Protease Digestion

Protease treatment is applied to sections to be used for in situ hybridization of DNA or RNA but not to those sections utilized for immunohistochemistry or enzyme histochemistry. An appropriate protease digestion of sections removes some of the proteins from the cellular network to make DNA or RNA more accessible for in situ hybridization. It subsequently enhances the hybridized signal of interest, reducing background as well.

1. Prepare a sufficient volume of protease solution as follows: Proteinase K solution: 1 mg/ml in 50 mM Tris-HCl, pH 7.5.

Pepsin solution: 2 mg/ml in dd.H2O, pH ~2.0 (40 mg pepsin in 19 ml dd.H2O and 1 ml 2 N HCl). Trypsin solution: 2 mg/ml in dd.H2O (40 mg pepsin in 19 ml dd.H2O and 1 ml 2 N HCl).

Notes: (1) Protease solutions can be stored at -20°C for up to 2 weeks. (2) Although each of the three protease solutions works equally well in our laboratories, we recommend using pepsin or trypsin solution. Proteinase K digestion is not readily controlled compared with pepsin or trypsin. As a result, overdigestion of tissue usually occurs. (3) Proteinase K is relatively difficult to inactivate or wash away from the tissue. In contrast, it is easy to inactivate and wash pepsin or trypsin by simple adjustment of the pH from 2.0 to 7.2 to 7.5, which is the standard pH of PBS.

2. Carefully overlay the tissue sections with an appropriate volume of the protease solution.

Add protease solution to the tissue sections instead of immersing the slide rack into a staining dish containing a large volume of protease solution. Otherwise, protease covers the entire surface of the slides and it is not readily washed away after treatment.

3. Carry out digestion of tissues at 37°C for 10 to 30 min.

Notes: Protease digestion is a tricky step for the success of in situ hybridization. Insufficient protease digestion may result in a diminished or completely absent hybridization signal. If overdigestion occurs, the tissue morphology may be poor or even completely destroyed. In order to obtain a good signal-to-noise ratio and a good preservation of tissue morphology, it is recommended to determine the optimal digestion time for each tissue sample. Different tissues may have different optimal digestion times. We usually set up a series of protease treatment times (5, 10, 15 and 30 min) for tissue sections fixed for 0.5 to 2, 3 to 5, 7 to 10 and 12 to 18 h, respectively. Each test should consist of two or three replicates. It is hoped that these trial tests will provide investigators with a useful guide to determine the optimal digestion time for specific in situ hybridizations.

4. After protease digestion is complete, stop proteolysis by rinsing the slides in a staining dish containing PBS (pH 7.4) with 2 mg/ml glycogen. Perform three changes of fresh PBS, 1 min each. Alternatively, wash the slides for 1 min in a solution containing 100 mM Tris-HCl and 100 mM NaCl.

5. Post-fix the specimens in freshly prepared 10% buffered-formalin solution or 4% PFA fixative for 6 min at room temperature.

6. Rinse the slides with PBS three times, 1 min for each rinse.

7. Rinse the slides in 100% ethanol for 1 min and air-dry. At this stage, the sections will be subjected to different treatments, depending on in situ hybridization with cellular DNA or with RNA.


This treatment serves to remove cellular DNA from tissue. We have found that genomic DNA may compete with the RNA species of interest in hybridizing with a specific probe. Subsequently, DNase digestion can reduce nonspecific background for in situ hybridization of RNA.

1. Overlay the tissue sections with an appropriate volume of commercially available RNase-free DNase solution (e.g., Boehrinnger Mannheim Corporation).

2. Allow digestion to proceed for 30 min at 37°C.

3. After digestion is complete, inactivate DNase by rinsing the slides in a staining dish containing a solution of 10 mM Tris-HCl (pH 8.0), 150 mM NaCl and 20 mM EDTA for 5 min.

4. Wash the slides in TE buffer (pH 8.0) for 3 x 2 min and 1 min in 100% ethanol and air-dry. At this stage, the slides are ready for hybridization or can be stored in a slide box with desiccant at -80°C until use.

