Gary Drews, Tom Jack, Detlef Weigel, Hajime Sakai , Bobby Williams, Mark Running, Steve Jacobsen, and Beth Krizek.

Meyerowitz Lab


I. Fixation, Embedding, and Sectioning

II. Probe Synthesis

III. Hybridization

IV. Wash

V. Autoradiography and Staining

VI. Viewing and Photographing slides




Cox et al. (1984) Detection of mRNAs in sea urchin embryos by in situ hybridization using asymmetric RNA probes. Develop. Biol. 101, 485-502.

Cox and Goldberg (1988) Analysis of plant gene expression. In Plant Molecular Biology, A Practical Approach. C.H. Shaw, ed. (Oxford: IRL Press), pp 1-34.

Fink (1987) Some new methods for affixing sections to glass slides. I. Aqueous adhesives. Stain Technol. 62, 27-33.

Hayashi et al. (1978) Acetylation of chromosome squashes on Drosophila melanogaster decreases the background in autoradiographs from hybridization with [125I]-labeled RNA. J. Histochem. Cytochem. 26, 677-679.

Huang et al. (1983) Improved section adhesion for immunocytochemistry using high molecular weight polymers of L-lysine as a slide coating. Histochemistry 77, 275-279.

McFadden et al. (1988) A simple fixation and embedding method for use in hybridization histochemistry on plant tissues. Histochem J. 20, 575-586.

Meyerowitz (1987) In situ hybridization to RNA in plant tissue. Plant Molec. Biol. Rep. 5, 242-250

Moench et al. (1985) Efficiency of in situ hybridization as a function of probe size and fixation technique. J Virol. Methods 11, 119-130.




A. FAA Fixation:



Use bulk 95% or 100% ethanol except for the 100% ethanol steps.
For the 100% ethanol steps, use high quality ethanol (e.g. Gold Seal)
Glacial acetic acid
37% formaldehyde
20 ml glass scintillation vials (e.g. S/P Baxter #R2550-14)


1. Fixative Mix:

% In Mixture Amount
Ethanol 50.0% 50 ml
Acetic Acid 5.0% 5 ml
Formaldehyde 3.7% 10 ml of 37%
Water 41.3% 35 ml
100 ml

2. Place 10-15 ml of fixative into 20 ml scintillation vials.

3. Cut a cluster of flowers at the apex of the floral stem that includes stages 1-14, and immediately immerse in fixative. Cut so that 5 mm or so of floral stem is present--this helps to orient the tissue when sectioning. Cut off older flowers if you are not interested in these stages. Place about 20 pieces of tissue in each vial.

4. The tissue will float in the fixative.

5. Place tissue/fixative in a vacuum (with an aspirator) for 15 minutes. Pull the vacuum very slowly. This step pulls the air out of the tissue, allowing the fixative to penetrate better. Usually, air bubbles will come out of the solution and get trapped in the trichomes, causing the tissue to float.

6. After 15 minutes of slow bubbling, release the vacuum very slowly. Incubate at room temperature, up to a total of 1-2 hours exposure to the fixative, one hour is best for antibody in situ experiments and works fine for RNA in situ experiments.

7. Remove fixative and add 50% ethanol. Incubate at room temperature for 30 minutes. Repeat this step.


B. Dehydration:


The tissue must be completely dehydrated when it hits the xylenes; otherwise, the water and xylenes will form a white emulsion. We also stain the tissue with Eosin Y during these steps to help orient the tissue when sectioning.


Use bulk 95% or 100% ethanol except for the 100% ethanol steps.
For the 100% ethanol steps, use high quality ethanol (e.g. Gold Seal)
Xylenes (Fisher)
Eosin Y; Eosin Yellow (Manufacturing Chemists #B286 or Sigma)


1. Remove 50% ethanol and replace with 60% ethanol. Incubate for 30 minutes.

2. Repeat for the following ethanol solutions: 70%, 85%, 95%.

3. Leave overnight in 95% ethanol with 0.1% Eosin (Eosin should be stored wrapped in aluminum foil and should be discarded if no longer bright orange).

4. Next day, remove as much as possible of the Eosin/95% ethanol and replace with 100% ethanol. Incubate for 1 hour. The tissue destains slowly in 100% ethanol, so do not incubate too long.

5. Remove as much as possible of the 100% ethanol, replace with fresh 100% ethanol, and incubate for 30 minutes. Repeat this step if it is impossible to remove virtually all of the solution in steps 4 and 5.


C. Clearing:


The tissue must be permeated with xylenes because paraffin is not miscible in ethanol. The tissue does not destain (Eosin) in xylenes.



