bayani2004

Fluorescence In Situ Hybridization
(FISH)
UNIT 22.4
As early as 1988 (Lichter et al., 1988), FISH was used to visualize labeled DNA probes hybridized to chromosome and interphase nuclei preparations. The improvement of cloned
DNA sources, antibodies, fluorochromes, microscopy and imaging equipment, and software has permitted a variety of scientific investigations.
This unit is divided into five parts—probe preparation, slide preparation, hybridization,
post-hybridization washes, and interpretation. Probes are prepared by nick translation (see
Basic Protocol 1) or degenerative oligonucleotide primer PCR (DOP-PCR; see Alternate
Protocol 1) using hapten- or fluorochrome-labeled nucleotide. The amount of hapten
label incorporated is quantified by dot blotting (see Support Protocol 1). Cytogenetic
slide preparations are suitable for FISH (see Basic Protocol 2) if they are appropriately
aged (see Support Protocol 2); these slides may be used even if they have previously been
G-banded (see Alternate Protocol 2). In addition, slides made from paraffin-embedded
tissues are suitable specimens for hybridization with DNA (see Alternate Protocol 3) or
protein nucleic acid (PNA) probes (see Alternate Protocol 4). Hybridization conditions
must be appropriate for both the sample and probe materials: cytogenetic slides can be
hybridized with DNA (see Basic Protocol 3) and PNA (see Alternate Protocol 5) probes,
as can paraffin sections (see Alternate Protocols 6 and 7). Post hybridization wash and
detection conditions vary depending on the probe—indirectly labeled DNA probes (see
Basic Protocol 4), directly labeled DNA probes (see Alternate Protocol 8 or 9), and
peptide nucleic acid (PNA) probes (see Alternate Protocol 10).
NOTE: Coplin jars containing solutions at temperatures other than room temperature
should be prewarmed and maintained in a water bath at the specified temperature.
LABELING DNA PROBES FOR FISH
This section describes methods for labeling probes for fluorescence in situ hybridization
(FISH) analysis. The first protocol (see Basic Protocol 1) describes DNA labeling by nick
translation either indirectly with a hapten (biotin or digoxigenin) or directly with a fluorochrome (APPENDIX 1E) conjugated to dUTP. This method is appropriate for most DNAs
listed in Table 22.4.1; however, smaller DNAs, such as cDNAs, may require labeling by
PCR methods, which are also described (see Alternate Protocol 1). The choice of label,
whether direct or indirect, is based on the type of DNA probe, its size, and the application. Small probe inserts (such as cDNAs) should be labeled with a hapten, enabling the
option of signal amplification. Larger inserts can be both directly or indirectly labeled
(Table 22.4.1). The probe’s signal strength can vary depending on the application and
sequence specificity (Table 22.4.2). Furthermore, the choice of label or fluorochrome will
also depend on the availability of conjugated antibodies, the nature of the experiment,
and the availability of filters for the microscope utilized for visualization. A list of fluorochrome properties can be found in Table 22.4.3. For both labeling methods discussed
here, assessment of probe labeling is determined by size, gel electrophoresis, and for
indirectly labeled probes, dot blot analysis of incorporation (see Support Protocol 1).
The labeled DNA is ethanol precipitated in the presence of excess unlabeled DNAs,
which serve as carriers (sonicated salmon sperm DNA) and as suppressors of repetitive
sequences (Cot-1 DNA). The final product is resuspended in a hybridization buffer and
is then ready for use in FISH experiments (see Hybridization). The labeling procedures
outlined can be applied to all species of DNA.
Contributed by Jane Bayani and Jeremy A. Squire
Current Protocols in Cell Biology (2004) 22.4.1-22.4.52
C 2004 by John Wiley & Sons, Inc.
Copyright Cell Biology of
Chromosomes
and Nuclei
22.4.1
Supplement 23
Table 22.4.1 Sources of DNA Probes for FISH Analysisa
Type
Application
Amplification of
signal required
Labeling methods
applicable
Indirect or
direct labeling
Plasmids (10–20 kb insert)
Locus specific
Yes
Nick translation and PCR Indirect
Cosmids (15–30 kb insert)
Locus specific
Yes
Nick translation and PCR Indirect
PACs (∼100 kb insert)
Locus specific
No
Nick translation and PCR Both
BACs (∼200 kb insert)
Locus specific
No
Nick translation and PCR Both
YACs (300–1.5 Mb insert)
Locus specific
No
Nick translation and PCR Direct
Genomic DNA from flow sorted
chromosomes
Chromosome
paints
No
PCR labeling
Both
Genomic DNA from
microdissected DNA
Locus specific
Yes
PCR labeling
Both
a Shown are the most common types of DNAs used as probes for FISH analysis. Many of these DNAs can be labeled by nick translation (see Basic
Protocol 1) or PCR (see Alternate Protocol 1). For probe inserts <2.0 kb, labeling by PCR is recommended.
Table 22.4.2 Different Classes of Commonly Used Probes for FISH Analysisa
Type
Fluorescence intensity
Typical applications
Centromere probes
Strong
Ideal probe for the beginner to learn basic FISH
techniques. Commonly used to enumerate chromosomal
monosomies and trisomies, and for sex determination in
transplantation studies.
Subtelomere specific probes
Moderate/weak
Used for determining whether small terminal
rearrangements near telomeres have taken place.
Chromosome paints
Strong/moderate
Useful for identifying small marker chromosomal
aberrations where a specific chromosome is suspected.
Also useful in confirming SKY or MFISH findings.
Translocation junction
unique-sequence probes
Moderate
Used for detecting the presence of specific translocations in
interphase cells or in metaphase spreads.
Microdeletion
unique-sequence probes
Moderate
Identification of small submicroscopic deletions using
metaphase preparations.
Probes detecting gene
amplification
Strong
For detecting oncogene copy number increases (gene
amplification) in interphase and metaphase cells
a Shown are the most common types of commercially available probes used in FISH experiments and their typical applications. Signal strength will
vary depending on the size and application of the probe as well as the target DNA specimen.
Another choice for FISH analysis is a peptide nucleic acid (PNA) probe. These probes also
bind DNA in a sequence-specific manner but are created by oligonucleotide synthesis. The
fabrication of PNA probes is not easily accomplished in a typical molecular cytogenetic
laboratory, so they are currently only available commercially (see Internet Resources).
BASIC
PROTOCOL 1
Fluorescence
In Situ
Hybridization
(FISH)
Labeling FISH Probes by Nick Translation
This protocol, adapted from Beatty et al. (2002), outlines the basic steps for either directly or indirectly labeling DNA probes for FISH analysis. Once labeled probes are
ethanol precipitated, they are ready for final preparation prior to hybridization to the
target specimen.
