Fluorescent in situ hybridization for mRNA detection in yeast
Last modified by T.L. To on Feb 17, 2009
For additional information, see Arjun Raj's website
- 1 Reagents
- 2 Fixation
- 3 Hybridization
- 4 Washing and Imaging
- 5 Probes preparation
Start with 300mL RNase free water in a bottle (freshly autoclaved) Add 218g sorbitol (MW 182.17) 17.4g Potassium Phosphate (dibasic) Mix this until it dissolves a bit, then add RNase free water up to 1L. Store at 4 degC
- 10mL Buffer B
- 100μl 200mM vanadyl ribonucleoside complex
- 20μl beta-mercaptoethanol (optional)
- Store at 4 degC
For 10 mL -
- 1mL formamide to 10% (Ambion)
- 10mg E. coli tRNA (=1μg/1μl * 10000μl) (Sigma, 83854)
- 1mL 20x SSC (RNase free, from Ambion)
- 40μl of RNase free BSA (stock is 50mg/mL = 5% solution from Ambion)
- 1g Dextran Sulfate (for 10% in 10mL)
- 100μl of 200mM vanadyl-ribonucleoside complex (NEB S1402S)
- Appropriate amount of nuclease free H2O
First add some of the water and the dextran sulfate to a 15mL tube and vortex to mix. Once dissolved, add in the rest of the ingredients and add water up to the final volume of 10mL. Before using, be sure to filter through small filters from Ambion. Store 50 uL aliquots at -20 degC.
NOTE: USE FILTER TIPS and wear gloves for all procedures to minimize contamination with RNases.
- Grow cells in rich or synthetic medium. Harvest cells at mid log-phase (OD 0.5) - take 10mL of culture and put it in a 15mL falcon tube (plug seal is preferred).
- Add the appropriate amount of formaldehyde to ~4% final formaldehyde (e.g., add 1.1mL of 37% formaldehyde to the 10mL of culture). Note that the formaldehyde stock can degrade over time.
- Let sit for 40 minutes at room temperature.
- Spin down for 3 minutes at 3000g.
- Aspirate fixative and resuspend in 10-15mL cold Buffer B, then spin down as before.
- Aspirate Buffer B, resuspend again in Buffer B and then spin down again (this washing procedure should remove all the remaining fixative from the solution).
- (Spheroplasting) Add 1mL spheroplasting buffer to 50mL tube, resuspend by pipetting and transfer to a 1.5mL eppendorf tube. (Optional: sonicate the cells at this point)
- Add 1-2 µl of lyticase (20000-25000U/ml, dissolved in 50mM Tris-HCl 7.5pH, 20% glycerol, 5mM beta-mercaptoethanol).
- Mix by pipetting and incubate at 30C for 15 minutes.
- Spin down at 3000rpm for 2 minutes.
- Aspirate supernatant and wash with 1mL Buffer B.
- Spin down at 3000rpm for 2 minutes.
- Aspirate supernatant and wash with 1mL Buffer B.
- Remove supernatant.
- Add 1mL of 70% EtOH.
- Let sit for at least 1 hour at room temperature before performing in situ hybridization. Alternately, one can leave the cells in EtOH at 4C for up to a week. Be sure to carefully seal the chamber or petri dish with parafilm, since the EtOH can quickly evaporate away and ruin the sample.