Note: In order to prevent RNase contamination, all the tools used for in situ hybridization of cellular RNA should be completely cleaned and rinsed with DEPC-treated dd.H2O. All the solutions should be treated with 0.1% (v/v) DEPC or prepared using DEPC-treated water. RNase contamination may completely ruin the experiment.


Note: This treatment functions to degradate cellular RNA in the tissue inasmuch as RNA can compete with DNA species of interest in hybridizing with a specific probe. As a result, RNase digestion can enhance the hybridized signal by reducing nonspecific background for in situ hybridization of DNA.

1. Overlay the tissue sections with an appropriate volume of RNase solution (DNase free).

2. Allow digestion to proceed for 30 min at 37°C.

3. After digestion is complete, inactivate RNase by rinsing the slides in a staining dish containing a solution of 10 mM Tris-HCl (pH 8.0), 150 mM NaCl and 20 mM EDTA for 5 min.

4. Wash the slides in TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) for 3 x 2 min and 1 min in 100% ethanol and then air-dry. At this stage, the slides are ready for hybridization or can be stored in a slide box with desiccant at -80°C if the time schedule is not convenient.


Synthesis of Probes for DNA Hybridization Using Random Primer Labeling of dsDNA

Random primer labeling kits are available from commercial companies such as Pharmacia, Promega Corporation, and Boehringer Mannhein Corporation. A kit usually includes a population of random hexanucleotides synthesized by an automated DNA synthesizer. Such a mixture of random primers contains all four bases in each position. The primers can be utilized to primer DNA synthesis in vitro from any denatured, closed circular or linear dsDNA as a template using the Klenow fragment of E. coli DNA polymerase I. Inasmuch as this enzyme lacks 5' to 3' exonuclease activity, the DNA product is produced exclusively by primer extension instead of by nick translation. During the synthesis, one of the four dNTPs is radioactively labeled and incorporated into the new DNA strand. By primer extension, it is relatively easy to produce a probe with extremely high specific activity (108 cpm/mg) because more than 70% of the labeled dNTP can be incorporated into the new DNA strand. In a typical reaction, the amount of DNA template can be as low as 25 ng.

1. Place 1 ml of DNA template (0.5 to 1.0 mg/ml in dd.H2O or TE buffer) in a sterile microcentrifuge tube. Add an appropriate volume of dd.H20 or TE buffer (19 to 38 ml) to the tube to dilute the DNA template to 25 to 50 ng/ml.

2. Cap the tube and denature the dsDNA into ssDNA by boiling for 6 min, and quickly chilling the tube on ice H2O for 2 to 4 min. Briefly spin down.

3. Set up a labeling reaction by adding the following components, in the order listed:

Denatured DNA template (25 to 50 ng), 1 ml

5X Labeling buffer, 10 ml

Acetylated BSA (10 mg/ml) (optional), 2 ml

Three unlabeled dNTP mixture (500 mM each), 2 ml

[35S]dCTP or [35S]dATP(1000 to 1500 Ci/mmol), 2.5 to 4 ml

Klenow enzyme, 5 units

Add dd.H20 to final volume of 50 ml.

Note: The standard amount of DNA template for one reaction should be 25 to 50 ng. Because the specific activity of a probe depends on the amount of the template, the lower the amount, the higher the specific activity of the probe. [35S]dCTP or [35S]dATP (1000 to 1500 Ci/mmol) should not be >5 ml used in the reaction to prevent potential high background.

Caution: [35S]dCTP and [35S]dATP are dangerous and should be handled carefully using an appropriate Plexiglas protector shield. Any gloves and waste buffer/solutions containing radioactive materials should be placed in special containers for isotopic disposal.

4. Mix well and incubate the reaction at room temperature for 60 min.

5. Stop the reaction by adding 5 ml of 200 mM EDTA and place on ice for 2 min.

6. Determine the percentage of [35S]dCTP incorporated into the DNA and purify the probe from unincorporated nucleotides using the protocols described next.