Materials
DNA probe (i.e., cosmid, plasmid, PAC, BAC, or YAC; Table 22.4.1)
10× nick translation buffer (see recipe)
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Table 22.4.3 Fluorescent Properties of Labels and DNA Stains Used for FISHa
Label
Excitation (nm)
Emission (nm)
Digoxigenin
NA
NA
Biotin
NA
NA
Alexa 488c
490
520
c
525
550
c
555
570
c
Alexa 594
590
615
Amino-methyl coumarin
399
445
Cascade Blue
Haptens
b
Fluorochromes
Alexa 532
Alexa 546
400
420
d
489
506
d
550
570
d
649
670
d
Cyanine 7
743
767
Fluorescein isothiocyanate (FITC)
495
523
Rhodamine B
Cyanine 2
Cyanine 3
Cyanine 5
560
580
e
433
480
e
655
675
497
524
530
555
Spectrum Orange
559
588
Spectrum Rede
587
612
Tetramethylrhodamine isothiocyanate
(TRITC)
550
570
Texas Red
595
610
Chromomycin A3
430
570
4 ,6-Diamindine-2-phenylindole (DAPI)
538
461
Ethidium bromide
518
615
Hoechst 33258 (bis-benzimide)
352
461
Propidium bromide
535
617
Spectrum Aqua
Spectrum FRed
e
Spectrum Green
e
Spectrum Gold
e
DNA stains
a Shown are the commonly used fluorochromes and DNA stains used in FISH analysis. Of these, DAPI is
the most widely used.
b Haptens, such as digoxigenin and biotin are detected with antibodies conjugated to one of the fluorochromes
listed.
c The Alexa family of fluorochromes can be purchased from Molecular Probes.
d The Cyanine family of fluorochromes can be purchased from Amersham Biosciences.
e The Spectrum family of fluorochromes can be purchased from Vysis.
DNase I dilution buffer (see recipe)
1 mM dNTP mixture (see recipe)
Fluorochrome/dTTP or hapten/dTTP mixture (see recipes)
3 mg/ml DNase I (see recipe)
10 U/µl E. coli DNA polymerase I (Roche)
5 × loading dye (see recipe)
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2% (w/v) agarose gel (see recipe)
100-bp DNA ladder
Sonicated salmon sperm DNA standards (i.e., 12.5, 25.0, and 500 ng/µl; see recipe)
1× TBE buffer (see APPENDIX 2A)
300 mM EDTA (APPENDIX 2A)
10 mg/ml sonicated salmon sperm DNA (Invitrogen)
1 µg/µl human or mouse Cot-1 DNA (Invitrogen)
3 M sodium acetate (APPENDIX 2A)
100% ethanol
70% ethanol, cold
Hybridization buffer (see recipe)
Water bath or PCR machine
0.5-ml PCR tube (optional)
Additional reagents and equipment for agarose gel electrophoresis (APPENDIX 3A)
and determination of hapten incorporation by dot blot analysis (see Support
Protocol 1)
Prepare equipment and reagents
1. Set water bath or PCR machine to 15◦ C.
2. Put DNA probe and all labeling reagents (i.e., 10× nick translation buffer, DNase I dilution buffer, 1 mM dNTP mixture, and fluorochrome/dTTP or hapten/dTTP mixture)
on ice.
3. Prepare 10× DNase I solution (1.0 ml total) by combining 1.0 µl of 3 mg/ml DNase
I and 999.0 µl DNase I dilution buffer. Place on ice.
Perform labeling reaction
4. For each DNA to be labeled, prepare the following mixture in a 1.5-ml microcentrifuge
or 0.5-ml PCR tube, adding the 10× DNase I solution and E. coli polymerase I last:
2 µg DNA probe
10.0 µl 10× nick translation buffer
5.0 µl 1 mM dNTP mixture (i.e., dATP, dCTP, dGTP)
4.0 µl fluorochrome/dTTP or hapten/dTTP mixture
10.0 µl 10× DNase I solution
2.5 µl E. coli DNA polymerase.
Adjust the volume to 100 µl with water.
The amount of DNase I solution added will vary (see Commentary).
A probe prepared using hapten/dTTP is indirectly labeled, while one prepared with
fluorochrome/dTTP is directly labeled.
5. Incubate 90 to 120 min at 15◦ C (see Table 22.4.22 and Critical Parameters).
6. After incubation, place tubes on ice.
Determine size of products from labeling reaction
7. Transfer 10 µl labeling reaction to a fresh tube and mix with 4 µl of 5× loading dye.
Load onto a 2% agarose gel (APPENDIX 3A) along with a 100-bp DNA marker and 10 µl
of each salmon sperm concentration standard.
Fluorescence
In Situ
Hybridization
(FISH)
The salmon sperm standards are used to help determine the amount of labeled DNA probe,
as well as to act as a molecular-size marker.
8. Electrophorese at 100 V for ∼20 min or until the dye front is two thirds of the way
through the gel.
22.4.4
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Figure 22.4.1 Nick translated DNA fragments electrophoresed on a 2% agarose gel. A volume of
10.0 µl labeled product was loaded. The fragment sizes range from 200 to 500 bp as determined
from the molecular marker. The estimated concentration of the probe is 200 ng in 10 µl, yielding
∼20 ng/µl.
The final fragment sizes should be between 200 and 500 bp, with ∼200 ng labeled DNA
present in the gel, assuming the original amount of labeled probe was 2 µg. This can be
ascertained using the sonicated salmon sperm concentration standards (Figure 22.4.1).
9. If the fragment sizes are too large, return the labeling reaction to 15◦ C for 20 to 30 min
(or time as required) and reassess the size of the fragments (i.e., repeat steps 5 to 8).
It may be necessary to spike the labeling reaction with additional DNase I solution and
E. coli DNA polymerase I. If the fragments are too small, labeling (step 4) will need to be
repeated with an adjustment in the amount of DNase I added to the labeling reaction. If
there is no significant change in fragment size following further digestion, the investigator
should start again and review Table 22.4.22 for possible reasons of insufficient digestion.
Determine amount of labeled probe
10. Once the proper labeling size has been achieved, add 10 µl of 300 mM EDTA to the
labeling reaction to stop the action of the enzymes.
11. Estimate the amount of labeled probe on the gel using the salmon sperm standards
as a guide. Calculate the amount of labeled DNA remaining in the tube.
Be sure to consider the volumes removed from the starting volume for loading onto the
gel(s) when calculating the amount of labeled DNA left in the tube.
Based on the concentration standards, the labeled probe in Figure 22.4.1 appears to
be somewhere between 125 ng/10 µl (1.25 ng/µl) and 250 ng/10 µl (25.0 ng/µl). A
conservative estimate is 200 ng/10 µl (20 ng/µl). Thus, from an original 100 µl starting
volume, 10 µl was removed for a gel, leaving 90 µl. If the estimated concentration is
20 ng/µl, the remaining labeled probe is 1800 ng (20 ng/µl × 90 µl = 1800 ng).
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Add carrier and supressor of repetitive sequences
12. For each 1 µg labeled DNA in the tube, add 5 µl of 10 mg/ml sonicated salmon sperm
DNA (50 µg).
For example, for 1.8 µg DNA, add ∼100 µg sonicated salmon sperm DNA.
13. For each microgram labeled DNA in the tube, add 5 to 10 µg human or mouse
Cot-1 DNA.
For the example given above (1.8 µg labeled DNA), ∼20 µg Cot-1 DNA is required
The amount of Cot-1 used is dependent on the number and extent of repeat elements in
the DNA insert used as the probe. Generally, cDNA probes will have fewer repeats than
genomic probes, so they will require relatively less Cot-1 suppression. BAC and YAC
probes may need more suppression.
Determine incorporation
14. If the DNA probe has been labeled with hapten, check incorporation by dot blot
analysis (see Support Protocol 1).