High quality ethanol (e.g. Gold Seal)


1. Remove 100% ethanol and replace with 25% xylenes:75% ethanol. Incubate at room temperature for 30 minutes.

2. Repeat for the following xylene:ethanol solutions:

50% xylenes:50% ethanol
75% xylenes:25% ethanol

3. Remove the 75% xylenes and replace with 100% xylenes. Incubate at room temperature for 1 hour.

4. Repeat step 3 two times. Fill the vial half full with xylenes the final time.


D. Infiltration:



Paraplast Plus Chips.
Molten Paraplast Plus. Place a beaker full of paraplast chips in an oven at 60oC well in advance; it will take >12 hours to melt.


1. Add 20 chips of paraplast to each vial. Incubate overnight at room temperature.

2. Next morning, the paraplast will be only partially into solution. Place vials in a 42oC incubator. After about 30 minutes, the paraplast chips will be in solution.

3. Add 20 more chips, and incubate at 42oC until chips are in solution--about 30 minutes. Swirl occasionally.

4. Repeat step 3 until the vial is full (4-5 times; total of about 100 chips).

5. Pour off xylene/paraplast solution. Add molten paraplast. Swirl to mix. Incubate at 59-62oC for at least 4 hours.

6. Repeat step 5 for a total of 6 changes over 3 days (i.e. change twice a day).


E. Pouring Boats:



Molten paraplast plus.
Index cards (3x5).
Variable temperature hot plate (Eberbach, Corp., Ann Arbor, MI)


1. Fold up an index card so that it has raised edges of about 1 cm. Tape the edges with Scotch tape outside.

2. Place the boat on the hottest part of the variable temperature hot plate.

3. Pour the infiltrated tissue into the boat.

4. Top off the boat with molten paraplast plus.

5. Arrange the tissue into a regular array. Orient tissue pieces with the stems either straight up or lying on their side. Tissue pieces must be at least 5 mm apart.

6. Slowly move the boat to the cooler part of the hot plate. You probably will need to arrange the tissue again.

7. Label each boat by inserting a flagged string.

8. Let harden completely. This will require overnight at room temperature. If in a hurry, you can float the boat for 2 hours (1 hour per side) in water.


F. Sectioning:



Poly-L-lysine-coated slides (Fisher 12-550-15).
TissuePrep Flotation Bath (e.g. Lab-Line # 26103/Baxter # M7655) set to 42oC .
Slide warmer (e.g. Fisher # 12-594) set to 45-50oC.
VWR single edged razors


1. Cut out blocks of embedded tissue and mount onto microtome blocks.

2. Section tissue at 8 um. If the wax is good quality, 5-6 um sections can be made, which will give more tissue per sample.

3. Cut ribbons into 1.5 cm pieces. Float ribbon pieces on 42oC water for >1 minute. This step takes the compression out of the tissue.

4. Put slide in water just under the floating ribbon.

5. Bring slide up so as to catch the ribbon (don't put sections to close to any edge of the slide). Use a wooden applicator to position the ribbon. Label the slide with a pencil (ink dissolves in the ethanol/xylene steps).

6. Repeat for as many different ribbon pieces as you want to place on a given slide, frame the sections so that they will fit under the 24 X 50 mm cover slips.

7. Incubate slide on a slide warmer overnight to "bake" the ribbon piece onto the slide. This is a very important step. This must be done immediately, and the slide warmer must be set to >45oC (but not >50oC), or the sections may fall off during the hybridization and wash steps. Use the dust cover.

8. Assess quality of slides and choose the worst slides to be used as tester slides for estimating exposure length. You may also want to use some of the intermediate quality slides for the sense RNA control probe.

9. After baking, slides can be stored in slide boxes at 4C.




Requires DEPC-treated water and RNAse-free TE, 3M NaOAc, Ethanol, DTT, and tRNA. Also in vitro transcription kit, phenol, chloroform, and radiolabelled UTP.


A. The probes:


Generally two probes are synthesized. The first probe hybridizes with the RNA of interest, and is generally called antisense, anti-mRNA, or (+) strand probes. You must also synthesize a control probe that will not hybridize with the RNA of interest. The most convenient thing to do is to synthesize an RNA probe in the opposite orientation as the anti-RNA probe. Probes of this type are called sense, mRNA, or (-) strand probes. A second control probe that can be used is [3H]-poly(U). This control is used to describe the distribution of total poly(A) RNA in the tissue and to assess whether the RNA within a cell is accessible to the in situ hybridization probes. We purchased our [3H]-poly(U) from Amersham, but they no longer sell it. It can probably be purchased from another supplier.


B. Template preparation:


1. Digest 5 ug or more with the appropriate restriction enzyme to linearize the template so that a "run-off" transcript can be generated. Avoid the use of enzymes that generate 3' overhangs--they can act as promoters for the polymerases.

