Protocols

Protocols

Overview

On this page, we describe the general procedure for coupling and purifying the probes, fixing various types of samples, performing the in situ hybridization itself, and imaging the samples on a fluorescence microscope.

For the design of the probes, please visit this page on our website where we give a thorough description of the important points and where we have a program to assist in the designing of the probes themselves. In each section, we give a list of reagents and equipment required, followed by the procedure itself.

Critical points and warnings will be identified as follows:

Watch out!

Coupling and purification

Reagents and equipment

For coupling:

DMSO
0.1M Sodium Bicarbonate (made with RNase free water)
1M Sodium Bicarbonate (made with RNase free water)
Ethanol (95% pure is fine)
3M Sodium Acetate, pH 5.2
Fluorophore with succinimidyl ester group

For purification:

0.1M Triethylammonium acetate for HPLC (Buffer A)
Acetonitrile for HPLC (Buffer B)
HPLC with a dual wavelength detector to measure both DNA and fluorophore absorbtion
C18 Column for HPLC, 218TP104
Speedvac rated for acetonitrile

Procedure

Coupling
  1. Order oligonucleotides from a company (e.g. BioSearch Technologies) with 3' amine groups at the smallest possible synthesis scale.
  2. Pipette around 1nmole of each oligo into a single microcentrifuge tube (this is 10µl of a 100µM stock).
  3. Add 0.11 volumes 1M sodium bicarbonate to give a final concentration of 0.1M sodium bicarbonate. If the total volume at this stage is < 0.3mL, add some 0.1M sodium bicarbonate to bring total volume to 0.3mL.
  4. Dissolve a small amount (roughly 0.2mg) of dye into 50µl of 0.1M sodium bicarbonate.
    • Note: TMR can be hard to dissolve in aqueous solution, so one must first dissolve it in a small volume (<5µl) of DMSO before adding 50µl of 0.1M sodium bicarbonate.
  5. Add the dissolved fluorophore to the oligos.
  6. Cover the tube in foil to prevent photobleaching and let the reaction proceed overnight at room temperature with gentle rocking.
  7. In the morning, precipitate the oligos by adding 10% volume/volume of sodium acetate and then adding 2.5 volumes of 100% EtOH. Store this at –70C for at least 1 hour (up to overnight).
  8. Spin down the sample in a 4C microcentrifuge for at least 15 minutes at maximum speed (~16K RCF).
  9. After centrifugation, one should see a small colored pellet at the bottom of the tube. Aspirate away the fluorescent supernatant (containing uncoupled dye molecules) as completely as possible.

    10. The pellet is stable and can be stored at -20C.

Purification
  1. Resuspend pellet in appropriate volume (0.1->0.5mL nuclease free water, depending on your HPLC)
  2. Inject coupled probe into HPLC and run a program in which the percentage of buffer A varies from 7% to 30% over the course of around 30 minutes.
    • Set the detector to monitor DNA absorption (260nm) and the absorption of the coupled fluorophore (e.g., 555nm for TMR).
  3. One will observe two broad peaks. The first contains uncoupled probes and will only show a peak in the 260nm channel. The second contains pure coupled probes and will show peaks in both channels. Collect this entire peak in a series of microcentrifuge tubes.

    4. Be sure to collect the entire fraction and not just the "peak".

  4. Dry the purified probes in a speedvac (~ 3-5 hours for 0.5mL).
    • Be sure to prevent any light from hitting the probes during the drying process to prevent photobleaching, especially for photolabile dyes such as Cy5.
  5. Resuspend all tubes together in a total of 100µl of TE, pH 8.0. This is the probe stock.
  6. Dilute this probe 1:10, 1:20, 1:50 and 1:100 in TE to make working stocks for testing probe concentration.