Ethanol precipitate DNA
15. Add 0.1 vol. of 3 M sodium acetate followed by 2.5 vol of 100% ethanol.
16. Incubate overnight at −20◦ C or 2 to 3 hr at −80◦ C.
The recovery of DNA during the precipitation process can be increased by lengthening the
time at −20◦ C or −80◦ C.
17. Microcentrifuge 20 min at 13,000 rpm, 4◦ C.
Resuspend DNA
18. Carefully remove the supernatant and wash the pellet with cold 70% ethanol.
19. Microcentrifuge 20 min at 13,000 rpm, 4◦ C.
20. Remove the ethanol and allow the pellet to air dry.
21. Resuspend the pellet in hybridization buffer to a final concentration of 10 ng/µl. Store
labeled probe at −20◦ C up to several months (directly labeled probe) or several years
(indirectly labeled probe).
The final concentration should be based on the labeled DNA content, not the total DNA
content, which includes the unlabeled Cot-1 and salmon sperm DNA. Thus, for a labeled
DNA amount of 1.8 µg, 180 µl hybridization buffer should be added to reach a final
concentration of 10 ng/µl.
The labeled probe can now be used for FISH experiments.
The amount of labeled probe used will depend on the area of the slide to be hybridized.
For example, 10 µl is sufficient to cover a 22 × 22–mm coverslip, 20 µl is sufficient to
cover a 22 × 30–mm coverslip, and 30 µl is sufficient to cover a 22 × 50–mm coverslip.
ALTERNATE
PROTOCOL 1
Labeling Probes for FISH by Degenerative Oligonucleotide Primer Polymerase
Chain Reaction (DOP-PCR)
This protocol describes direct and indirect labeling procedures using PCR labeling. The
user has various choices in the primer sets. These may be specific primers for a region,
specific primers for vector sequences, universal primers, or DOP primers.
Additional Materials (also see Basic Protocol 1)
Fluorescence
In Situ
Hybridization
(FISH)
DNA template (Table 22.4.1)
10× PCR buffer: 100 mM Tris·Cl, pH 8.3 (APPENDIX 2A)/500 mM KCl (store up to
several months at −20◦ C)
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2 mM dNTP mixture (i.e., dATP, dCTP, dGTP; see recipe)
5 µM each primers 1 and 2 (APPENDIX 3F)
50 mM MgCl2
5 to 10 U/µl Taq DNA polymerase
Water, sterile
Mineral oil, sterile
1. For each labeling reaction, mix the following:
1 to ng DNA template
5.0 µl 10× PCR Buffer
4.0 µl 2 mM dNTP
2.0 µl 1 mM fluorochrome/dTTP or hapten/dTTP
1.5 µl 5 µM primer 1
1.5 µl 5 µM primer 2
1.5 µl 50 mM MgCl2
0.5 µl 5–10 U/µl Taq DNA polymerase.
Adjust final volume to 50.0 µl with sterile water. Overlay the mixture with sterile
mineral oil and put on ice until ready to place in the thermocycler.
The 10× PCR buffer is often supplied with the enzyme.
2. Place in a thermocycler and program the following steps:
Initial step:
35 cycles:
Final step:
Hold:
5 min
30 sec
30 sec
90 sec
5 min
indefinitely
93◦ C
94◦ C
55◦ C
72◦ C
72◦ C
4◦ C.
(denaturation)
(denaturation)
(annealing)
(extension)
(extension)
3. Mix 5 µl labeled reaction with 1 µl of 5× loading dye and run on a 2% (w/v) agarose
gel with molecular weight markers and sonicated salmon sperm DNA concentration
standards as described (see Basic Protocol 1, steps 7 and 8).
The final PCR product should fall between 200 and 500 bp.
4. Calculate the amount of labeled DNA remaining in the tube, add carrier and supressor DNA, precipitate, and resuspend in hybridization buffer as described (see Basic
Protocol 1, steps 11 to 20).
5. If the DNA probe has been labeled with hapten, check incorporation by dot blot
analysis (see Support Protocol 1).
Determination of Hapten Incorporation by Dot Blot Analysis
Gel electrophoresis of the labeled DNA FISH probe allows the investigator to determine
whether the DNA has been nicked (see Basic Protocol 1) or amplified (see Alternate
Protocol 1) to the appropriate sizes. It does not, however, allow determination of whether
the hapten has been incorporated into the DNA. Using a dot blot assay, the degree of
hapten incorporation can be assessed to determine the efficiency of the labeling protocol
and the activity of the DNA polymerase I or Taq DNA polymerase. This method is useful
in determining whether the lack of FISH signal is due to a poorly labeled probe or to
hybridization or slide preparation factors. This method involves spotting labeled product
onto a nylon filter along with a control. The labeled DNA is detected by immunolabeling
and colorimetric visualization.
SUPPORT
PROTOCOL 1
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Chromosomes
and Nuclei
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Current Protocols in Cell Biology
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Materials
100 mM Tris·Cl (pH 7.5)/15 mM NaCl (APPENDIX 2A)
Indirectly labeled probe (i.e., with biotin or digoxigenin; Table 22.4.3)
Control DNA labeled with biotin or digoxigenin (i.e., labeled DNA probe known to
produce good signal strength)
0.5% (w/v) BSA in 100 mM Tris·Cl, pH 7.5 (APPENDIX 2A)/15 mM NaCl (store up to
6 months at 4◦ C)
AP-labeled antibody mixture (see recipe)
100 mM Tris·Cl (pH 9.5)/100 mM NaCl/50 mM MgCl2 (APPENDIX 2A)
NBT/BCIP (see recipe)
Charged nylon membrane (∼5 × 5 cm per sample)
Filter paper
37◦ C dry oven
Rotating platform
Prepare and load nylon membrane
1. Cut a charged nylon membrane to a size appropriate for the number of samples.
2. Soak the membrane 5 min in 100 mM Tris·Cl (pH 7.5)/15 mM NaCl, room temperature. Blot the membrane with filter paper.
3. Pipet aliquots of indirectly labeled probe and control DNA labeled with biotin or
digoxigenin in a dilution series (e.g., 200, 100, 50 ng) onto the membrane, leaving
ample space between spots. Incubate 5 to 10 min, room temperature.
Block and label membrane
4. Incubate the membrane 1 min in 100 mM Tris·Cl (pH 7.5)/15 mM NaCl, making sure
it is saturated.
5. Incubate the membrane 30 min in 0.5% (w/v) BSA in 100 mM Tris·Cl (pH 7.5)/15 mM
NaCl at room temperature.
6. Transfer the membrane to an alkaline phosphatase–labeled antibody mixture in a
plastic container and incubate 30 min in a 37◦ C dry oven on a rotating platform,
making sure the membrane is completely saturated.
Develop dot blot
7. Remove the membrane and wash 15 min with 100 mM Tris·Cl (pH 7.5)/15 mM NaCl.
8. Remove the membrane and wash 2 min with 100 mM Tris·Cl (pH 9.5)/100 mM
NaCl/50 mM MgCl2 , room temperature.
9. Remove the membrane and incubate 5 to 10 min in NBT/BCIP solution under dimmed
lighting until the blot develops.
10. Wash the membrane with water and air dry.
The membrane can be stored at room temperature. Over time the intensity of the chemical
stain will fade. Typical results are shown in Figure 22.4.2.
