Protocols

This page contains a bunch of specific protocols for RNA FISH. It is by no means comprehensive, and assumes some basic familiarity with the protocol as specified by Biosearch Technologies. The basic overview is fix, permeabilize, hybridize, wash, and image. This page gives protocols in specific situations that maybe differ somewhat from what's in the literature. The idea is NOT to necessarily provide super detailed step-by-step instructions, but to give some sense of things to watch out for. Note also that some of these protocols are from other people, and so I take no responsibility for any FISH disasters–peruse at your own risk! Just sayin'...

RNA FISH

Vanilla, i.e., mammalian cells

The original! Here's the latest version of what we used to do. First, fix for 10 minutes in 4% formaldehyde, 1x PBS. You can use paraformaldehyde if you want to, but formalin (37% formaldehyde) is fine. Wash 2x with 1x PBS. Then permeabilize in 70% ethanol overnight at 4C (or just 1 hour at room temperature). Then add 1mL wash buffer for 1-2 minutes. Then add 50µl (or so) of hybridization solution and cover with a coverslip (to spread the solution and prevent drying). Then put the sample in a sealed dish with a wet Kimwipe (to humidify) and put at 37C overnight. In the morning, add 1mL wash buffer and remove the coverslip. Aspirate and add more wash buffer, incubate at 37C for 30 minutes. Repeat the 1mL wash for another 30 minutes (adding some DAPI) and then add either 2x SSC or do the glucose oxidase routine and image!

Iterative hybridization (i.e., hybridization, strip, then rehybridization)

Written 6/22/16 Sydney M. Shaffer

Reagents needed:

• Stripping buffer: 60% formamide, 2X SSC
• 1X PBS

Start: begin with hybridized sample. Then:

1. Turn on heat plate and set to 37C.
2. Remove the current buffer from your sample and add 1mL (this is the volume for a 2-well chamber) of stripping buffer containing 60% formamide and 2X SSC.
3. Place the sample on the heat plate and incubate for 15 minutes.
4. Remove the stripping buffer and replace with 1mL of PBS.
5. Remove the PBS and replace it with another 1mL of PBS.
6. Place the sample on the heat plate and incubate for 15 minutes.
7. Repeat steps 5-6 two more times (for a total of three 15 minute incubations with PBS).
8. Remove the PBS and wash with 1mL of wash buffer.
9. Set up your next RNA FISH!

Yeast

This comes courtesy of Lenny Teytelman, all around cool guy and yeast FISHerman extraordinaire. All experiments done using CSM media/plates (supposedly more reproducible results than YPD).

Fixation

Want the cells continuously in exponential phase, never reaching stationary, so do not do overnight inoculation as in the transformation protocol.

1. Around 10am, start a cell culture in a 50ml tube, using 10ml of CSM.
2. Grow for 8-10 hours in a shaker at 30C.
3. Measure OD in the evening and dilute into 250ml glass flasks, starting 45ml of CSM for overnight growth. Aiming for OD 0.2-0.4 around 10am the next morning. (Better to dilute more and wait till the next morning than to over-grow; don’t want to fix at OD>0.5.)
4. Transfer to 50ml falcon tubes.
5. Add 5ml of Formaldehyde, invert a few times, set at benchtop for 45min.
6. (Optional, NOT recommended per Anne Dodson, Marc Sherman Lenny Teytelman) - Transfer to gentle rocking overnight at 4C for 18-24 hours.
7. Spin down at 3,000rpm for 4min.
8. Decant and wash by pipetting with 1ml of ice-cold Buffer B. Transfer to 1.5ml tubes.
9. Spin down and wash once more with ice-cold Buffer B.
10. Resuspend in 1ml of Buffer B, add 2.5ul of Zymolyase and digest at 30C until most of the cells turn dark when checked by phase contrast microscope.
    1. NOTE (per Dan Pollard): if you're not sure what zymolyase treatment looks like, do a 10X positive control for 3 hours. Marc Sherman adds: if the positive control cells don't look markedly different (essentially black/opaque) from untreated control, try zymolyase treatment immediately after fixation (i.e., skip step 6).
11. Wash twice with ice-cold Buffer B, spinning 5-6min at 2,000rpm.
12. Resuspend in 1ml of 70% Ethanol overnight at 4C.

NOTE: * Zymolyase digestion usually takes 45-90 minutes, depending on the cell concentration.

NOTE: * Do not spin at 3K rpm after digestion with zymolyase - cells burst and deform because of the digested cell wall.

Hybridization

- Spin down 300ul of fixed cells in ethanol, aspirate away the ethanol completely

- Add 50ul of 10% Formamide hybridization buffer and 1ul of the probe to each sample. Hybridize at 30C overnight.

- In the morning, add 100ul of 10% Wash Buffer, spin at 2,000 for 5 minutes, aspirate carefully.

- Add 1ml Wash Buffer and incubate for 30 minutes at 30C.

- Spin, aspirate the Wash Buffer and add 1ml of Wash Buffer with 1ul of DAPI

Imaging

- Spin down, aspirate, and add 10ul of Glox Buffer without enzymes (resuspend).

- Keep all samples at 4C.

- Prepare 5x stock of Glox Buffer + enzymes (5ul of catalase, 5ul of Glucose Oxidase, 5ul of 200uM Trolox).

- Add 10ul of the 3X Glox Buffer + enzymes to sample, directly before the imaging.

- Apply 2ul of the cells to a glass slide, gently lower a coverslip onto a glass slide

- Put a kimwipe on top of the slide, place between 3-4 paper towels, and apply gentle pressure to the coverslip.

* First time, trying different probe concentrations from the original TE suspension (1:10,1:20,1:50,1:100).

* If making a master mix, hybridization buffer is very viscous -- if need 200ul of mix, make 300ul total.

* After O/N hybridization, easy to lose cells in aspiration after the first spin. Okay to leave 10-20ul of liquid. Also in subsequent washes, don’t be aggressive in aspirating.

Thanks Lenny!