Tissue Handling: Note that all unfixed human tissue should be handled as BioSafety Level 2 materials (wear gloves, lab coat, etc). All plasticware (tubes, tips, etc.) should be RNase free and always handled with gloves to avoid contamination of the sample with RNases. In general, it is not necessary to treat plastic ware to eliminate RNase contamination- just keep separate, unopened stocks of tubes and tips for autoclaving that are always handled while wearing gloves. Similarly, all chemicals used for RNA should be dedicated for that purpose and handled with gloves to avoid contamination.

 

 

 

1.      Prepare a scintillation vial (Fisher #3-341-13) for each sample containing 1.5 ml of Trizol (Invitrogen#15596-026). Exercise proper laboratory practice when handling Trizol, as it is toxic and corrosive (i.e., wear gloves, lab coat, eye protection). Also label a 14 ml round bottomed polypropylene centrifuge tube (Falcon #2059) for each sample.

 

 

2.      In a cryostat at -20° C, cut the desired number of sections and place in the scintillation vial containing Trizol. The scintillation vial is used because the large opening allows the sections to be placed directly in Trizol to prevent degradation of the RNA and sticking of the sections to the sides of the tube. The number of sections to be cut can be estimated as follows:

 

 

Pre-cut a thin section (5 m m) and H&E stain it to determine if the sample contains sufficient tumor cells (we prefer at least 70% tumor cells). Measure the length and width of the section. The number of sections needed is then estimated as follows:

 

 

Length (mm) ´ width (mm) ´ thickness of sections (mm) ´ # sections = X

where X has a value between  and 20 and 100 mm³

 

 

 

For example, a 4´ 5 mm section would require a minimum of 20 50m m sections (4´ 5´ 0.05´ 20= 20 mm³ )

 

 

Sometimes it is not possible to cut the minimum number of sections to meet the above criteria. If this is the case, cut as many sections as possible making sure to leave enough tissue to take an “AFTER” section for H&E staining to verify the tumor cell content.

 

 

3.      Homogenize the tissue for 30 seconds in the scintillation vial so that no large pieces of tissue are evident. Perform the homogenization in a Biosafety cabinet to avoid exposure to aerosolized tissue, with the vial in a beaker filled with ice to keep the sample cold.

 

 

4.      Transfer the homogenate to the Falcon 2059 tube that is in an ice bucket. Rinse the scintillation vial out with 0.5 mL of Trizol in order to get any residual tissue and add to the Falcon tube. Dispose of the scintillation vial in chemical waste.

 

 

5.      Homogenize for 2 minutes, keeping the tube in a small beaker filled with ice to prevent overheating. Homogenization in this tube gives a smaller surface area, allowing more efficient homogenization. Once homogenized, the tissue can be stored at -80° C for at least one month prior to extraction.

 

 

6.        Transfer the Trizol to two 1.5ml microfuge tubes (1 ml in each).

 

 

7.      Add 200 m l of chloroform for each ml of Trizol.

 

 

8.      Vortex for ~10 seconds to mix the Trizol-chloroform.

 

 

9.      Spin at 12,000´ g at 4° C for 15 minutes in a microfuge.

 

 

10.  Transfer the clear aqueous upper phase to a fresh microfuge tube, being careful to avoid the white interphase, which contains DNA and proteins. The remaining Trizol can be saved for extraction of DNA or disposed of in chemical waste.

 

 

11.  Add 2.5 m l of 5mg/ml linear acrylamide (Ambion #9250) to the extracted aqueous solution (final working concentration should be in the 10-20 m g/ml range). Linear acrylamide acts as a coprecipitant to aid in precipitating small amounts of RNA. Alternatively, glycogen can be used as a coprecipitant (final working concentration of 50-150 m g/ml- available from Ambion as 5mg/ml #9510).

 

 

12.  Add 0.9 vol of isopropanol (typically 540 m l of isopropanol added to 600 m l of aqueous solution).

 

 

13.  Precipitate RNA for at least 1 hour (up to overnight) in a -20° C freezer.

 

 

14.  Pellet the RNA by spinning at 12,000´ g at 4° C for 15 minutes in a microfuge.

 

 

15.  Remove supernatant and rinse with 500 m l of ice cold 70% ethanol.

 

 

16.  Spin down in microfuge at 7,500´ g at 4° C for 5 minutes.

 

 

17.  Quickly remove supernatant. Failure to remove the 70% ethanol quickly can result in breakup of the pellet and potential loss of RNA while removing the supernatant. Respin the tube as in step 17 if you see evidence of the pellet fragmenting.

 

 

18.  Air-dry the pellet. Since there are two tubes, it is a good idea to dissolve one pellet in formamide for long-term storage, and the other in nuclease free water for immediate use. Typically we dissolve the RNA from sections in 30-50 m l of water or formamide. When the pellet dries, it will become clear (it will be white in color prior to this). When using formamide, dissolve the pellet immediately after it becomes clear. If the pellet dries out more, it will become very difficult to dissolve. To recover the RNA stored in formamide, add NaCl to a final concentration of 0.2 M, add 4 volumes of 100% ethanol, incubate 5 minutes at room temperature, then spin for 5 minutes at 10,000´ g at 4° C in a microfuge. Rinse the pellet with 70% ethanol, spin 5 minutes at 7500´ g at 4° C, and air dry pellet. When dry, dissolve in nuclease free water.

 

 

19.  Take a small aliquot of the RNA solution for measurement by UV spectrophotometry (typically a 1 in 25 or 1 in 50 dilution in 1´ TNE). Make sure that the 260/280 ratio is at least 1.7 (high quality RNA gives a ratio of 2.0) otherwise the concentration readings will not be accurate. Re-extract the RNA if the ratio is below 1.7.

 

 

20.  Calculate the amount required to run 100ng on a standard 1% formamide agarose gel. For 100 ml gel volumes, add 1 g of agarose to 87 ml of depc-treated water and boil until dissolved. Cool to 60° C, then add 10 ml of 10´ MOPS and 1.6 ml of formaldehyde, mix well, and pour into casting tray. Use 1´ MOPS as the running buffer. Mix the RNA with water and 2.0 ul of RNA loading buffer (Sigma, R4268) so that the total volume is 8 m l, then heat the sample to 65° C for 10 minutes. The RNA loading buffer contains ethidium bromide, so use caution when handling this solution. Transfer the sample to ice for 5 minutes then load into the gel. It is also useful to include a size marker (Promega, G3191) for size comparisons. Run until the loading dyes have separated well, then photograph using UV illumination source. The RNA should give sharp 28S (~4.7 kb) and 18S (~1.86 kb) ribosomal bands, with the 28S band appearing denser. The 5S band should also be visible, and with very good quality RNA, should appear as two bands (tRNA and 5S ribosomal RNA). Note that samples dissolved in formamide will migrate at a slightly slower rate than the samples dissolved in water.