TARGET SLIDE PREPARATION
Fluorescence
In Situ
Hybridization
(FISH)
Slide quality can greatly influence the success of FISH assays. When slide preparations
are less than optimal, treatment with a protease can decrease the risk of high background
and increase access of the probe to the DNA target by removing protein barriers to allow
efficient hybridization. FISH carried out on paraffin sections requires more aggressive
22.4.8
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Figure 22.4.2 Dot-blot analysis of biotin incorporation in probes shown in Figure 21.4.1. Compared to control labeled DNA, the labeled probes appear to have incorporated biotin well, with
probe 4 showing the greatest incorporation compared to probe 1. The concentration of each probe
was estimated at 20 ng/µl; however, it is evident that the labeling efficiency was greater in some
probes over others.
protease treatment and assessment. Following optimal slide pretreatment, the specimen
is subjected to denaturation, causing double-stranded DNA to become single stranded.
This will permit the efficient hybridization of the denatured probe to the DNA target on
the slide.
Preparation of Cytogenetic Specimens for FISH
The following protocol describes the pretreatment of cytogenetic slides (UNIT 22.2) for
FISH analysis with DNA probes (in-house or commercial) or commercially obtained
PNA probes. In this protocol, pepsin is recommended for use as the protease; however,
proteinase K is also often used. The concentration and conditions for proteinase K treatment are also noted in the protocol. The treatment with pepsin will vary depending on the
extent of digestion required on the slide. In some cases, pepsin treatment is not needed.
BASIC
PROTOCOL 2
Materials
Cytogenetic slide preparation (UNIT 22.2): age naturally at least 2 days at room
temperature or artificially (see Support Protocol 2)
10% (w/v) pepsin—100 mg pepsin powder (Sigma) in 1.0 ml H2 O; store in 20-µl
aliquots up to several months at −20◦ C—and 0.01 M HCl, 37◦ C or
14 mg/ml proteinase K (Roche Diagnostics) and proteinase K buffer—i.e., 20 mM
Tris·Cl, pH 7 (APPENDIX 2A)/0.2 mM CaCl; store up to several months at room
temperature
1× PBS (APPENDIX 2A)
70% formamide/2× SSC (pH 7.0), 72◦ C (see UNIT 18.6 for SSC): prepare fresh
70% ethanol, ice-cold (for DNA probes)
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Table 22.4.4 Coplin Jars Needed for Preparation of Cytogenic Specimens for FISHa
No. Coplin jars
1
Contents
Temperature
70% ethanol
1
70% ethanol
1 or 2b
b
1 or 2
Room temperature
4◦ C
b
80% ethanol
Room temperature
100% ethanol
Room temperature
b
72◦ C
1
70% formamide in 2× SSC
1
1× PBS
Room temperature
1
Pepsin/0.01 HCl or proteinase
K/buffer
37◦ C or room temperature
(respectively)
a See Basic Protocol 2.
b For DNA probes only.
Phase contrast microscope (UNIT 4.1)
Coplin jars
Additional reagents and equipment for hybridization (see Basic Protocol 3 and
Alternate Protocol 5)
CAUTION: Formamide is a carcinogen and should be handled with care. Discard according to biohazard rules of the institution.
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.4.
Detect and remove cytoplasmic contamination by protease treatment
1. Using a phase-contrast microscope, determine the extent of cytoplasmic residue on
the cytogenetic slide preparation. If there is no cytoplasm, proceed to step 6.
2a. For pepsin digestion: Add 10 to 15 µl of 10% (w/v) pepsin to 50 ml warm 0.01 M
HCl in a Coplin jar. Incubate the slides in the protease 5 min at 37◦ C.
2b. For proteinase K digestion: Dilute 14 mg/ml proteinase K to a final concentration of
0.1 µg/ml in proteinase K buffer in a Coplin jar. Incubate the slides in the protease
6.5 min at room temperature.
Remove protease and determine efficacy
3. Wash the slide 5 min at room temperature in a Coplin jar containing 1× PBS.
4. Pass the slide through a dehydrating series of 70%, 80%, and 100% ethanol in Coplin
jars for 5 min each. Allow to air dry after the final ethanol treatment (∼5 min).
5. View the slide using phase-contrast microscopy to determine the extent of protein
digestion. If necessary, repeat the protease step.
Prepare slides for hybridization
6a. For PNA probes: Proceed to hybridization (see Alternate Protocol 5).
6b. For DNA probes: Denature the slide for the recommended time (see Table 22.4.5) in
70% formamide/2× SSC (pH 7.0), 72◦ C.
Fluorescence
In Situ
Hybridization
(FISH)
The time will vary according to the age and quality of the slide.
7b. Promptly place the slide into a Coplin jar containing ice-cold 70% ethanol for 5 min.
Pass the slides through a dehydration series of 80% and 100% ethanol for 5 min each.
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Table 22.4.5 Formamide Denaturation Guide for Cytogenetic Specimensa
Slide criteria
Suggested
denaturation timeb
Fresh slide stored 1–2 days at room temperature
60–90 sec
Fresh slide artificially aged 1–2 days
60–90 sec
Aged slide stored 1–2 weeks at room temperature
1.5–2 min
Aged slide stored 2–4 weeks at room temperature
2 min
Aged slides stored 1–2 months
2–3 min
Previously G-banded slides >1-week old
30–40 sec
Previously G-banded slides <1-week old
20–30 sec
a Shown are general guidelines for denaturation of slides of varying ages and conditions. The investigator
should pay attention to the slide quality and monitor the changes in denaturation conditions as the slides age
or are processed. Each laboratory will possess different environmental conditions as well as slide making
procedures, which will affect the denaturation procedures significantly.
b Denaturation conditions are 72◦ C in 70% formamide/2× SSC.
8b. Air-dry the slide after the final ethanol treatment and proceed to hybridization (see
Basic Protocol 3).
Artificial Aging of Cytogenetic Slide Preparations for FISH
The chromosomes on freshly prepared slides are often too fragile for immediate use—
i.e., the high temperatures used during the slide denaturation procedure can damage the
DNA, making it less optimal for hybridization. Such slides require aging, which can be
achieved by allowing them to naturally age a few days at ambient temperature; however,
an experiment must occasionally be performed immediately. The protocol below outlines
a method for artificially aging freshly prepared slides so that FISH results are available
within 12 to 24 hr after preparation from the cytogenetic suspension. This method may
also be used when the relative humidity in the laboratory is high due to local weather
conditions.
SUPPORT
PROTOCOL 2
To age, incubate a freshly prepared cytogenetic slide (UNIT 22.2) at least 1 hr (up to 3 hr)
in a Coplin jar containing 2× SSC (UNIT 18.6), 37◦ C. Remove the slide and pass through
a dehydration series of 70%, 80%, and 100% ethanol for 5 min each, and air dry (5 to
10 min). Proceed with enzyme digestion if required.
Note that a list of all Coplin jars used in this protocol is given in Table 22.4.6.
Preparation of Previously G-Banded Cytogenetic Specimens for FISH
The following method outlines the steps involved in pretreating a previously Giemsa
(G)-banded cytogenetic slide (UNIT 22.3) for subsequent use in FISH analysis. The slides
must be completely free of any oils and must also be destained. Since the slide has already
been treated with a protease, it will require no additional protease digestion; however,
stripping the proteins, which are protective to the DNA, will cause it to become much
more sensitive to degradation during the denaturation process. The ability to re-use a
previously banded slide for FISH will be dependent on the age of the slide, the degree of
trypsin digestion during the banding, and whether residual immersion oils or mounting
buffers have degraded the DNA.