2. Phenol/chloroform extract twice, and chloroform extract twice with an equal volume. Use RNAse free eppendorf tubes.

3. Precipitate the DNA with 1/10 volume RNAse free 3M NaOAc and 2.5 volumes EtOH. Wash with RNAse free 70% EtOH, and resuspend in RNAse-free TE at about 1 mg/ml. Assess digest extent and concentration of DNA on a gel.


C. Probe synthesis:



Transcription kit. We use the Promega Transcription kit (P1460, with SP6 and T7 RNA polymerase), but other kits work equally well.
RNAse-free tRNA (We use BRL # 5401 or Boeringher Manheim #?)
RNAse-free DNase (We use Promega RQ1 DNAse, # M6101 (also available in Promega Transcription Kit))
[35S]-UTP: We purchase our label from Amersham . Generally the 650 Ci/mmole specific activity (catalog number SJ 263) is sufficient.


1. Dry down [35S]-UTP (25 ul of 650 Ci/mmol 250 mmol for a single reaction, 50 ul for a double reaction). Resuspend in 20 ul RNase free H2O for a single reaction, and 40 ul for a double reaction. A half reaction is sufficient for many slides.

2. Set up transcription reaction:

Mix the following at room temperature for a single reaction:

H2O + [35S]-UTP 20 ul
5x Transcription Buffer 10 ul
100 mM DTT 5 ul
RNasin 2 ul
ATP, GTP, CTP mix (3 ul each, 2.5 mM each) 9 ul
linearized plasmid (1ug/ul) 1 ul
H2O 2 ul
T7 or SP6 polymerase 1 ul
50 ul

3. Incubate at 40°C for 1 hr. Take 1 ul sample for total counts.

4. Add 1 ul RNAse-free DNase per ug of template, and incubate at 37°C for 15 min.


From here there are two options:


-- The Beth/Steve approach: This method does not use a column for purification, but could result in higher background in some cases.

5B-S. Add up to 200 ul TE
2 ul 1M DTT
2ul 10mg/ml tRNA
phenol/chloroform extract
chloroform extract
1/10 vol 3M NaOAc, ph 5.2
2.5 vol Ethanol
15 min spin
70 % wash
resuspend in 55 ul DEPC H2O

6B-S. Count 1ul in glass scintillation vial with 5 ml scintillation fluid, Program User 4. Make sure you have enough probe for each slide at 2-4 million counts/slide.

7B-S. Take off a sample of 107 cpm to run on a gel (pre-hydrolysis sample). If less than 107 cpm/ 3 ul, just take 3 ul and add to 2 ul of Sequenase stop buffer.


-- The Hajime/Bobby approach:

5H-B. Phenol/chloroform extract twice, chloroform extract once. Run through G-50 columns (Boeringer Manheim) as instructed. Use one column for each single reaction.

6H-B. Count 1ul in glass scintillation vial with 5 ml scintillation fluid, Program User 4. Make sure you have enough probe for each slide at 2-4 million counts/slide.

7H-B. Take off a sample of 107 cpm to run on a gel (pre-hydrolysis sample). If less than 107 cpm/ 3 ul, just take 3 ul and add to 2 ul of Sequenase stop buffer.

8H-B. Precipitate with 1/10 volume 100 mM DTT, 2ul tRNA, 1/10 volume 3M NaOAC, and 2.5 volumes of Ethanol. Resuspend in 50 ul H20


D. Probe Hydrolysis:


We reduce the size of our probes to allow better penetration into the tissue. According to Moench et al. (1985), probes of mean length 70 nt give higher signals than larger probes (150 nt and larger). We chemically degrade our probes to a mean length of 75-100 nt. The RNA probes are chemically degraded by incubation in alkali at 60oC. The length of time for incubation is determined by the formula below:

t = (Lo - Lf) / (K)(Lo)(Lf)
Lo = starting length (kb), including all parts of the vector that are transcribed
Lf = final length (kb) = .075-.1 kb
K = .11
Calculated time will be in minutes.
Sample calculation, 1 kb transcript to 100 nt mean size:
(1-.1)/(.11)(1)(.1) = 82 minutes.


1. Hydrolysis buffers:

(a) 200 mM Na2CO3 (Sodium Carbonate) (0.21g in 10 ml)
(b) 200 mM NaHCO3 (Sodium Bicarbonate) (0.17g in 10 ml)
NOTE: Make buffers fresh each time and do not autoclave.

2. Hydrolysis reaction:

(a) Mix the following:

50 ul RNA
30 ul 200 mM Na2CO3
20 ul 200 mM NaHCO3

(b) Incubate at 60oC for the calculated time.