Protocol 1. DE-81 Filter-Binding Assay

1. Dilute 1 ml of the labeled mixture in 99 ml (1:100) of 200 mM EDTA solution (pH 8.0). Spot 3 ml of the diluted sample onto two pieces of Whatman DE-81 filter papers (2.3 cm diameter). Air-dry the filters.

2. Rinse one filter with 50 ml of 0.5 M sodium phosphate buffer (pH 6.8) for 5 min to remove unincorporated cpm. Repeat washing twice. The other filter will be utilized directly to obtain the total cpm of the sample.

3. Add an appropriate volume of scintillation fluid (~10 ml) to each vial containing one of the filters. Count the cpm in a scintillation counter according to the manufacturer's instructions.

Protocol 2. TCA Precipitation

1. Dilute 1 ml of the labeled reaction mixture in 99 ml (1:100) of 200 mM EDTA solution and spot 3 ml on a glass fiber filter or on a nitrocellulose filter for determination of the total cpm in the sample. Allow the filter to air-dry.

2. Add 3 ml of the same diluted sample into a tube containing 100 ml of 100 mg/ml carrier DNA or acetylated BSA and 20 mM EDTA. Mix well.

3. Add 1.3 ml of ice-cold 10% trichloroacetic acid (TCA) and 1% (w/v) sodium pyrophosphate to the mixture. Incubate the tube on ice for 20 to 25 min to precipitate the DNA.

4. Filter the precipitated DNA on a glass fiber filter or on a nitrocellulose filter using vacuum. Rinse the filter with 5 ml of ice-cold 10% TCA four times under vacuum. Wash the filter with 5 ml of acetone (for glass fiber filters only) or 5 ml of 95% ethanol. Air-dry the filter.

5. Transfer the filter to two scintillation counter vials and add 10 to 15 ml of scintillation fluid to each vial. Count the total cpm and incorporated cpm in a scintillation counter according to the manufacturer's instructions.

Protocol 3. Calculation of Specific Activity of the Probe

1. Calculate the theoretical yield:

mCi dNTP added X 4 X 330 ng / nmol ng theoretical yield =

specific activity of the labeled dNTP (mCi / nmol)

2. Calculate the percentage of incorporation:

cpm incorporated

% incorporation =-x 100

total cpm

3. Calculate the amount of DNA produced:

ng DNA synthesized = % incorporation x 0.01 x theoretical yield

4. Calculate the specific activity of the prepared probe:

total cpm incorporated (cpm incorporated x 33.3 x 50)

The factor 33.3 is derived from the use of 3 ml of 1:100 dilution for the filter-binding or TCA precipitation assay. The factor 50 is derived from the use of 1 ml of the total 50 ml reaction mixture for 1:100 dilution. For example, given that 25 ng DNA is to be labeled and that 50 mCi[35S]dCTP (1000 Ci/mmol) is utilized in 50 ml of a standard reaction, and assuming that 4.92 x 104 cpm is precipitated by TCA and that 5.28 x 104 cpm is the total cpm in the sample, the calculations are as follows:

Theoretical yield =-= 66 ng

1000 mCi/nmol

Specific activity-= 9.5 X10 cpm / mg DNA

Protocol 4. Purification of a Radioactive Probe

It is recommended to carry out separation of a labeled probe from unincorporated isotopic nucleotide so as to avoid a high background (e.g., unexpected black spots on the hybridization filter). Use of chromatography on Sephadex G-50 spin columns or Bio-Gel P-60 spin columns is a very effective way for separation of a labeled DNA from unincorporated radioactive precursors such as [35S]dCTP or [35S]dATP. The oligomers will be retained in the column. This is particularly useful when an optimal signal-to-noise ratio with 150 to 1500 bases in length probe is generated.

