1. Solutions and Reagents
   1. 1X Phosphate Buffered Saline (PBS): Dissolve 8g NaCl, 0.2g KCl, 1.15g Na2HPO4 and 0.2g KH2PO4 in 800mL distilled water (dH2O). Adjust the pH to 7.4 with HCl and the volume to 1 liter. Store at room temperature
   2. Poly-L-lysine solution: 0.1mg/ml in 1xPBS
   3. Glass coverslips No.1, 18mm dia.
   4. Fixation buffer: 4% paraformaldehyde in 1xPBS
   5. Permeabilization buffer : 0.1% Triton X-100 in 1xPBS
   6. Blocking buffer: 5% Fetal Bovine Serum (FBS) in 1xPBS
   7. Fluorescence-labeled secondary antibody
2. Fixation permeabilization
   1. Coat the coverslips with 0.1mg/ml poly-L-lysine solution at room temperature for 2hrs, dry, and then wash with 1xPBS buffer.
   2. Place the coated coverslip into each well of 12-well plate, and inoculate cells the day before immunocytochemistry experiment.
   3. Suck off the medium and rinse cells attached to cover slips twice with 1xPBS, removing liquid by gentle aspiration in this and subsequent steps.
   4. Fix cells with 4% paraformaldehyde in 1xPBS for 6 min at room temperature, and then rinse briefly twice with 1xPBS.
   5. The fixed cells can be permeabilize with 0.1% Triton X-100 in 1xPBS for 6 min.
   6. Wash cells briefly twice with 1xPBS, then block the coverslip with blocking buffer briefly at R/T.
3. Staining
   1. Dilute primary antibody with blocking buffer, and incubate the coverslip for 60 min at room temperature.
      Note: You may wish to leave one slip for a secondary antibody only control.
   2. Wash cells 3 times with 1xPBS, then 2 times with blocking buffer.
   3. Incubate cells with a dilution of the fluorescence-labeled secondary antibody in blocking buffer for 30–45 minutes at room temperature in the dark.
   4. Wash cells three times with 1xPBS.
   5. Mount the coverslip on a glass slide. Store the slides in the dark.

IHC Protocols

Immunohistochemistry Protocol for Paraffin-embedded Tissues

1. Solutions and reagents
   1. Xylene
   2. Ethanol, anhydrous denatured, histological grade (100%, 95%, 70%)
   3. Washing buffer:
      TBST washing buffer: 1XTBS/0.1% Tween-20
      To prepare stock solution of 10X TBS: add 24.2 g Trizma base and 80 g sodium chloride to 1L of dH2O. Adjust pH to 7.6.
      Working solution: 1XTBST/0.1% Tween-20: add 100ml 10XTBS to 900 ml dH2O. Add 1 ml Tween-20 and mix well.
   4. Distilled water (dH2O)
   5. Antigen Retrieval Solution:
      0.01M Sodium Citrate Buffer, pH 6.0
      To prepare stock solutions: Solution A. 0.1 M citric acid solution: dissolve 21.0 g of citric acid, monohydrate (C6H8O7.H2O) in 100 ml of dH2O. Solution B. 0.1M sodium citrate solution: dissolve 29.4 g trisodium citrate dihydrate (C6H5Na3O7.2H2O) in 100 ml of dH2O.
      Working solution: Add 9 ml of Stock solution A and 41 ml of stock solution B to 450 ml of dH2O. Adjust pH to 6.0.
   6. 3% Hydrogene Peroxide
   7. Blocking buffer:
      1xPBS: This buffer is made by dissolving 8g of NaCl, 0.2g of KCl, 1.44g of Na2HPO4 and 0.24g of KH2PO4 into 800ml of distilled water. Then adjust the pH to 7.4 with HCl, and add H2O to 1 liter. Add 10% serum to make the final blocking buffer (serum origin depends on the host of the secondary antibody)
   8. Hematoxylin QS (catalog #H-3404 from Vector Laboratories, Inc.)
   9. Permanent Mounting medium (VectaMount, catalog# H-5000 Vector Laboratories, Inc.)
      