- Warm up the hybridization solution to room temperature
- Decant ethanol and wash cells once with washing buffer (10% formamide, 2xSSC). Formamide is toxic so fume hood is needed (also, high quality deionized formamide should be used – can be ordered from Ambion). Cells can be vortexed but centrifuging steps must be gentle (< 2000 rpm)
- Re-suspend cells in 40 uL hybridization solution
- Add probes (they are photosensitive and should be kept away from light). The concentration of probes in the final hybridization mixture should be ~ 0.1-1ng/mL. The probes are difficult to quantify accurately so the concentration is only estimated by eyeballing
- Incubate at room temperature for at least 8 hours. Keep in dark
Washing and Imaging
- Spin, decant hybridization solution, and wash with 1mL 10% formamide 2xSSC. Incubate at 30 degC for half an hour
- In the meantime, use poly-lysine to coat the coverglass for imaging. Make 1mg/mL poly-lysine stock solution, aliquot and store at -20 degC. Add about 200-300 uL to each coverglass and let it sit for 30 minutes. Pipet to remove poly-lysine but leave a thin layer. (Note: use tweezers to handle coverglass to avoid RNAse)
- After the first wash (30 min), spin, decant, and resuspend in 1mL 10% formamide 2xSSC, (Optional) add 1 uL of DAPI (5mg/mL) to mark the nucleus. Incubate at 30 degC for half an hour
- After the second wash (30 min), spin, decant and resuspend in 1mL 2xSSC (no formamide). 2xSSC (at right pH) maintains hybridization (once probes come off they won’t rebind)
- Usually, can wait after the third wash before imaging. However, it’s better to do imaging a.s.a.p. if there are unknown parameters (mismatches, not enough probes, etc)
- Imaging – simply take Z-stacks. Need 100x objective, appropriate filter set, and mercury lamp (Xenon lamp does not have enough power)
- For TMR, use chroma filters D546/10x and D610/60, exposure time of 3000 ms, and EM gain of 3500 (this acquisition setting is saved as “Rhodamine” in the Metamorph software).
- Poly-lysine is from Sigma (Product no. P1274). The final concentration is 1 mg/mL (0.1%). Store small aliquots (~1mL) at -20 degC.
- Order probes from Biosearch Technologies Inc. via Joanne Giffra [email@example.com]. Let her know you are referred by Arjun Raj
- Fill out the Custom Oligonucleotide Order Form.
- Plate orders need to be a minimum of 48 oligos but can be any number above that
- Prepare 1M and 0.1M sodium bicarbonate solutions. Also, add 1M NaHCO3 to the probes to bring [NaHCO3] to 0.1M
- TMR (tetramethylrhodamine) dye is not water soluble and has to be dissolved in DMSO. Mix a few uL of TMR dye with about 10uL of DMSO. Color change should occur (solution turns pink). This amount of dye should be good for ~10 coupling reactions.
- Add 0.1M NaHCO3 (36 uL per sample) to the dye, mix by pipetting.
- Add 36 uL of dye-NaHCO3 mix to each sample.
- Incubate at 37 degC for overnight in dark.
- The oligo probes need to be precipitated prior to purification. Precipitation can be done using the standard protocol (sodium acetate, 100% EtOH, -80degC for an hour)
- Spin down pellets. A nicely coupled pellet should show uniform color.
- Discard supernatant. No washing step is needed. The fraction of unlabelled dye is negligible and it can be picked up by HPLC.
- Bring microtubes, pipet, filter tips, gloves, freezer box, and lab markers to the HPLC facility
- HPLC facility is located at the Biopolymer Lab at E17-FL4. The main unit is Agilent 1100/1200 Flow Series (Diode Array and Multiple-wavelength Detector) G1315 C/D, G1365 C/D. The right column is VyDAC Catalog # 218TP104, protein and peptide C18. The automatic collector is Gilson FC203B. It knows how to tell jokes.
Setting up the HPLC
Contact Richard Cook (Dick) cook@MIT.EDU to set up appointments
- If buffers need to be swapped, wash the lines with water (flow-rate = 4mL/min) and replace the buffer bottle. For FISH, Triethylammonium acetate (Buffer A) and acetonitrile (Buffer B) are used. Buffer A is at pH 7.0, which facilitates bacterial growth and thus resulting in pH changes (which affect the heavily charged oligos). Buffer A is also mildly volatile. No desalting is needed if it runs out.
- Clean out all lines with Buffers A and B with 4mL/min flows for 5 minutes. In the meantime, change the column. The flow should be directed to the waste at this point.
- Turn the black knob clockwise until the second catch to run the column.
- To equilibrate signals (the traces on the signal plots), run the HPLC at 1 mL/min for about 15min. Use 50% Buffer A and 50% Buffer B (what flows through the column).
- The pressure should become 50 bar when Buffer A is 93% and Buffer B is 7%
Important note: TURN OFF the flow when swapping buffers to avoid air from getting into the system.