1. Suspend 2 to 4 g Sephadex G-50 or Bio-Gel P-60 in 50 to 100 ml of TEN buffer and allow equilibration to occur for at least 1 h. Store at 4°C until use. Alternatively, such columns can be purchased from commercial companies.

2. Insert a small amount of sterile glass wool into the bottom of a 1-ml disposable syringe using the barrel of the syringe to tamp the glass wool in place.

3. Fill the syringe with the Sephadex G-50 or Bio-Gel P-60 suspension up to the top.

4. Place the syringe containing the suspension in a 15-ml disposable plastic tube and place the tube in a swinging-bucket rotor in a bench-top centrifuge. Carry out centrifugation at 1600 x g for 4 min at room temperature.

5. Repeat addition of the suspended resin to the syringe and continue cen-trifugation at 1600 x g for 4 min until the packaged volume reaches 0.9 ml in the syringe and remains unchanged after centrifugation.

6. Add 0.l ml of 1X TEN buffer to the top of the column and recentrifuge as described previously. Repeat this step three times.

7. Transfer the spin column to a fresh 15-ml disposable tube and add the labeled DNA mixture to the top of the resin dropwise using a Pasteur pipette.

8. Centrifuge at 1600 x g for 4 min at room temperature. Discard the column containing unincorporated radioactive nucleotides into a radioactive waste container. Carefully transfer the effluent (about 0.1 ml) from the bottom of the decapped microcentrifuge tube or the 15-ml tube to a fresh microcentrifuge tube. The purified probe can be stored at -20°C until it is used for hybridization.

Preparation of [35S]UTP Riboprobe for RNA Hybridization by Transcription in Vitro Labeling

Recently, a number of plasmid vectors have been developed for subcloning the cDNA of interest. These vectors contain polycloning sites downstream from powerful bacteriophage promoters T7, SP6, or T3 in the vector. The cDNA of interest can be cloned at the polycloning site between promoters SP6 and T7 or T3, generating a recombinant plasmid, or transcribed in vitro into single-strand sense RNA or antisense RNA from a linear plasmid DNA with promoters SP6, T7 or T3. During the process of in vitro transcription, one of the rNTPs is radioactively labeled and can be incorporated into the RNA strand, which is the labeled riboprobe. The probe has a high specific activity and is much "hotter" than a ssDNA probe. As compared with DNA labeling, a higher yield of RNA probe can be obtained because the template can be repeatedly transcribed. RNA probes can be easily purified from DNA template merely with treatment of DNase I (RNase free). The greatest merit of an RNA probe over a DNA probe is that it can produce much stronger hybridization signals.

1. Prepare linear DNA template for transcription in vitro. The plasmid can be linearized by an appropriate restriction enzyme for producing run-off transcripts. The recombinant plasmid should be digested with an appropriate restriction enzyme that cuts at one site very close to one end of cDNA insert.

a. Set up a plasmid linearization reaction mixture on ice. Recombinant plasmid DNA (mg/ml), 5 mg Appropriate restriction enzyme 10 X buffer, 5 ml Appropriate restriction enzyme (10 to 20 u/ml), 20 units Add dd.H2O to a final volume of 50 ml.

b. Allow digestion to proceed for 2 h at the appropriate temperature, depending on the particular enzyme.

c. Carry out extraction by adding one volume of TE-saturated phenol/chloroform and mix well by vortexing. Centrifuge at 12,000 x g for 5 min at room temperature.

d. Transfer the top, aqueous phase to a fresh tube and add one volume of chloroform:isoamyl alcohol (24:1) to the supernatant. Mix well and centrifuge as described in the preceding step.

e. Carefully transfer the top, aqueous phase to a fresh tube. Add 0.1 volume of 2 M NaCl solution or 0.5 volume of 7.5 M ammonium acetate (optional), and 2.5 volumes of chilled 100% ethanol to the supernatant. Precipitate the linearized DNA at -80°C for 30 min or at -20°C for 2 h.

f. Centrifuge at 12,000 x g for 5 min, decant the supernatant and briefly rinse the DNA pellet with 1 ml of 70% ethanol. Dry the pellet for 10 min under vacuum and dissolve the DNA in 15 ml dd.H2O.

g. Use 2 ml of the sample to measure the concentration of the DNA by UV absorption spectroscopy at A260 and A280 nm. Store the DNA at -20°C until use.