ALTERNATE
PROTOCOL 2
Materials
Banded cytogenetic slide specimen (UNIT 22.3)
Xylene
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Table 22.4.6 Coplin Jars Needed for Artificial Aging of Cytogenetic Slide
Preparations for FISHa
No. Coplin jars
Contents
Temperature
1
70% ethanol
Room temperature
1
80% ethanol
Room temperature
1
100% ethanol
Room temperature
1
2× SSC
37◦ C
a See Support Protocol 2.
Table 22.4.7 Coplin Jars Needed for Preparation of Previously G-Banded
Cytogenetic Specimens for FISHa
No. Coplin jars
Contents
Temperature
1
70% ethanol
4◦ C
1
70% ethanol
Room temperature
2
80% ethanol
Room temperature
2
100% ethanol
Room temperature
1
70% formamide/2× SSC,
pH 7.0
72◦ C
1
Methanol
Room temperature
1
b
Xylene
Room temperature
a See Alternate Protocol 2.
b CAUTION: Maintain in a fume hood.
Methanol
70%, 80%, and 100% ethanol
70% formamide in 2× SSC (pH 7.0; UNIT 18.6), 72◦ C: prepare fresh
70% ethanol, ice-cold
Coplin jars
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.7.
1. Remove residual oils from previously banded cytogenic slide specimen by incubating
5 min in a Coplin jar containing xylene under a vented chemical hood.
2. Destain by incubating the slide ∼5 to 10 min in a Coplin jar containing roomtemperature methanol.
3. Pass the slide through a dehydrating series of 70%, 80%, and 100% ethanol for 5 min
each. Allow the slide to air dry (5 to 10 min) after the final ethanol treatment.
4. Denature slide 20 to 30 sec in 70% formamide/2× SSC (pH 7.0), 72◦ C.
The time will vary according to the age and quality of the slide (Table 22.4.5).
5. Promptly place the slide into a Coplin jar containing ice-cold 70% ethanol for 5 min
following denaturation.
Fluorescence
In Situ
Hybridization
(FISH)
6. Proceed through a dehydration series of 80% and 100% ethanol for 5 min each. Airdry the slide after the final ethanol treatment and proceed with hybridization (see
Hybridization).
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Preparation of Paraffin-Embedded Specimens for Hybridization with DNA Probes
FISH analysis on paraffin sections is becoming increasingly important in both diagnostic
and research laboratories. This protocol outlines the steps involved in pretreating paraffin
sections for FISH. While the preparation of paraffin-embedded specimens can be modified, the most critical component is the quality of the starting section (see Commentary).
Different histopathology laboratories will have various methods of tissue fixation which
should be taken into consideration when assessing the success of an experiment.
ALTERNATE
PROTOCOL 3
Materials
5- to 10-µM paraffin sections on silanized slides
Xylene
100% ethanol
0.5% (w/v) pepsin in 0.85% (w/v) NaCl (pH 1.5), 45◦ C (see recipe)
2× SSC, pH 7.0 (UNIT 18.6)
Propidium iodide (PI) or DAPI in antifade (see recipes)
70%, 80% and 100% ethanol
Coplin jars
45◦ C water bath
Fluorescence microscope (UNIT 4.2) with a FITC, and PI or DAPI filter set
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.8.
Remove wax
1. Add 5- to 10-µM paraffin sections on silanized slide to a Coplin jar containing
xylene. Gently agitate 5 min at room temperature. Transfer slides to a second Coplin
jar containing fresh xylene for an additional 5 min with gentle agitation.
2. Transfer the slide to a Coplin jar containing 100% ethanol. Soak 5 min, then transfer
slides to fresh 100% ethanol and soak another 5 min. Agitate 2 to 3 times during each
5-min period.
3. Remove the slide from ethanol and allow to air dry (5 to 10 min).
Perform protease digestion and counterstain
4. Place slide in a Coplin jar containing 0.5% (w/v) pepsin in 0.85% NaCl, 45◦ C.
Incubate 15 to 20 min in a 45◦ C water bath (see Table 22.4.5).
5. Rinse the slide 20 to 30 sec in a Coplin jar containing 2× SSC, pH 7.0.
Table 22.4.8 Coplin Jars Needed for Preparation of Paraffin-Embedded
Specimens for Hybridization with DNA Probesa
No. Coplin jars
Contents
Temperature
1
70% ethanol
Room temperature
1
80% ethanol
Room temperature
3
100% ethanol
Room temperature
1
0.5% pepsin in 0.85% NaCl
45◦ C
4
2× SSC, pH 7.0
Room temperature
2
b
Room temperature
Xylene
a See Alternate Protocol 3.
b CAUTION: Maintain in a fume hood.
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Table 22.4.9 Troubleshooting Guide for Pretreatment of Paraffin-Embedded Sections
Problem
Cause
Green autofluorescence Under-digestion with
upon inspection/or poor pepsin solution
uptake of counterstain
Solution
Remove coverslip and wash with 2× SSC
as described (see Alternate Protocol 3, step
5). Return the slide to pepsin digestion
buffer (see Alternate Protocol 3, step 4).
The time required for subsequent protease
treatment will be dependent on the existing
degree of digestion.
Should repeated attempts at digestion fail,
consider another enzyme such as proteinase
K (use at 10 mg/ml in 2× SSC for 20 min at
37◦ C) in place of the pepsin buffer (see
Alternate Protocol 3, step 4) or in
combination with the pepsin buffer.
Lifting of tissue from
slide
Holes in tissue or
“ghosty” appearance
Loosening by protease
treatment
Occasionally the protease treatment will
loosen the tissue from the slide. Try to work
carefully such that the tissue is relatively
undisturbed.
Glass slides not coated
with silane
Associated with this is the use of charged or
silanated slides. Many pathology labs will
use coated slides. Check if the sections
were mounted onto such slides. If not, try to
obtain sections on these slides.
Over-digestion
Over-digestion will be evident from the
degradation of the tissue. Start again and
decrease the tissue treatment.
Figure 22.4.3 Digestion of paraffin-embedded specimen for FISH. (A) Under-digestion is indicated by weak uptake of the DAPI stain. (B) Increased digestion permits better access to the DNA
and increase uptake of DAPI. This black and white facsimile of the figure is intended only as a
placeholder; for full-color version of figure, see color plates.
6. Apply 10 µl PI or DAPI in antifade to counter stain and coverslip.
The choice between PI or DAPI depends on the label color of the probe being analyzed.
Fluorescence
In Situ
Hybridization
(FISH)
7. View the slide using a fluorescence microscope with FITC, and PI or DAPI filter
set. Evaluate the tissue sections for under-digestion, appropriate digestion, or overdigestion using the guidelines in Table 22.4.9 and Figure 22.4.3.
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For certain types of tissue additional troubleshooting steps may also be necessary (see
Troubleshooting).
8. Carefully remove the coverslip and rinse slide three times in separate Coplin jars
containing 2× SSC, pH 7.0, for 5 min each, agitating 5 to 10 sec in each rinse.