(c) Stop the reaction by putting on ice and adding the following:

3 ul 3 M Sodium Acetate (pH 6.0)
5 ul 10% Glacial Acetic Acid

3. Take off a sample of 107 cpm to run on a gel (post-hydrolysis sample).

4. Precipitate the probes:

Volume = 108 ul and already has 3 ul of 3 M Sodium Acetate.
Add: 1 ul 1M DTT
1 ul 10 mg/ml tRNA carrier
8 ul 3 M Sodium Acetate
250 ul Ethanol

4. Run pre- and post-hydrolysis samples on a denaturing sequencing gel to determine the size of the probe. Add 3ul to 2 ul of seq loading dye. Heat 75 2min, cool on ice, load 2.5 ul on gel. Just do short run. Second dye runs at about 75 nt.

5. Store the probe as an ethanol precipitate until needed. After spinning down the probe, resuspend it in 25-50 ul of TE + 10 mM DTT. Count 1 ul in Scintillation fluid.




Setting up RNAse-Free glassware

Two days before hybridization day, wash with soap and water a minimum of 28 staining dishes and lids (VWR 25445-009) for the first rack of 30 slides you plan to do, and a minimum of 6 dishes for each additional rack of 30 slides. (Try to do extra, since dishes tend to break in certain steps). Also wash a minimum of 2 2L glass cylinders and 2 500ml glass cylinders, slide racks, slide rack holders. and one 1L flask for each set of 30 slides. Let dry several hours to overnight. The day before the experiment, bake the glassware and racks in the Rothenberg oven for 5 hours. (The temperature is already set.) Turn the oven off and let the glassware cool with the oven door closed for at least 8 hours.


A. Solutions (all are DEPC-treated or made with DEPC-treated water):


1. Prepare about 10L of DEPC-water for the first rack of slides, then 3-4L for each additional rack.

2. Set up 10 staining dishes containing the following ethanol solutions: 100%, 100%, 95%, 85%, 70%, 50%, 30%, 15%, H2O, H2O.

3. 1 M Tris (7.5) RNAse-free 500 ml stock:

Use unopened bottle of tris. Add 60.55 g to about 400 ml DEPC-H2O. Add about 65 ml conc. HCl; confirm with pH paper.

4. Proteinase K Solution:

Need 500 ml for each rack of slides. Prewarm to 37°C.
100 mM Tris (7.5)-- from 10x stock above
50 mM EDTA-- from 10X DEPC-EDTA

5. 2X SSC--500 ml for each rack of slides.

6. 10X PBS-- about 500ml 10X needed for each rack of slides

NaCl 76g
Na2HPO4.2H2O 12.46g (18.76g of Na2HPO4.7H2O)
NaH2PO4.2H2O 4.48 (4.14g of anhydrous NaH2PO4)
H2O to 1 Liter

7. 10x Hybridization Salts:

3 M NaCl
100 mM Tris (7.5)
10 mM EDTA (8)

8. 50% Dextran Sulphate. We use Dextran Sulphate from Pharmacia (# 17-0340-01).

9. 1M Dithiotheitol (DTT)

10. 10 mg/ml RNAse-free yeast tRNA We use tRNA from BRL (# 5401)

11. 50X Denhardt's Solution. We filter it just before use.

12. Other Reagents:

(a) Distilled Formamide (We use BRL # 15515)
(b) Triethanolamine (We use Sigma # T1377)
(c) Acetic Anhydride (We use Mallinckrodt # 2420-200)


C. Remove paraplast from sections:


1. Put slides in a staining dish slide holder.

2. Fill a staining dish with xylenes, and place a small stir bar at the bottom. Place slides into this staining dish.

3. Stir for 10 minutes. Stir very slowly--just fast enough to mix the solution a bit.

4. Repeat with fresh xylenes.

5. If you are processing many racks of slides, you can use the same two xylenes solutions.


D. Remove xylene from slides:


1. Remove slides from xylenes and place into 100% ethanol (95% ethanol will not work). Dip up and down about 15 times or until the "streaks" go away.

2. Repeat with the second 100% ethanol.



E. Hydration:


1. Process the slides through the following ethanol solutions: 95%, 85%, 70%, 50%, 30%, 15%, H2O, H2O. Once again, dip up and down 15 times or until the "streaks" go away. Begin with the 95% ethanol and end with the H2O, so as to hydrate the tissue gradually.

2. Change the last water for each rack of slides.

3. Save the ethanol solutions for late steps.


F. Denaturation


1. Incubate slides in 0.2N HCl for 20 minutes at room temperature (8.6 ml conc. HCl in 500 ml H2O). This step denatures proteins and nicks DNA, but will also partially reverse the fixation step.

2. Incubate in 2X SSC, 70°C, for 15 minutes. NOTE: THIS IS A STEP WHERE DISHES ARE SUSCEPTIBLE TO BREAKING. To avoid breakage, place the dishes in a tupperware container partially filled with water, and place in the RT water bath, then turn on the heat to 70 C. This step denatures RNA and also removes some of the proteins to make the RNA more accessible to hybridization.