    7. All done! The probes can be stored at -20C for years.

Fixation

Reagents and equipment

For all samples:

4% formaldehyde, 1xPBS (mix 37% formaldehyde/100% formalin in RNase free water 1:10)
1x PBS (made with RNase free water)
70% Ethanol (mix 100% ethanol with RNase free water)

For yeast:

Buffer B (from Singerlab protocols)
Spheroplasting buffer (10mL Buffer B + 100µl of 200mM vanadyl ribonucleoside complex)
Zymolyase
Concanavalin A

Formaldehyde is a known carcinogen and should be used in a chemical fume hood.

Procedure

Adherent mammalian cell lines

This protocol is adapted from the Singer lab's protocol.

  1. Grow cells in Lab-Tek chambered coverglass (with #1 coverglass on the bottom).
  2. Aspirate growth medium.
  3. Wash with 1x PBS.
  4. Add fixation solution and incubate at room temperature for 10 minutes.
  5. Wash 2x with 1xPBS.
  6. Add 70% EtOH and store at 4C at least overnight and up to 1 week.

    7. All done! Cells can be hybridized up to a week after being fixed, perhaps longer.

C. elegans larvae
  1. Grown larvae in a plate seeded with OP50.
  2. Add 5mL M9 buffer and swirl in plate to release worms from surface, then move worms to a 15mL conical centrifuge tube.
  3. Spin down worms and aspirate.
  4. Wash with 5mL M9 buffer.
  5. Spin down worms and aspirate.
  6. Add 1mL fixation solution, transfer to microcentrifuge tube, and incubate for 45 minutes.
  7. Wash 2x with 1mL 1x PBS.
  8. Resuspend in 1mL 70% EtOH and leave at least overnight at 4C.

    9. All done! Larvae can be hybridized up to a week after being fixed, perhaps longer.

C. elegans embryos
  1. Add 5mL M9 buffer to a plate of gravid hermaphrodites and swirl to release worms from surface. Move worms to a 15mL conical centrifuge tube.
  2. Spin down and add bleaching solution.
  3. Vortex for roughly 4-8 minutes until worms disappear and only embryos remain.
  4. Spin down and aspirate, then wash 2x in M9 buffer.
  5. Resuspend in 1mL fixation solution and incubate at room temperature for 15 minutes.
  6. Vortex and then immediately submerge tube in liquid nitrogen for 1 minute to freeze crack the embryos’ eggshells.
  7. Thaw in water at room temperature.
  8. Once thawed, vortex and place on ice for 20 minutes.
  9. Wash 2x with 1mL 1x PBS.
  10. Resuspend in 1mL 70% EtOH and store at least overnight at 4C.

    11. All done! Embryos can be hybridized up to a week after being fixed, perhaps longer.

D. melanogaster wing imaginal discs
  1. Submerge 3rd instar larvae in 1mL 1x PBS and dissect to release wing imaginal discs.
  2. Place discs at the bottom of a chambered coverglass. They should stick readily.
  3. Fix wing discs by aspirating PBS and adding 1mL fixation solution; incubate at room temperature for 45 minutes.
  4. Wash 2x with 1mL 1x PBS to remove fixative.
  5. Add 1mL 70% EtOH and leave at least overnight at 4C.

    6. All done! Discs can be hybridized up to a week after being fixed, perhaps longer.

Yeast (S. cerevisae)

This protocol is adapted from the Singer lab's protocol.

  1. Grow yeast to an OD of around 0.1-0.2 in a 45mL volume of minimal media.
  2. Add 5mL of 37% formaldehyde directly to growth media and let sit for 45 minutes.
  3. Wash 2x with ice cold Buffer B.
  4. Add 1mL of spheroplasting buffer, transferring to a new microcentrifuge tube.
  5. Add 1µl of zymolyase and incubate at 30C for 15 minutes.
  6. Wash 2x with ice cold Buffer B, spinning at low speed (~2000 rpm).
  7. Add 1mL 70% EtOH and leave overnight at 4C.