9. Dehydrate slide in a series of 70%, 80%, and 100% ethanol washes for 1 min each.
10. Allow the slide to air dry. Proceed to hybridization with labeled DNA probes.
Preparation of Paraffin-Embedded Specimens for Hybridization with PNA Probes
PNA probes are also applicable to paraffin sections. Like pretreatment for hybridization
with DNA probes (see Alternate Protocol 3), pretreatment for hybridization with PNA
probes requires some minor modifications as described below.
ALTERNATE
PROTOCOL 4
Materials
5- to 10-µM paraffin sections on silanized slides
Xylene
100% ethanol
100 µg/ml RNase I (see recipe)
2× SSC (UNIT 18.6)
1 M sodium thiocyanate, 80◦ C
5 mg/ml pepsin in 0.85% (w/v) NaCl, 45◦ C (see recipe)
0.1 M triethanolamine, pH 8.0
Acetic anhydride (Sigma)
1× PBS (APPENDIX 2A)
70%, 80%, and 100% ethanol
45◦ C hot plate or slide warmer
Coplin jars
37◦ C oven
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.10.
Remove wax
1. Preheat 5- to 10-µm paraffin sections on silanized slide on a 45◦ C slide warmer or
hot plate for ∼30 min (i.e., until the wax melts).
Table 22.4.10 Coplin Jars Needed for Preparation of Paraffin-Embedded Specimens for Hybridization with PNA Probesa
No. Coplin jars
Contents
Temperature
1
70% ethanol
Room temperature
1
80% ethanol
Room temperature
4
100% ethanol
Room temperature
2
1× PBS
Room temperature
1
5 mg/ml pepsin in 0.85% NaCl
45◦ C
1
1 M sodium thiocyanate
80◦ C
5
2× SSC
Room temperature
1
Triethanolamine, pH 8.0
Room temperature
2
Water
Room temperature
3
b
Xylene
a See Alternate Protocol 4.
b CAUTION: Maintain in a fume hood.
Room temperature
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2. Dewax by transferring the slide to a Coplin jar containing 50 ml xylene and incubating
5 to 10 min at room temperature. Repeat twice with fresh xylene.
Time for dewaxing is dependent on many factors (e.g., tissue section thickness, surface
area, type of tissue).
3. Remove xylene by soaking slide 5 to 10 min in 100% ethanol. Repeat twice with
fresh ethanol.
4. Allow slides to air dry (5 to 10 min).
The tissue should turn white at this point.
Treat with RNase, SSC, and sodium thiocyanate
5. Add 40 µl of 100 µg/ml RNase I to the slide. Cover with a coverslip and incubate
1 hr at 37◦ C.
6. Wash twice in Coplin jars containing fresh 2× SSC for 5 min each time.
7. Incubate slide 8 min in a Coplin jar containing 1 M sodium thiocyanate, 80◦ C.
8. Rinse twice in Coplin jars containing fresh distilled water for 1 min each.
Digest with protease
9. Incubate 7 to 9 min at 45◦ C in a Coplin jar containing 5 mg/ml pepsin in 0.85% NaCl,
pH 1.5.
10. Wash twice for 1 min each in Coplin jars containing fresh 2× SSC.
Acetylate
11. Start acetylation by placing slide in a Coplin jar containing 0.1 M triethanolamine,
pH 8.0.
12. While stirring gently, slowly add 125 µl acetic anhydride to give a final concentration
of 0.25% (v/v). Incubate 10 min at room temperature.
The acetylation procedure is carried out to reduce the electrostatic binding of probe to
positive charges on the tissue, thereby reducing the background.
Wash and dehydrate
13. Rinse 5 min in a Coplin jar containing 1× PBS. Repeat the rinse using a fresh Coplin
jar of PBS.
Washes from this point forward can be reduced in duration if adherence of tissue to the
slide is poor.
14. Rinse 5 min in a fresh Coplin jar containing 2× SSC.
15. Dehydrate through a series of 70%, 80%, and 100% ethanol and allow to air dry.
16. Proceed to hybridization with PNA probes (see Alternate Protocol 7).
HYBRIDIZATION
Fluorescence
In Situ
Hybridization
(FISH)
In the following section, hybridization of the probe to the slide is described. The prepared
DNA probe is denatured to a single-stranded state in the presence of denaturing buffer.
Depending on the probe type, there may be a preannealing step. The denatured probe
is applied to the denatured cytogenetic specimen and allowed to hybridize overnight.
Additionally, this section also describes the hybridization conditions for paraffin sections,
where the slide and probe are simultaneously denatured. If commercial DNA probes are
being used, the manufacturer’s instructions for probe denaturation are followed and the
hybridization conditions are adjusted accordingly. If a PNA probe is utilized, the probe
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Current Protocols in Cell Biology
is applied to the slide, and together the probe and target are denatured and permitted to
hybridize for at least 1 hr.
Hybridization of Labeled DNA Probes to Cytogenetic Specimens
Hybridization of a cyotgenetic specimen with the probe requires heat-denaturation of
the DNA probe in the hybridization buffer, and a pretreated and denatured slide specimen; together they are permitted to hybridize at 37◦ C. This protocol can be applied to
cytogenetic specimens that are unstained or were previously G-banded (see Alternate
Protocol 2).
BASIC
PROTOCOL 3
Materials
DNA probe (see Basic Protocol 1 and Alternate Protocol 1)
Pretreated and denatured cytogenetic slide specimen (see Basic Protocol 2 and
Support Protocol 2)
Rubber cement
75◦ C water bath or PCR machine
37◦ C dry oven
Glass coverslips
Hybridization box, slightly dampened (e.g., black plastic video tape box lined with
slightly moistened gauze or paper towel)
Hybridization container (i.e., any plastic container with lid or a black video cassette
box)
1. Heat-denature the required amount of labeled DNA probe 5 min in a 75◦ C water bath
or PCR machine. Preanneal 1 hr in a 37◦ C dry oven (see Critical Parameters).
2. Add the appropriate amount of DNA probe to the pretreated and denatured cytogenetic
slide specimen using guide given in Table 22.4.11. Coverslip and ring the perimeter
with rubber cement to seal in place.
Table 22.4.11 Guide to Appropriate Probe Volume
for a Given Coverslip Size
Probe volume
Coverslip size (mm)
10 µl
22 × 22
20 µl
22 × 30
30 µl
22 × 50
Table 22.4.12 Examples of Minimal Hybridization Times Required for Representative
DNA Probes
Minimal hybridization
time (37◦ C)
Probe type (300–500 bp)
Centromere Probesa
4 hr
a
Subtelomere chromosome-specific probes
24 hr
Chromosome paints
4 hr
Translocation junction unique-sequence probes
24 hr
Microdeletion unique-sequence probes
24 hr
Probes detecting gene amplification
24 hr
a Protein nucleic acid (PNA) probes, which are commercially available, require a minimum of 60 min for
hybridization.
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3. Place slide in a slightly dampened hybridization box.
4. Transfer slide to a hybridization container and incubate overnight in a 37◦ C dry oven
(see Table 22.4.12).
5. Proceed to post-hybridization washes and immunofluorescent detection of indirectly
(see Basic Protocol 4) or directly labeled DNA probes (see Alternate Protocol 8)
ALTERNATE
PROTOCOL 5
Hybridization of PNA Probes to Cytogenetic Specimens
PNA probes, as discussed below (see Critical Parameters), can be commercially obtained.