3. Wash slides in 1X PBS for 2 minutes at room temperature.


G. Proteinase K digestion:


Proteinase K is used to partially digest the tissue to allow better probe penetration. This step increases the signal. For Arabidopsis floral tissue, the best incubation time is 30-45 minutes. Less time gives a weaker signal, and greater time tends to destroy the morphology of the tissue. The timing was calibrated with a Proteinase K solution that was prepared just before the incubation.


1. Prewarm the Proteinase K Solution to 37o C in a staining dish, using the 37 C incubator.

2. Add Proteinase K to the prewarmed solution to a final concentration of 1 ug/ml. I make up a fresh 5 mg/ml solution and add 100 ul of this to each 500 ml of solution.

3. Put slides in solution and incubate for 30 minutes at 37oC. Agitate every 10 min.

4. After the incubation, remove from the Proteinase K solution and place into a staining dish filled with H2O. Dip up and down a few time to rinse off the Proteinase K.

5. Repeat the H2O wash.

6. Rinse in 1X PBS

7. Incubate in 2mg/ml glycine in 1X PBS for 2 minutes at room temperature (dissolve dry glycine in PBS). This step blocks the protease.

8. Rinse in 1X PBS, 2 times for 2 minutes.


H. Post-Fix


1. Make post-fix solution (4% formaldehyde in PBS), fresh 30 min before use. Dissolve 20g of paraformaldehyde in 450ml water (in a 1L flask which has been treated with chloroform or baked), bring pH up to 9-10 with 2ml of 2N NaOH (400 ul of 10N NaOH). Check with pH paper. Microwave to about 60°C until the paraformaldehyde goes into solution. Check temperature of formaldehyde solution with a thermometer that was dipped in chloroform.

2. Bring pH back down to pH 7.0 with 2ml of 2N HCl (345 ul Conc. HCl). Then add 50ml of 10X PBS and cool in ice H2O down to room temperature.

3. Post-fix slides for 20 minutes in staining dish.

4. Rinse in 1X PBS, 2 x 5 minutes at room temperature.


I. Acetylation Reaction:


This reaction acetylates any remaining positive charges in the tissue or on the slides, and is done to reduce background Hayashi et al. (1978). One important point, the half life of acetic anhydride in aqueous solution is less than 1 minute.


1. Make up 500 ml of 100mM Triethanolamine for each rack of slides (6.7 ml Triethanolamine per 500 ml H2O). Adjust pH to 8.0 with about 2 mls of concentrated HCl. Check with pH paper.

2. Set up a staining dish that has 500 ml of 100 mM Triethanolamine and a chloroform treated stir bar at the bottom.

3. Stir the bar very hard and add 1.25 ml of acetic anhydride. Wait about 5 seconds to mix, turn off stir bar, and add slides.

4. Incubate at room temperature for 5 minutes.

5. Rinse in 1X PBS, 2 x 5 minutes at room temperature.


J. Dehydration:


The purpose of this step is to completely dry the sections so that when the hybridization solution is applied, it is sucked into the tissue.


1. Progress through the following ethanol solutions so as to dehydrate the tissue gradually (except for the 100% and 95% ethanols, use the ethanol solutions from step C): 15%, 30%, 50%, 70%, 85%, 95%, 100%.

2. Dry under a vacuum (will take about 1 hour). Alternatively one can let sit overnight to dry: wrap in aluminum foil.


K. Pre-Hybridization:


1. Make several humidified boxes. We place several layers of paper towels that are soaked with 50% formamide/300 mM NaCl at the bottom of a tupperware box. Make sure to use a container with a flat bottom. It is a good idea to elevate the slides so that the probe does not get wicked away. We place the slides onto a pair of 10 ml plastic pipets.

2. Use 175 ul of pre-hyb solution per slide. Make a little extra in case you run out. Keep at RT.

50% Formamide
1X Hybridization salts
1X Denhardt's Solution
70 mM DTT
150 ug/ml tRNA

3. Apply Pre-hyb solution to one end of slide. Slowly place coverslip down on slide and make sure the pre-hyb is distributed over all the tissue. (Prehyb solution can be kept at room temperature.)

4. Place slides in 45° C incubator for 1-2 hours.


L. Hybridization:


1. Amount of hybridization solution to make:

Use 175 ul per slide for a surface area of 24 x 50 mm. Make 10-20% extra in case you run short.