    8. All done! Yeast can be hybridized up to a week after being fixed, perhaps longer.

Hybridization

One of the key parameters in hybridization is the concentration of formamide used for the hybridization and washing steps. This effectively sets the stringency, with higher formamide concentration being more stringent. We have generally found that 10% formamide works well in most cases, but if your mRNA target has high GC content, you may want to increase this to 20% or higher. The recipes given here are based on those from the Singer lab.

Reagents and equipment

Hybridization buffer (10mL):

Dextran sulfate (1g)
E. coli tRNA (10mg)
Vanadyl ribonucleoside complex (NEB) (100µl of 200mM stock)
BSA (RNase free) (Ambion) (40µl of 5mg/mL)
20X SSC (nuclease free, Ambion) (1mL)
Formamide (deionized, Ambion) (1mL for 10% final concentration)
Nuclease free water (Ambion) (to 10mL final volume)

Wash buffer (50mL):

20X SSC (Ambion) (5mL)
Formamide (deionized, Ambion) (5mL for 10% final concentration)
Nuclease free water (Ambion) (to 50mL final volume)

Formamide is a teratogen that is easily absorbed through the skin and should be used in a chemical fume hood.

Be sure to let the formamide warm to room temperature before opening the bottle.

Anti-bleach buffer and enzymes:

10% glucose in nuclease free water
2M Tris CL, pH 8.0
20X SSC (Ambion)
Nuclease free water (Ambion)
Glucose oxidase (Sigma) (diluted to 3.7mg/mL stock)
Catalase (Sigma)

Mix together 0.85mL of NF water and add 100µl of 20x SSC, 40µl of 10% glucose and 5µl of 2M Tris CL. Vortex and then transfer 100µl of this "glox" buffer to another tube, to which one should add 1µl of glucose oxidase stock and 1µl of (nicely vortexed) catalase suspension. The remainder will be used as an equilibration buffer.

Procedure

Hybridization in solution
  1. Prepare the hybridization solution: to 100µl of hybridization buffer, add 1-3µl of probe at the appropriate concentration, then vortex and centrifuge.
    • Be sure to warm the hybridization solution to room temperature before opening it.
    • For the initial test of a set of probes, it is best to start 4 separate hybridization reactions by adding 1µl each of the 1:10, 1:20, 1:50 and 1:100 working dilutions of probes to see which one is optimal.
  2. Centrifuge the fixed sample and aspirate away the ethanol.
  3. Resuspend in 1mL wash buffer that contains the same percentage formamide as the hybridization buffer you will be using. Let stand for 2-5 minutes.
  4. Centrifuge sample and aspirate wash buffer, then add hybridization solution. Incubate in the dark overnight at 30C.
  5. In the morning, add 1mL of wash buffer to the sample, vortex, centrifuge and aspirate, then resuspend in another 1mL of wash buffer and incubate at 30C for 30 minutes.
  6. Vortex, centrifuge and aspirate the wash buffer, then resuspend in another 1mL of wash buffer containing 5ng/mL DAPI for nuclear counterstaining. Incubate at 30C for 30 minutes.
  7. If you are imaging without using glucose oxidase (glox) anti-fade solution (e.g., if you used TMR), then just resuspend in an appropriate volume (> 0.1mL) of 2x SSC and proceed to imaging.
  8. If you are imaging with the glucose oxidase (glox) anti-fade solution, aspirate the buffer and resuspend in the glox buffer without enzymes for equilibration; incubate for 1-2 minutes.
  9. Aspirate the buffer and resuspend in the 100µl of glox buffer to which the enzymes (glucose oxidase and catalase) have been added.
  10. Proceed to imaging