Use of PNA probes requires minor modifications in the hybridization to both cytogenetic
specimens and paraffin-embedded sections, as described below.
Additional Materials (also see Basic Protocol 3)
Labeled PNA probe (Applied Biosystems)
Pretreated cytogenetic specimen, not denatured (Basic Protocol 2)
80◦ C oven, hot plate, or HYBrite (Vysis)
1. Add the required amount of labeled PNA probe to the pretreated, undenatured, cytogenetic specimen.
2. Coverslip and seal with rubber cement. Allow the rubber cement to set and dry.
3. Place the slide in an 80◦ C oven or HYBrite, or on an 80◦ C hot plate for 90 sec.
4. Remove the slide and place it into a hybridization box. Hybridize at least 1 hr at room
temperature.
5. Proceed to post-hybridization washes for specimens hybridized with PNA probes
(see Alternate Protocol 10).
ALTERNATE
PROTOCOL 6
Hybridization of Labeled DNA Probes to Sections from Paraffin-Embedded
Material
Unlike cytogenetic specimens, tissue sections from paraffin-embedded material require
higher denaturation temperatures and longer denaturation times. Hybridization to these
sections requires co-denaturation of the probe and tissue section simultaneously. All experiments using previously paraffin-embedded material and a DNA probe should include
a minimal 24-hr hybridization.
Additional Materials (also see Basic Protocol 3)
Deparaffinized and enzyme-digested specimen (see Alternate Protocol 3)
90◦ C oven, hot plate, or HYBrite (Vysis)
1. Apply DNA probe to the deparaffinized and enzyme-digested specimen using the
guide given in Table 22.4.11. Add a glass coverslip and apply rubber cement along
the perimeter of the coverslip. Allow the rubber cement to dry.
2. Denature the probe and target DNA simultaneously by placing the slide in a 90◦ C
oven or HYBrite, or on a 90◦ C hot plate for 12 min.
Fluorescence
In Situ
Hybridization
(FISH)
3. For slide denatured in an oven or on a hot plate: Transfer the slide to a hybridization
container lined with a wet paper towel or damp gauze, and incubate overnight in a
37◦ C dry oven.
4. For slide denatured in a HYBrite: Program the unit to hold at 37◦ C. Incubate overnight.
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5. Proceed to post-hybridization washes and immunofluorescent detection of indirectly
or directly labeled DNA probes (see Basic Protocol 4 or Alternate Protocols 8 or 9).
Hybridization of PNA Probes to Sections from Paraffin-Embedded Material
The following protocol describes hybridization of commercially available PNA probes to
pretreated sections from paraffin-embedded samples. Unlike DNA probes, PNA probes
require a minimum of 60 min for hybridization.
ALTERNATE
PROTOCOL 7
Additional Materials (also see Basic Protocol 3)
PNA probes (Applied Biosystems)
Deparaffinized and enzyme-digested specimen (see Alternate Protocol 4)
80◦ C oven, hot plate or HYBrite (Vysis)
1. Apply the appropriate amount of PNA probe (as suggested by the manufacturer) to the
deparaffinized and enzyme-digested paraffin section using the guidelines provided in
Table 22.4.11. Coverslip and seal with rubber cement. Allow the rubber cement to
dry.
2. Place in an 80◦ C oven or HYBrite, or on an 80◦ C hot plate for 3 min.
3. Place in a hybridization container and allow to hybridize at least 1 hr at room
temperature.
4. Proceed to post-hybridization wash for slides hybridized with PNA probes (see
Alternate Protocol 10).
POST-HYBRIDIZATION WASHES AND DETECTION
Following hybridization, unbound probes, whether DNA or PNA, must be removed from
the specimen. This is accomplished through washes of appropriate stringency, using
formamide and SSC in varying amounts. After immunodetection, final washes contain
detergents to remove unbound antibodies. The slides are counterstained and mounted in
an antifade medium for visualization. Variations on washing procedures reflect the nature
of the probe, whether directly or indirectly labeled.
Post-Hybridization Washes and Immunofluorescent Detection of Indirectly
Labeled DNA Probes
This protocol describes post-hybridization washes and detection of indirectly labeled
DNA probes. Following overnight hybridization, the coverslip is removed and the slide
is immersed in a solution of formamide to remove any unbound probe. The slide is then
washed in stringent SSC washes and blocked with a solution of BSA. Depending on the
type of DNA probe used, amplification of the signal may be required. This is achieved by
the addition of primary and secondary antibodies that may or may not be conjugated with
a fluorochrome. Detergent washes are carried out after each antibody incubation; it is
critical that the slide not be permitted to dry out at any point of the assay. The slide is then
counterstained and ready for visualization. In multicolor FISH experiments, the user may
have both indirectly and directly labeled probes on the same specimen. If this is the case,
it is important to keep the ambient light dim to prevent quenching of the signal; Coplin
jars with lids are useful for this purpose. If using multiple indirectly labeled probes, be
sure that the primary and secondary antibodies are raised in different animals such that
there is no cross-reaction. This protocol is applicable to cytogenetic slides or sections
from paraffin-embedded samples; however, more stringent or additional washes may be
required for paraffin experiments.
BASIC
PROTOCOL 4
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and Nuclei
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Supplement 23
Table 22.4.13 Coplin Jars Needed for Post-Hybridization Washes and
Immunofluorescent Detection of Indirectly Labeled DNA Probesa
No. Coplin jars
Contents
Temperature
3
50% formamide in 2× SSC
45◦ C
3
1× SSC
45◦ C
9
0.1% Tween 20 in 4× SSC
45◦ C
a See Basic Protocol 4.
Materials
Hybridized slides in a hybridization box (see Basic Protocol 3 and Alternate
Protocol 6)
50% formamide in 2× SSC, 45◦ C (prepare fresh)
1× SCC, 45◦ C (UNIT 18.6)
Blocking solution: 1% BSA (w/v)/0.1% (v/v) Tween 20 in 4× SSC (store
indefinitely at −20◦ C or up to several months at 4◦ C)
Primary, secondary, and tertiary antibodies (see recipe for antibodies)
0.1% (v/v) Tween 20 in 4× SSC, 45◦ C
DAPI in antifade (see recipe)
Clear nail polish
Coplin jars
22 × 50–mm glass coverslips
37◦ C oven
Fluorescent microscope and appropriate filter sets (UNIT 4.2)
CAUTION: Formamide is a carcinogen and should be handled with care. Discard according to biohazard rules of the institution.
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.13.
Remove coverslip
1. After 24 hr, remove hybridized slide from the hybridization box.
2. Carefully peel the rubber cement from the hybridized slide and immerse in a Coplin
jar containing 50% formamide in 2× SSC, 45◦ C.
3. Allow the coverslip to fall off and let stand 5 min in solution with gentle agitation.
Wash and block
4. Remove slide and transfer to second Coplin jar containing formamide/SSC, 45◦ C,
for 5 min. Agitate gently. Repeat with a third Coplin jar containing formamide/SSC,
45◦ C.
5. Wash the slide for 5 min each in three consecutive Coplin jars containing 1× SSC,
45◦ C, agitating gently between washes.
6. Drain excess solution but do not allow the slides to dry. Add 80 µl blocking solution,
coverslip with 22 × 50–mm glass, and place back in hybridization box. Incubate
40 min at 37◦ C.
