By Susan L. Naylor, John R. McGill, and Bernhard U. Zabel
In situ hybridization is the direct hybridization of a nucleic acid probe to metaphase chromosomes on a slide. Figure 1 depicts the overall scheme for in situ hybridization. Basically, metaphase chromosomes are immobilized on a slide and denatured. A tritiated labeled probe is also denatured and hybridized to the chromosomal DNA. After washing, slides are coated with autoradiographic emulsion. The sites of hybridization are revealed as silver grains produced as a result of the tritium acting on the autoradiographic emulsion. After developing the emulsion-coated slide, the chromosomes are banded, stained, and analyzed. Because of the structure of a metaphase chromosome, accessibility of chromosomal sites complementary to probe can be limiting, that is, not every complementary site will hybridize. Consequently, distribution of the grains over the metaphase chromosomes is analyzed statistically to distinguish specific from nonspecific background hybridization.
Copyright © 1987 by Academic Press, Inc. METHODS IN ENZYMOLOGY, VOL. 151 All rights of reproduction in any form reserved.
Carnoy's fixative (methanol: glacial acetic acid, 3:1) Prepared slides (cleaned with 7X detergent and alcohol)
3H-labeled nucleoside triphosphates (New England Nuclear) dCTP (specific activity 40-60 Ci/mmol) dTTP (specific activity 90-110 Ci/mmol) dATP (specific activity 40-60 Ci/mmol) dGTP (unlabeled) (PL Biochemicals) DNase I (1 mg/ml; Worthington DPFF) DNA polymerase I Nick translation buffer (1 OX) Tris-HCl, pH 7.8, 0.5 M MgCl2, 75 mM
BSA, 150 /ig/ml (nuclease free) 2-Mercaptoethanol, 0.1 M 0.5 MEDTA
10% Trichloroacetic acid (TCA)
Sephadex G-50 medium hydrated in column buffer (10 mM Tris, pH
7.6, 1 mM EDTA, and 0.1% SDS) Oligo labeling buffer LS: 1 M HEPES, pH 6.6/DTM/OL in ratio 25/25/7, store at -20° DTM: 100 nMdGTP, 250 mM Tris-HCl, pH 8.0, 25 mM MgCl2,
50 mM 2-mercaptoethanol OL: 1 mM Tris-HCl, pH 7.5, 1 mM EDTA, 90 OD units/ml oligo primers (hexamers from PL Biochemicals)
RNase I (2 mg/ml; Sigma) place stock in boiling water bath for 10 min 20X SSC (standard saline citrate) 3 M NaCl
0.3 M sodium citrate pH to 7.0 with HC1 and autoclave Dextran sulfate (50% in water, autoclaved)
Salmon sperm DNA (10 mg/ml in water, sheared through an 18-gauge needle, autoclaved) formamide (BDH or AnalR) (should be ~pH 7.0, after opening bottle, store remainder —20° in 50 ml conical tubes) 1 MNaOH
1 M sodium phosphate buffer pH 6.0 (made from 1 M stocks of monobasic and dibasic phosphate buffers in the ratio 2:1)
Kodak NTB-2 nuclear track emulsion Kodak D-19 developer, diluted 1:1 with water Safelight, dark red filter (Kodak No. 2, Cat. 152 1723)
Wright stain stock (EM Sciences: 0.25% in methanol, acetone free) Hoeschst H33258 (1 ¿¿g/ml in 2X SSC; Sigma) Giemsa stain (7% in 0.2 M phosphate buffer, pH 6.8)
Two alternative methods for chromosome preparation and banding are presented. One method produces elongated prometaphase chromosomes (approximately 1000 bands) for precise gene localization. The other method yields metaphase chromosomes of 400 bands, and are easier to recognize for the inexperienced eye. It is of the utmost importance to use the best chromosome spreads possible for in situ hybridization.
Chromosome Preparation (Method 1, Elongated Chromosomes; Adapted
Five milliliters of blood is drawn into a syringe containing 500 units heparin. The blood should be stored at room temperature and used while fresh. Twelve drops of heparinized blood are added to 4 ml chromosome medium 1A with PHA in 12.5 ml Falcon culture tubes and cultured for 72 hr. BrdUrd is then added to 200 /ig/ml. After 16-17 hr, the cells are washed twice with medium and then incubated in new culture tubes with 4 ml chromosome medium 1A with PHA and 10~5 M thymidine. After 7 hr cells are harvested by centrifugation (5 min at 1000 rpm) and resuspended in 6 ml warm (37°) 0.075 MKC1. The cells are incubated in the hypotonic solution for 8 min at 37°. The cells are pelleted by centrifugation (10 min, 1000 rpm) and fixed by resuspending them in Carnoy's fixative for 20 min. After three more washes with Carnoy's fixative, the cell suspension is