Samples preparation for HPLC
- Take sample tubes and caps.
- Resuspend the probes (in pellet form) with at least 113uL of nuclease free water. The HPLC will suck up 100uL but extra volume is needed (avoid sucking up air)
- Filter if there are clumps in the sample.
- When pipetting the sample, one should make sure there is no air bubble (extremely important as bubbles can totally kill the signals). Suck out any air bubbles by pipetting.
- Put on the cap, and squeeze it tightly with the squeezer (which is sitting next to the HPLC).
- Put the sample into the plate located inside the HPLC. Jot down the position (e.g. D03).
Setting up the HPLC software
- Make sure the method has been loaded: >Instrument>Load method and Arjun’s method is stored at C:\Chem32\1\METHOD\OLIGO_7_25_55_ARJUN.M
- First, go to >Sequence>Sequence parameter
- Change ‘Subdirectory’ to date_details – e.g. 7_17_08_med_tmr
- Press Ok to create the directory
- Second, go to >Sequence>Sequence table
- Change the location (to the position of the sample tube)
- Change the sample name (e.g. med_tmr)
- Double check the method name (which should be OLIGO_7_25_55_ARJUN)
- Hit OK
- Third, go to >Sequence>Save Sequence As
- Type in the name of the file (e.g. 7_17_08_med_tmr.S)
- Hit start and save changes
Running the HPLC
- Click start
- Run at 1mL/min. Initially, Buffer A is 93% and Buffer B is 7%
- There will be no signals during the first 3-4 minutes (before solvent and solutes can be detected). After startup there might be fluctuations in signals. If a huge peak is seen then something is wrong. For a real signal it should read ~200 mAU
- Auto-suction machinery injects sample into the column
- The time axis of the signal plot is zeroed at the time of injection
- Signals to be looked at: DNA: 260 (reference off); TMR: 550; Alexa: 594; CY5: 650
- Optional: run a water blank, or run small fraction of the sample (diluted)
- Troubleshooting - Oscillatory background signals can be due to air bubbles. To get away with that, turn the black knob (so that the fluid flows are directed to the waste), and purge the lines by running the buffers at 5mL/min (max speed).
- Collect 500uL per tube. Start collecting when the right peak starts (not when it reaches maximum) and wait until the signal falls back to the baseline.
- Take anything from the outlet that is colored
- In Arjun’s program, Buffer B goes from 7% to 23%. The stripping step runs at 70% Buffer B
- After stripping Buffer B goes back to 7%, let it run at 7% Buffer B for at least 15minutes before the next sample.
- TMR is single isomer so there’s only 1 peak. For some other dyes (e.g. Alexa) there can be another peak at later times. This peak still represents coupled probes (possibly duplex). CY5 dominates hydrophobicity and has a long retention time (sharp elution peak)
Shut down the HPLC
- Run 100% Buffer B for 10 minutes to strip everything out.
- Go to >Instrument>Systems off
Drying the probes
- SpeedVac is available at 68-616C (Keating’s lab). Basically, it is a freeze drying machine. Solvent is first solidified (by refrigeration). Vacuum is then used to generate a very low temperature at which the solid directly sublimes to gas without turning into liquid. Freeze drying is a preferred method for preserving perishable materials and simplifying transports of materials
- Turn on the refrigeration to cool it down, and wait
- Turn on the pump, and turn on the valve to test if the vacuum line works properly. Turn off the valve after testing.
- Put the samples into the block (which looks like microcentrifuge’s)
- Turn on the rotor. Make sure ‘heating’ is off. When the rotor starts spinning turn on the vacuum pump. The lid should be sealed.
- Cover the lid with aluminum foil to avoid photo-bleaching
- Drying takes 3-4 hours.
- When drying is done, turn the valve of the pump to off position
- CAUTION: turn off the valve (or the pump) before turning off the rotor.
- Purge the vacuum in pump’s line
- Turn off the pump
Storage of probes
- Resuspend probes in about 75 uL of TE (from Ambion)
- Store probes at -20 degC
- Dilute ~20 fold to make working stock