2. Hybridization Solution for anti-mRNA and mRNA probes:

50% Formamide
1X Hybridization salts
1X Denhardt's Solution
10% Dextran Sulfate
70 mM DTT
150 ug/ml tRNA
500 ug/ml Poly A
2-4 x 10*6 cpm/slide (boil 3 min, quick cool in ice H2O, then 50C block)

3. Hybridization solution for [3H]-poly(U) probe:

300 mM NaCl
10 mM Tris (7.5)
1X Denhardt's Solution
150 ug/ml tRNA
25 units/ml RNasin
1 ng/ul [3H]-poly(U)

4. Remove prehybridization solution and slides by holding the slides vertically on a paper towel.

5. Apply hybridization solution to slides:

Use appropriate precautions for radioactive work.
(a) Pre-warm the hybridization solution to 50oC in a heat block. This makes it easier to spread.
(b) Apply hybridization solution in a pool to one side of the slide. Spread with a yellow tip. Make sure the tissue is wetted--sometimes the hybridization solution has a tendency to surround but not penetrate the tissue. This step is made easier by using a lot (175 ul) of hybridization solution.
(c) Cover slide with coverslip. When all of the tissue sections are wet, slant the slide briefly, so that a small pool of hybridization solution accumulates to one end. Hold the coverslip at one end with a jeweler's forceps and place the other end on the end of the slide that has the small pool of hybridization solution. Slowly lower the coverslip to completely cover the slide. Avoid bubbles--If you get a lot of bubbles, start over.

6. Place slides in the humidified box and incubate at 45oC (45-48 is fine) overnight.




These steps do not require RNAse free glassware or solutions.


A. Solutions:


1. RNase Buffer:

Need 500 ml for each set of 30 slides. Preheat to 37°C.
500 mM NaCl
10 mM Tris (7.5)

2. First Wash Buffer-- Need 1.5 L in 3 dishes

5 mM DTT 1.16 g

3. Low Stringency Wash Buffer-- Need at least 3 L, 3 dishes plus big container.

5 mM DTT 2.32 g

4. High Stringency Wash Buffer-- Need 4 L, preheat to 57C

5 mM DTT 1.54 g to each 2L, add just before adding slides.


B. Remove coverslips and hybridization solution:


1. Dip the slides individually in a staining dish filled with First Wash Buffer until the coverslip falls off. Dispose in radioactive liquid waste.

2. Place slides back into a rack and into a fresh solution of First Wash Buffer. Dip up and down a few times.

3. Repeat so that the slides have been rinsed 3 times with First Wash Buffer.


C. RNase Treatment:


This step can reduce background significantly (2-5 fold). In an experiment with the AGAMOUS probe, I tried to reduce background further by raising the RNase concentration to 100 ug/ml and 200 ug/ml. Both concentrations failed to reduce background over the reduction achieved with 25 ug/ml RNase. More significantly, the slides incubated with 200 ug/ml RNase had no signal.


1. RNase Precautions:

(a) Making up an RNase stock solution:
Never weigh out RNase!!! You do not need to make up RNase fresh each time. To avoid weighing out RNase, buy a bottle, and add RNase Buffer directly to the bottle to achieve a concentration of 25 mg/ml. Store frozen at -20oC. This is a 1000X stock. This RNase stock will be good for >1 year. We use Ribonuclease A from Sigma (catalog # R-5503). 100 mg size: add 4 ml of RNAse buffer.
(b) RNase glassware:
We have a special set of staining dishes that are used for the RNase incubation and the first 3 washes. Use a plastic 1 ml pipet to measure out the 500 ul of stock solution. Do not put the RNase glassware through the normal wash. Without splashing, rinse off the RNase glassware >10 times to dilute the RNase.

2. Prewarm RNase Buffer to 37oC. You will need 500 ml for each rack of 20 slides. The best way to prewarm is to fill the large staining dishes with RNase Buffer and place them into a 37oC water bath. This way, the glass gets warmed up too. Use the RNase staining dishes.

3. Thaw the 25 mg/ml RNase stock, and dilute to 25 ug/ml by adding 500 ul to each staining dish filled with 500 ml of RNase Buffer. Mix by stirring with a plastic pipet. RNase Precaution--Do not use a pipetman to measure out the RNase stock--use a plastic 1 ml pipet.

4. Place slides into the RNase solution, and incubate at 37oC for 30 minutes. Dip up and down every 10 minutes during the incubation to remove bubbles.

5. Fill 3 staining dishes with Low Stringency Wash Buffer. Use the RNase staining dishes.

6. After the 30 minute incubation, remove slides from the RNase solution and place into the first staining dish containing Low Stringency Wash Buffer. Dip up and down 15 times to rinse off the RNase.

7. Repeat with the second and third solutions of Low Stringency Wash Buffer. Total of 3 washes.


D. Low Stringency Wash:


1. Fill a large Tupperware container (4 L container) with Low Stringency Wash Buffer and place on top of a magnetic stirrer.