2. Blunt the 3' overhang ends by the 3' to 5' exonuclease activity of Klenow DNA polymerase. Even though this is optional, we recommend that the 3' protruding end be converted into a blunt end because some of the RNA sequence is complementary to that of the vector DNA.

Note: Enzymes such as Kpn I, Sac I, Pst I, Bgl I, Sac II, Pvu I, Sfi and Sph I should not be utilized to linearize plasmid DNA for transcription in vitro. a. Set up an in vitro transcription reaction mixture on ice as described below:

5X Transcription buffer, 8 ml 0.1 M DTT, 4 ml rRNasin ribonuclease inhibitor, 40 units Linearized template DNA (0.2 to 1.0 g/ml), 2 ml Add dd.H2O to a final volume of 15.2 ml.

b. Add Klenow DNA polymerase (5 u/mg DNA) to the preceding mixture and incubate at 22°C for 15 min.

c. Add the following components to the reaction and mix well: Mixture of ATP, GTP, UTP (2.5 mM each), 8 ml

SP6 or T7 or T3 RNA polymerase (15 to 20 u/ml), 2 ml d. Incubate the reaction at 37 to 40°C for 60 min.

3. Remove the DNA template with DNase I.

a. Add DNase I (RNase free) to a concentration of 1 u/mg DNA template.

4. Purify the RNA probe as described for DNA labeling or by phenol/chloroform extraction followed by ethanol precipitation.

5. Determine the percentage of incorporation and the specific activity of the RNA probe, which is similar to the method in the preceding DNA labeling.

Note: The specific activity of the probe can be calculated as follows. For example, if 1 ml of a 1:10 dilution is utilized for TCA precipitation, 10 x cpm precipitated = cpm/ml incorporated. In 40 ml of a reaction mixture, 40 x cpm/ml is the total cpm incorporated. If 50 mCi of labeled [35S]UTP at 1000 mCi/nmol is used, then 50/1000 = 0.05 nmol of UTP added to the reaction. Assuming that there is 100% incorporation and that UTP represents 25% of the nucleotides in the RNA probe, 4 x 0.05 = 0.2 nmol of nucleotides incorporated, and 0.2 x 330 ng/nmol = 66 ng of RNA synthesized. Then, the total ng RNA probe = % incorporation x 66 ng. For instance, if a 1:10 dilution of the labeled RNA sample has 2.2 x 105 cpm, the total cpm incorporated is 10 x 2.2 x 105 cpm x 40 ml (total reaction) = 88 x 106 cpm; hence, % incorporation = 88 x 106 cpm/110 x 106 cpm (50 mCi) = 80%. Therefore, total RNA synthesized = 66 ng x 0.80 = 52.8 ng RNA. The specific activity of the probe = 88 x 106 cpm/0.0528 mg = 1.6 x 109 cpm/mg RNA.


After treatment of specimens with DNase (see the section on DNase treatment for in situ hybridization of RNA) or RNase (see the section concerning RNase treatment for in situ hybridization of DNA) and preparation of a specific probe (see the preceding section) are complete, one can proceed to in situ hybridization. In general, the procedure for DNA hybridization is similar to that for RNA hybridization except for the differences indicated at specific steps.

1. Carry out blocking of nonspecific sulfur-binding sites on tissue specimens prior to hybridization.

a. Remove the slides from the freezer and place them in a slide rack. Allow the slides to warm up to room temperature for a few minutes.

b. Immerse the slide rack in PBS buffer containing 10 mM DTT in a staining dish, and allow the specimens to equilibrate for 12 min at 45°C in a water bath.

c. Transfer the slide rack to a fresh staining dish containing a sufficient volume of freshly prepared blocking solution. Wrap the entire staining dish with aluminum foil and allow blocking of the specimens to proceed for 45 min at 45°C.