2. Protocol
   1. Deparaffinization/Rehydration
      1. Heat slides in an oven at 65 °C for 1 hour.
      2. De-paraffinize/hydrate using the following series of washes: two Xylene washes (5 min each), followed by two 100% ethanol rinses (5 min each), followed by 95% ethanol, 70% ethanol, 50% ethanol, 30% ethanol, followed by H2O and a TBST wash for 5 min on a shaker.
   2. Antigen Retrieval
      1. Immerse slides into staining dish containing Antigen Retrieval Solution.
      2. Place covered staining dish into the rice cooker. Add 120 mL d H2O and press “cook”.
      3. When “cook” is turned to “warm” (about 20–30 min), unplug the cooker and remove the staining dish to the bench top.
      4. Allow to cool down, without cover, for 20 min.
   3. Staining
      1. Wash slides with TBST for 5 min on a shaker.
      2. Inactivate endogenous peroxidase by covering tissue with 3% hydrogen peroxide for 10 min.
      3. Wash slides three times with TBST (3 min each on a shaker).
      4. Block slides with the blocking solution for 1 hour.
      5. Dilute primary antibody in the blocking buffer per recommendation on the data sheet.
      6. Apply primary antibody to each section and incubate overnight in the humidified chamber (4 ºC).
      7. Wash slides three times with TBST (3 min each on a shaker).
      8. Apply to each section secondary HRP-conjugated anti-rabbit antibody diluted in the blocking solution per manufacturer’s recommendation; incubate for 1 hour at room temperature.
      9. Wash slides three times with TBST (3 min each on a shaker).
      10. Add freshly prepared DAB substrate to the sections.
      11. Incubate tissue sections with the substrate at room temperature until suitable staining develops (generally 2–5 min).
      12. Rinse sections with water.
      13. Counterstain with Hematoxylin.
      14. Rinse sections with water.
      15. Dehydrate samples using two rinses with 100% Ethanol (20 dips per rinse) followed by two rinses with Xylene (30 dips per rinse).
      16. Mount coverslips on slides using Permount medium.

Western Blot Protocols

Western blot analysis

1. Run SDS-PAGE gel, and then Western transfer the protein samples to nitrocellulose (NC) membrane for immunoblot analysis.
2. After transfer, transfer the membrane to western-blot tray, briefly wash the NC membrane with distilled water.
3. (Optional) Visualize the proteins on the membrane by Ponceau’s staining.
4. Wash off the red stain with distilled water.
5. Block the membrane with 5-10ml blocking buffer (made by 5% non-fat milk in 1xPBST) for 30 minutes at R/T.
6. Dilute the primary antibody with blocking buffer according to the suggested dilution factor on datasheet (In case of anti-DDK mouse monoclonal antibody (TA100011, do 1:4000 dilution).
7. Remove the blocking buffer and add enough diluted primary antibody to cover the membrane.
8. Incubate the membrane with primary antibody for 1hr at R/T. (Note: Or you can do overnight incubation at 4C, make sure you cover the western-blot tray to prevent excessive evaporation). To prevent uneven coverage, the western-blot tray can be rocked on a rocker platform.
9. Collect the primary antibody and store them at 4C for up to two weeks. (If you would like to store them longer, you can freeze the diluted antibody at –20C. Remember frequent freezing and thawing will gradually decrease the antibody titer.)
10. Briefly wash the membrane with 1xPBST once to remove any excessive primary antibody.
11. Add enough 1xPBST to cover the membrane and leave the Western-blotting tray on a rocker platform.
12. Wash the membrane for 15 minutes. (Note: If the background is high, repeat this step for two to three times.), turn on the developer during the wash time.
13. Dilute HRP-conjugated secondary antibody with blocking buffer (1:5000 or higher dilution is usually good for Goat anti-mouse-HRP; TA100015).
14. Incubate the membrane with secondary antibody for 30 minutes to 1hr.
15. Wash the membrane with 1xPBST for 15 minutes, and then 3 times (5 min/time).
16. Prepare the chemiluminescence development substrate mixture by mixing equal amount of solution1 and 2 (TA100016; Normally 1ml will be enough for one membrane).
17. Prepare a plastic saran film, lay the film on a flat surface, and dispense 1ml of substrate mixture for one membrane on the plastic saran film.
18. Use a forceps to take washed the blot from the western-blotting tray, flip it, lay on the substrate mixture, and then incubate for 1 to 5 minutes. (Note: To avoid air bubbles, always lay the blot by touching one edge first.)
19. Remove excess Chemiluminescence Reagent and wrap the membrane in plastic. Place inside X-ray cassette.
20. Expose to film and develop

Buffer preparation

1xPBS: This buffer is made by dissolving 8g of NaCl, 0.2g of KCl, 1.44g of Na2HPO4 and 0.24g of KH2PO4 into 800ml of distilled water. Then adjust the pH to 7.4 with HCl, and add H2O to 1 liter.

1xPBST: 0.05% Tween 20 in 1xPBS

Reference

Sambrook, Fritsch, and Maniatis (1989) Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, volume 3, apendix B.12

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