2. Place the slides (still in slide holder) in the buffer, and wash with stirring for 30 minutes.

3. At the end of the low stringency wash, remove the poly-U slides and place in a staining dish containing low stringency wash buffer until the other slides are processed. No high stringency wash for the poly-U slides!


E. High Stringency Wash:


1. Place a 4 L beaker containing 2 L of High Stringency Wash Buffer in a 57oC water bath to preheat. Measure the temperature of the buffer to make sure it is actually at 57oC--sometimes the temperature of the water bath must be set above 57oC to achieve this temperature in the buffer. Keep another 2L warm in the 60oC oven.

2. Place the slides (still in slide holder) in the 57oC High Stringency Wash Buffer, and incubate two times for 1 hour each time. Don't forget the DTT. If possible, agitate the buffer by stirring or shaking. Dip slides up and down occasionally--bubbles accumulate on the slides.


F. Dehydration:


1. Dehydrate the tissue gradually by processing through the following ethanol series: 15%, 30%, 50%, 70%, 85%, 95%, 100%.

2. Dry under a vacuum (requires about 1 hour).


G. Quick Autoradiography:


Optional. Expose slides overnight on Biomax MR X-Ray film to assess background. If there is signal in the larger structures, this will be visible.




NOTE: Steps A-C should be carried out in complete darkness. Turn off all light sources (i.e. computers) in the darkroom, and carefully cover the waterbath light and all sides of the waterbath. Minimize entering and leaving, and put a sign on the darkroom door indicating in situs are in progress.


A. Emulsion:


1. Purchase NTB-2 emulsion from IBI/Kodak (1-800-752-9128). This should be used fresh, or an older batch tested before use by developing a blank slide dipped in the emulsion.

2. Dilute the emulsion 1:1 with H2O.

(a) Prewarm the emulsion to 45°C in a water bath in a 500 ml flask. Place an equal volume of H2O in the flask and prewarm to 45°C.
(b) Pour the emulsion into the flask, swirl to mix, return to the 45°C water bath.

3. Aliquot into slide mailers (S/P Baxter # M6271). Most slide mailers can be filled by using a 25 ml plastic pipette, filled to maximum volume with a pink pipette dispenser.

4. Wrap in 2 layers of aluminum foil, place into a dark box, and store at 4°C.


B. Dipping Slides:


1. Melt emulsion in a 45°C water bath.

2. Dip a blank slide in the emulsion to check the emulsion level, and to insure that the emulsion is well mixed. Look at it outside of the darkroom.

3. Dip each slide into liquid emulsion. Dip slowly, and one time only to avoid streaks.

4. Dry slides completely--place in a rack, inside a black box with desiccant, and put in dark room cupboard. Will take 30-60 minutes.

5. Place slides into dark boxes (VWR # 4844-003) with desiccant for exposure. Wrap desiccant in kimwipe, and place in box supported by slides. Wrap with 2 layers of aluminum foil. Use separate boxes for slides to test for exposure time.

6. Expose at 4°C for the appropriate time. For unknown probes, develop testers after 2 and 6 weeks, and develop remaining slides once the signal becomes visible.


C. Develop slides:


Materials: Available from Lee-Mac or other photography supply stores

Kodak D-19 Developer--to preserve the developer, blow the O2 out of the bottle with N2 after each use. In an emergency, the developer will work when brown. Kodak DEKTOL developer will also work.
Kodak Fixer (Do not use Rapid Fixer).


1. Make fresh developer and fixer:

Developer - while stirring, slowly add contents of packet to 1 gallon of water (3.8 L).
Put in 65°C incubator for several hours, till chemicals are completely dissolved. Makes 2x solution.
Fixer - while stirring, slowly add contents of packet to 3L of water at room temperature.
When powder is dissolved, bring volume up to 1 gallon (3.8 L)

2. For each experiment use freshly diluted developer (1:1 in water).

3. Fill 3 staining dishes with developer, water, and fixer. Put dishes in ice until they cool down to 15oC.

4. Remove the appropriate slides from the darkboxes and place into a slide holder.

5. Place slides into developer, dip up and down 5 times, and let sit in developer for a total of 4 minutes (2 minutes if using DEKTOL developer). Longer developing times preferentially produce grains in the emulsion background.

(note old protocol was D19 full strength for 2.5 min. Hajime. does 1:1 for 2 min in Dektol, Bobby does 1:1 D19 4 min.)

6. Remove slides from the developer and drain briefly.

7. Place slides into the water and dip up and down continuously for 30 seconds.

8. Remove slides from the water and drain briefly.

9. Place slides into fixer, dip up and down 5 times, and let sit in fixer for 5.5 minutes

10. Remove slides from fixer and drain briefly.

11. Place into water until ready to stain. Rinse in running water for 30 minutes to remove fixer.


D. Stain sections:



Toluidine Blue O (Sigma # T-3260)
0.1% Toluidine Blue in 0.1% sodium borate (filtered)


1. Soak slides in 0.1% Toluidine Blue until tissue is deeply stained. The emulsion and tissue will be stained very deeply. With fresh Toluidine Blue, this takes 15-30 seconds. If using an old Toluidine Blue solution, stain for 1-2 minutes.