1 B. Dutrillaux and E. Viegas-Pequignot, Hum. Genet. 57, 93 (1981).
dropped onto cold slides (0°). The slides are air dried and stored at room
Chromosome Preparation (Method 2; Modified from Yunis and Chandler2)
Three milliliters of peripheral blood is drawn into a heparinized (0.1 ml sodium heparin, 1000 U/ml). The blood is expelled into 42.4 ml RPMI 1640 containing fetal bovine serum, antibiotics, and PHA (1.6 ml of reconstituted PHA from GIBCO). The mixture is divided into eight T-25 flasks which are incubated at 37°. Seventy-two hours later sterile methotrexate (60 ¡A of 10-5 M stock) is added to each flask and incubated at 37° for 17 hr. The contents of 2 flasks are combined into each of four 15 ml conical centrifuge tubes. The tubes are spun for 8 min at 400 g. Aspirate supernatant to just above pellet (note: lymphocyte layer is on top of pellet). Ten milliliters of warm (37°) RPMI 1640 without supplements is added to the cells and the cells are resuspended. The cells are pelleted and the wash repeated. Add 10 ml of RPMI 1640 with fetal bovine serum and 10~5 M thymidine. Place tubes into a 370 water bath for 5 hr (2 tubes) and 5 hr and 10 min (2 tubes). After incubation times are complete add 60 ¡A of colce-mid (10 /ig/ml) to tubes. Place tubes back into 37°waterbath for exactly 10 min. Centrifuge 8 min at 400 g. Aspirate supernatant; carefully add 10 ml warm (37°) hypotonic (0.075 M KC1) solution to tube and gently but thoroughly resuspend pellet. Incubate 10-20 min at 37° (time varies depending on blood sample). Centrifuge tubes 5 min at 400 g. Aspirate supernatant except for 0.5 ml above pellet. Gently resuspend pellet. Add freshly prepared Carnoy's fixative dropwise to 6-8 ml. Let tubes sit 1 hr. Wash cell pellet 4-5 times in fresh fixative; prepare slides.
DNA can be labeled by the same methods used in Southern filter hybridization. Both nick translation and random priming with oligonucleotides using tritiated nucleotides have been successful in our laboratory. The nick translation protocol is essentially that of Rigby et al? and the random priming method is that of Feinberg and Vogelstein.4
Nick Translation. To obtain the best hybridization, the nick translated probe should average 400 bases and have a specific activity of 107 cpm//ig
2 J. J. Yunis and M. E. Chandler, Prog. Clin. Pathol. 7, 267 (1977).
3 W. J. Rigby, M. Dieckmann, C. Rhodes, and P. Berg, J. Mol. Biol. 113, 237 (1977).
4 A. Feinberg and B. Vogelstein, Anal. Biochem. 132, 6 (1983).
1. Whole plasmid (0.25 to 1 ßg) is added on ice to a mixture of 6 ßM labeled nucleotides ([3H]dATP, [3H]dTTP, and [3H]dCTP), 60 ßM cold dGTP, IX nick translation buffer, and sterile water. The total volume of the reaction after addition of DNase I and DNA polymerase I should be between 20 and 30 ßl. If the nucleotides are supplied in ethanol, first
2. Dilute the stock of DNase I to 1 X 10-5 mg/ml with sterile water. Add DNase I at a final concentration of 10"~7 mg/ml in the reaction and mix. Let sit on ice for 30 min. Then heat for 10 min at 65°.
3. Add 5 units Escherichia coli polymerase I and incubate at 15° for 1 - 5 hr until at least 107 cpm//ig DNA is obtained. The reaction is monitored by TCA precipitation of the DNA onto glass fiber filters and counting in a scintillation counter. Stop the reaction by adding 5 ßl of 0.5 M
4. Remove the unincorporated label by column chromatography with Sephadex G-50 hydrated in column buffer. Apply the contents of the reaction tube to a column made in a siliconized Pasteur pipet. Elute with column buffer. Collect 100 ßl fractions into each of 15 tubes. Count a 2-jul aliquot of each fraction in aqueous scintillation fluid. Combine the tubes from the first peak and freeze at —20° until needed. The tritium-labeled plasmid is stable and can be used for up to 1 year.
Random Priming with Oligonucleotides; Adapted from Lin et al.5
1. Add the following reagents in order into a microcentrifuge tube: 0.75 nmol each of [3H]dATP, [3H]dCTP, and [3H]dTTP, water to a volume of 36.5 ßl, 10 //I of oligo-labeling buffer containing 96.8 ßM dGTP, 0.5 ßl bovine serum albumin (50 mg/ml), and 2 ß\ DNA (100 ngJß\). Denature in a boiling waterbath for 5 min and immediately quench in an ethanol-dry ice bath. Add 1 ßl (4 units) Klenow fragment of DNA polymerase I and incubate at room temperture for 1.5 to 4 hr.