Note: Blocking nonspecific sulfur-binding sites on specimens can reduce the background a great deal when using [35S]-labeled probes. However, it is not necessary for [32P]-labeled probes. Caution: The blocking solution contains toxic iodoacetamide and N-ethylmaleimide, which should be handled carefully. Waste solutions should be placed in special containers for toxic waste disposal.

d. Wash the slides 3 x 3 min in PBS buffer and 3 min in TEA buffer at room temperature.

e. Block polar and charged groups on tissue specimens by transferring the slide rack to fresh TEA buffer containing 0.5% acetic anhydride.

f. Incubate the specimens in 2X SSC for 5 min at room temperature.

g. Dehydrate the tissue specimens through a series of ethanol (50%, 75%, 95%, 100%, 100%, and 100%), 3 min each at room temperature.

h. Completely dry the slides by air-drying or in a desiccate and proceed to prehybridization or store at -80°C if the schedule is not convenient.

2. Perform prehybridization as follows:

a. Carefully overlay tissue specimens with an appropriate volume of pre-hybridization solution (15 to 20 ml/10 mm2) using a pipette tip.

b. Place the slides in a high-humidity chamber and allow prehybridization to proceed for 1 h at 50°C.

c. During prehybridization, prepare an appropriate volume of hybridization cocktail (see Reagents Needed section).

d. After prehybridization is complete, carefully remove the prehybridiza-tion solution from the slides using a 3MM Whatman paper strip or a pipette tip.

e. Immediately overlay each specimen with freshly prepared hybridization cocktail containing the specific probe (15 to 20 ml/10 mm2). Cover the cocktail with a piece of plastic coverslip slightly larger than the tissue section.

f. Simultaneously denature the probe and the target DNA or RNA by placing the slides on a hot plate for 3 to 4 min at 95 to 100°C for DNA hybridization or at 65 to 70°C for RNA hybridization.

Note: DNA or RNA in the specimens can be denatured prior to prehybridization. Labeled probes can be denatured immediately before preparation of the hybridization cocktail.

g. Quickly place the slides in a moisture chamber and carry out hybridization at 55 to 65°C for 2 to 4 h.

Notes: (1) Any bubbles underneath the plastic coverslips should be removed by gently pressing the coverslip or with a toothpick. (2) In order to prevent uneven hybridization, slides should be kept level in the chamber. (3) The humidity chamber must be tightly covered or sealed prior to being placed in a hybridization oven or an equivalent. If the specimens are dried during hybridization, a high background will occur.

h. Following hybridization, quickly place slides in a slide rack and insert the rack into a staining dish filled with washing solution containing 2X SSC and 2 mM b-mercaptoethanol. Wash the slides with three changes of fresh washing solution, each time for 15 min at 55°C with gentle shaking.

Caution: Any used washing solution containing isotope should be collected in an isotope waste container for disposal. Any areas that touch isotopic solution or slides should be surveyed afterward for potential contamination by following local guidelines.

i. Continue to wash the slides twice, each session for 5 min, in a fresh washing solution containing 0.2X SSC and 2 mM b-mercaptoethanol at room temperature with gentle shaking.

At this stage, unhybridized RNA in the tissue specimens can be removed by using an appropriate amount of RNase (DNase free, Boehringer Mannheim Corporation) or a mixture containing 45 mg/ml RNase A, 2 mg/ml RNase T1 (Sigma Chemicals), 5 mM EDTA and 250 mM NaCl in TE buffer, pH 7.5. RNase treatment should enhance the signal-to-noise ratio, thus reducing background. j. Dehydrate the slides through a series of ethanol (50%, 75%, 95%,

100%, 100%, and 100%), 3 min each. k. Air-dry the slides. Proceed to emulsion autoradiography.

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