2. Rinse off in water to get rid of excess stain.

3. Soak slides in water to destain. The stain will come out of the emulsion more quickly than the tissue. Soak just until the emulsion is well destained, but the tissue is still darkly stained; generally the emulsion will be faintly stained. This step can take > 1 hour. This step is accelerated by using a stir bar and rotating the slides every 30 min.

4. Dip 5-10 times in each of the following ethanol solutions: 25%, 50%, 75%, 100%, 100%. Move quickly through the ethanols because the tissue becomes destained in these solutions. If the emulsion is still somewhat stained, move through the ethanols more slowly to control the degree of staining. Note: If the tissue is too deeply stained, the hybridization grains will be obscured.

5. Dip slides in xylenes 15 times or until the streaks go away. The tissue will not destain in the xylenes.

6. Place slides into fresh xylenes.

7. Place coverslips on slides:

(a) Remove slide from xylenes and place onto a paper towel.
(b) Quickly add 2 drops of permount to the surface of the slide.
(c) Quickly place coverslip on slide.
(d) Squeeze out the excess xylene/permount--wipe surface of coverslip while holding down. When dried (next day), wipe surface with xylenes to clean.

8. The coverslips can be removed at any time by soaking in xylenes for 1-5 days.




A. Preparing slides for viewing


1. Scrape off emulsion from back of slide with a razor blade.

2. Rub off excess permount with a cotton-tipped applicator dipped in Xylene.

3. Examine slide, and repeat above steps if they are not clean.


B. Setting up microscope for phase contrast


1. Add water to top of condenser lens (directly below the stage). Turn condenser focusing knob until the condenser is all the way up. Turn on the illuminator, place a slide on the stage, and find a good section. Switch to 16x PH (phase contrast) lens.

2. Change the setting on the condenser ring (the ring just below the stage) to 2. The number 2 should be exactly aligned with the short white line on the right of the condenser ring. Press in the yellow filter knob (the knob farthest from the front of the microscope). This makes the light bluish.

3. Close the condenser light aperture (on the bottom of the microscope). A ring of light appears when viewed through the lens. Align this ring with the cross hairs, using knobs on the condenser ring.

4. Open condenser aperture. Move phase ring (on upper part of microscope) to PH setting. The phase ring contains two rings; the top ring for focusing, and the bottom one for changing the setting . Two donuts should appear, one light and one dark. Align donuts on top of each other with 2 knobs on the condenser ring.

5. Focus on the rings using the phase contrast focus ring (the upper ring). Set bottom of phase ring to 1.25

6. Take care to avoid bumping any of the phase contrast adjustment knobs. Repeat this procedure if this happens.


C. Photographing slides


Generally 3 pictures are advised: One bright field only, and two with both bright field and dark field (double exposures), with varying exposure times for dark field.

1. Use Kodak Ektachrome Elite 200 speed slide film. 100 speed also works. Load film in camera. (The shutter does not advance). Pull out metal shield.

2. Focus on the tissue in bright field. Pull upper knob all the way out to direct light to the camera instead of the eyepieces.

3. Turn on exposure apparatus (directly to the left of the microscope). Set lower left knob for the speed of the film (in most cases 200). The upper left knob should be on 3.

4. Adjust lower right knob until needle indicator is in the center. Use the upper right knob for fine adjustment. Press yellow button to take the picture.

5. Advance the film by pressing button on camera and winding film. Then Take another picture with the same setting, but do not advance the camera at this time (to allow for double exposure).

6. Switch to dark field. Switch condenser ring from 2 to D, turn off bottom blue filter (yellow knob out). Place yellow or red plastic plate filter directly on top of the condenser aperture. Push knob in to send light to the eyepieces.

7. Focus on grains (they are on a slightly different plane as the tissue). Pull knob all the way out to send light to the camera. Adjust exposure time as before, getting needle to the middle. Once it is in the middle, click the lower right knob back one (-1 exposure).

8. Press yellow button to expose film. It will take significantly longer than the bright field exposure. Once exposed, advance film as before. Take another dark field picture at -2 exposure by clicking the lower right knob back one more. Do not advance film after this exposure.

9. Switch back to bright field: Condenser ring from D to 2, turn on blue filter (yellow knob), remove red or yellow plastic filter, push knob to send light to eyepieces, and focus on the tissue.

10. Adjust exposure, take another picture, and advance film. This gives 3 total pictures per section, including two double exposures.