2. The reaction is stopped by adding 5 ßl of 0.5 M EDTA, pH 8.0.
3. The unincorporated label is removed by column chromatography through Sephadex G-50 as described above.
The in situ hybridization procedure is that of Harper and Saunders6
1. Incubate slides with RNase A (100 ßg/ml in 2X SSC) for 1 hr at 37°. Rinse slides 4 times in 2X SSC (2 min per rinse) and then dehydrate with an ethanol series (2 min each in 70, 80, and 95% ethanol).
5 C. C. Lin, P. N. Draper, and M. DeBraekeleer, Cytogenet. Cell Genet. 39, 269 (1985).
6 M. E. Harper and G. F. Saunders, Chromosoma 83, 431 (1981).
2. Two methods are described for denaturing the chromosomal DNA on a slide. The sodium hydroxide/ethanol appears to prevent the loss of DNA from the slide.
Formamide Method. Place coplin jar containing 70% formamide/2X SSC in a waterbath and heat to 70°. Incubate slides 2-6 min. Dehydrate slides in an ethanol series and air dry.
Sodium Hydroxide/Ethanol Method. Warm coplin jar containing 1 M NaOH to 37°. Incubate slides 2-6 min and dehydrate in an ethanol series. Air dry.
3. The amount of probe needed is calculated by considering the concentration to be used (usually 50-100 ng/ml), the number of slides needed, and the amount of probe per slide (80-100 fil). The final concentration of the hybridization mixture (pH 7.0 to 7.2) is 50% formamide, 2X SSC, 0.04 M sodium phosphate, pH 6.0, carrier salmon sperm DNA (10 ¿ig/ml) in 500 to 1000-fold excess over the probe DNA, and tritiated probe. The hybridization mixture is placed in a boiling waterbath for 5 min to denature the probe immediately before applying to the slides. Cool quickly in a dry ice-ethanol bath.
4. Hybridization mixture (80-100 fil) is added to each slide and covered with a 24 X 60 mm siliconized coverslip. The slides are incubated 18-48 hr at 37° in a humidified chamber saturated with 2X SSC.
5. The washing procedure is initiated by dipping the slides in 50% formamide/2X SSC (pH 7.0) at 39° to remove coverslips. The slides are then washed in Wheaton dishes: 4 washes of 50% formamide/2X SSC for 5 min at 39°, 4 washes in 2X SSC for 5 min at 39°, 3 washes of 2X SSC at room temperature for 5 min. Dehydrate the slides by an ethanol series (70, 80, and 95%) for 2 min each. Air dry slides.
Kodak nuclear track emulsion NTB2 is melted at 43-45° and diluted 1:1 with water using a safelight with a Kodak dark red filter (No. 2, cat. 152 1723). The emulsion is aliquoted into clean new 20-ml glass scintillation vials which are stored in light-proof containers at 4° until needed. Approximately 12 aliquots of 20 ml can be made from one bottle of diluted NTB2 emulsion.
To coat the slide warm an aliquot of emulsion to 43°. Pour warmed emulsion into a clean slide mailer box and place in a coplin jar. Dip slides one at a time and place slides in a slide rack. Place the racks into a light-tight, humid drying box. After drying (usually overnight), load coated slides into light-tight boxes and wrap with aluminum foil. Store slide boxes at 4° for 10-21 days.
While using the safelight with a dark red filter for illumination, develop slides in D-19 diluted 1:1 with water for 2 min at 15 °. The slides are placed in stop bath for 30 sec, fixed in Kodak fixer for 5 min, and rinsed in distilled water for 15 min. Scrape emulsion from the back of the slide with a razor blade and allow the slides to air dry.
Chromosome Preparation Method 1: Giemsa/Hoeschst 33258 Staining of Elongated Chromosomes. First stain the slides in Hoeschst fluorescent dye H 33258 (1 jug/ml in 2X SSC)at 15°, then rinse with distilled water and expose to long-wavelength UV light for 1 hr. The slides are exposed to the UV light covered with 2X SSC at a distance of 20 cm. After rinsing and air drying stain the slides in 7% Giemsa in 0.2 M phosphate buffer, pH 6.8, rinse in distilled water and air dry (see Fig. 2).
Chromosome Preparation Method 2: Wright Stain. Combine 1 ml Wright stain with 3 ml phosphate buffer (0.06 M), mix, and pour onto slide. Stain 5-10 min and then rinse with water. Air dry. Insufficient rinsing will result in particulate matter that has the appearance of silver
Fig. 2. Metaphase spread of elongated chromosomes (approximately 1000 bands) hybridized to a single copy probe located at chromosome 3ql2. The chromosomes were prepared using chromosome method 1 and the Giemsa/Hoescht 33258 staining method. Right, metaphase field focused on chromosome bands; left, focused on silver grains.
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