    11. The sample (either with or without anti-bleach solution) can be kept at 4C for a day's worth of imaging.

Hybridization for samples adhered to coverglass
  1. Prepare the hybridization solution: to 100µl of hybridization buffer, add 1-3µl of probe at the appropriate concentration, then vortex and centrifuge.
    • Be sure to warm the hybridization solution to room temperature before opening it.
    • For the initial test of a set of probes, it is best to start 4 separate hybridization reactions by adding 1µl each of the 1:10, 1:20, 1:50 and 1:100 working dilutions of probes to see which one is optimal.
  2. Aspirate the 70% ethanol off of the sample.
  3. Add 1mL wash buffer that contains the same percentage formamide as the hybridization buffer you will be using. Let stand for 2-5 minutes.
  4. Aspirate wash buffer and then add hybridization solution. Place a carefully cleaned coverslip over the sample to prevent drying of the hybridization solution during the incubation. Incubate in the dark overnight at 30C.
  5. In the morning, add 1mL of wash buffer to the sample, remove the coverslip, then incubate at 30C for 30 minutes.

    Be sure to remove the coverslip very carefully so as not to disturb the cells underneath very much.

  6. Aspirate the wash buffer, then resuspend in another 1mL of wash buffer containing 5ng/mL DAPI for nuclear counterstaining. Incubate at 30C for 30 minutes.
  7. If you are imaging without using glucose oxidase (glox) anti-fade solution (e.g., if you used TMR), then just add 1mL of 2x SSC and proceed to imaging.
  8. If you are imaging with the glucose oxidase (glox) anti-fade solution, aspirate the buffer and resuspend in 1x PBS
  9. Aspirate the PBS and add the glox buffer without enzymes for equilibration; incubate for 1-2 minutes.
  10. Aspirate the buffer and resuspend in the 100µl of glox buffer to which the enzymes (glucose oxidase and catalase) have been added.
  11. Place a carefully cleaned coverslip over the sample. This will spread the glox buffer over the entire sample and also slow evaporation.
  12. Proceed to imaging

    13. The sample (either with or without anti-bleach solution) can be kept at 4C for a day's worth of imaging.

Imaging

Microscopy equipment

Standard fluorescence microscope (e.g., Nikon TE2000 or equivalent)
Strong light source, such as a mercury or metal-halide lamp
Filter sets appropriate for the fluorophores chosen
Standard microscopy grade camera, ideally optimized for low-light level imaging rather than speed (13µm pixel size or less is ideal)
Nice 100x DIC objective (with an IR coating if you are using dyes like Cy5)

Tips for imaging

In general, the idea is simple: mount your samples and image them. There are, however, a number of potential issues that might cause you trouble. Here are some tips:

  • Be sure that the sample is as close to the coverglass (and hence objective) as possible (this is, of course, automatically the case with adherent cells). We have found that the particles become fuzzier the further they are from the coverglass. Also, we have not had good luck using coverglasses thicker than #1, but maybe you will have better luck.
  • Do not use commercial anti-fade mounting solutions. We found that they introduce a large background while dramatically decreasing fluorescence.
  • Do not seal the sample with nail polish. It causes a background autofluorescence in the red channels that can mask the mRNA signal.

Troubleshooting

In general, there are three important parameters that one can vary if the particular mRNA signal is not optimal: the concentration of probes used, the concentration of formamide used in the hybridization and washes, the temperature at which the hybridization is performed. If the probe concentration is too high, the background will be too large to yield good signals, whereas if the probe concentration is too low, the signals will be faint or undetectable. Optimal results can be obtained over a fairly large range of probe concentrations, though, and it is likely that one of the dilutions used in the initial pilot experiments will be acceptable. Increasing the formamide concentration increases the stringency of the hybridization and washing steps. This is particularly useful if the GC content of the mRNA is relatively high: we have found that 10% formamide is generally optimal when the GC content is between 40% and 50%, but if a large background is detected, then one can increase the formamide concentration up to 20%, which seems to work sometimes with probes whose GC contents range between 50% and 60%. Increasing temperature also increases stringency, although it can be hard to rigorously control temperatures through experiments and so we typically do not change that particular parameter.