Fluorescence
In Situ
Hybridization
(FISH)
Label with primary antibody
7. Remove coverslip and add 80 µl primary antibody. Coverslip and place back into the
hybridization box. Incubate 40 min at 37◦ C.
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Table 22.4.14 Common Primary and Secondary Antibody Systems and Final Concentrationsa
Hapten
Primary antibody
Secondary antibody
Tertiary antibody
Color
Probe Type
Biotin (no signal
amplification)
5 µg/ml FITC-avidin —
—
green
Centromere
chromosome paints
Biotin
5 µg/ml FITC-avidin 5 µg/ml biotinylated 5 µg/ml FITC-avidin Green
goat anti-avidin
Locus-specific
cDNAs
Biotin
5 µg/ml
rhodamine-avidin
5 µg/ml biotinylated 5 µg/ml
goat anti-avidin
rhodamine-avidin
Red
Locus-specific
cDNAs
Digoxigenin (no
2 µg/ml rhodamine
signal amplification) anti-Dig
—
—
Red
Centromere
chromosome paints
Digoxigenin (no
2 µg/ml FITC
signal amplification) anti-Dig
—
—
Green
Centromere
chromosome paints
Digoxigenin
0.5 µg/ml mouse
anti-digoxigenin
2 µg/ml digoxigenin 2 µg/ml rhodamine
anti-mouse
anti-Dig
Red
Locus-specific
cDNAs
Digoxigenin
0.5 µg/ml mouse
anti-digoxigenin
2 µg/ml digoxigenin 2 µg/ml FITC
anti-mouse
anti-Dig
Green
Locus-specific
cDNAs
a Shown in this table are the most common antibody systems for the detection of biotinylated and digoxigenin (Dig)-labeled DNA. Final concentrations
for antibodies are also stated but may require adjustment. A variety of antibody systems are available from Molecular Probes and Roche Diagnostics. If a
two–color FISH approach is being used, be sure that respective primary, secondary, and tertiary antibodies do not cross-react. This may require sequential
hapten detection rather than concomitant hapten detection. The scheme above uses antibodies raised against mouse and goat, thus no cross-reaction will
occur. Antibodies raised in rabbits will also offer more variety in hapten detection.
This antibody may or may not have a fluorescent moiety conjugated to it, depending on the
nature of the probe (see Critical Parameters). Common primary antibody and secondary
antibody systems are outlined in Table 22.4.14.
8. Remove coverslip and wash slide for 5 min each in three consecutive Coplin jars containing 0.1% Tween 20/4× SSC, 45◦ C, with gentle agitation. If proceeding with signal
amplification (see Table 22.4.15) continue to step 9. Otherwise, proceed to step 14.
9. Drain excess wash solution, but do not allow slide to dry. Add 80 µl blocking solution,
coverslip, and place back into the hybridization box. Incubate 10 min at 37◦ C.
Label with secondary antibody
10. Remove coverslip and add 80 µl secondary antibody (Table 22.4.14). Coverslip and
place back into the hybridization box. Incubate 30 min at 37◦ C.
11. Remove coverslip and wash slide 5 min each in three consecutive Coplin jars containing 0.1% Tween 20 in 4× SSC, 45◦ C, with gentle agitation.
Label with tertiary antibody
12. Remove coverslip and add 80 µl tertiary antibody (conjugated to fluorochrome).
Coverslip and place back into the hybridization box. Incubate 30 min at 37◦ C.
13. Remove coverslip and wash slide 5 min each in three consecutive Coplin jars containing 0.1% Tween 20 in 4× SSC, 45◦ C, with gentle agitation.
Counterstain and seal
14. Drain excess wash solution, but do not allow the slides to dry. Add 40 µl DAPI in
antifade counterstain. Coverslip and seal with clear nail polish.
The slides are now ready for visualization. When not in use, store slides up to several
weeks or months at −20◦ C, depending on the frequency of visualization.
15. Examine using a fluorescent microscope and appropriate filter sets.
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and Nuclei
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Table 22.4.15 Interpretation of Parallel Positive Control Experiments
Problem
Slide hybridized
with new probe
Slide hybridized
with good known Analysis
probe
Background
Present
Not present
New probe may still contain unincorporated dNTPs
or many small labeled fragments are present and
did not hybridize.
Slide quality is not the problem.
Washing conditions also do not appear to be the
cause.
Background
Present
Present
Problems likely related to washing conditions or
slide quality rather than issues with the probe.
Weak Signal
(indirect-labeled probe)
Yes
No
If using antibody detection system for indirectly
labeled probes, this scenario indicates that the
antibodies are in good working order. Look to
problems with incorporation of the hapten into the
DNA.
Weak signal
(direct-labeled probe)
Yes
No
This indicates a problem with the labeling of the
probe. Check that all enzymes and fluorochromes
are within their shelf life.
This also indicates that washing conditions are
correct.
Weak signal (indirect
labeled probe)
Yes
Yes
This indicates a problem with the antibody
detection system. Ensure that the concentrations are
correct and that the antibodies are fresh. It may also
indicate a problem with hybridization, insufficient
denaturation of the target DNA on the slide, or
overall quality of the DNA specimen.
Weak signal
(direct-labeled probe)
Yes
Yes
The problem may be in the post-hybridization
washes: too stringent or temperatures too high.
Poor chromosome
morphology with weak
signal
Yes
Yes
Over denaturation if the chromosomes appear puffy.
Good chromosome
morphology but no signal
Yes
Yes
Slides are likely too old and resistant to
denaturation. Change to a more fresh preparation.
Otherwise the experiment can be repeated but the
denaturation of the slide should be increased.
ALTERNATE
PROTOCOL 8
Post-Hybridization Washes and Detection of Directly Labeled DNA Probes
The protocol below outlines steps for washing slides hybridized with directly labeled
probes. If using commercially produced probes, follow the manufacturer’s instructions.
This protocol is applicable to hybridized cytogenetic slides or paraffin sections. See Basic
Protocol 4 for materials.
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.16.
Fluorescence
In Situ
Hybridization
(FISH)
1. After 24 hr hybridization, carefully peel off rubber cement from slide and immerse
in a Coplin jar containing 50% formamide in 2× SSC, 45◦ C. Allow the coverslip to
fall off and let stand 5 min.
2. Remove slide and transfer to a fresh Coplin jar containing formamide/SSC, 45◦ C.
Incubate 5 min. Repeat with a third Coplin jar containing formamide/SSC, 45◦ C.
22.4.22
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Table 22.4.16 Coplin Jars Needed for Post-Hybridization Washes and Detection of Directly Labeled DNA Probesa
No. Coplin jars
Contents
Temperature
3
50% formamide in 2× SSC
45◦ C
3
1× SSC
45◦ C
3
0.1% Tween in 4× SSC
45◦ C
a See Alternate Protocol 8.
Table 22.4.17 Coplin Jars Needed for Rapid Wash of Directly Labeled Probesa
No. Coplin jars
Contents
Temperature
1
0.3% (v/v) NP-40 in 2× SSC
72◦ C
1
0.3% (v/v) NP-40 in 2× SSC
Room temperature
1
2× SSC
Room temperature
a See Alternate Protocol 9.
3. Wash slide 5 min each in three consecutive Coplin jars containing 1× SSC, 45◦ C.
4. Wash slide 5 min each in three consecutive Coplin jars containing 0.1% Tween 20 in
4× SSC, 45◦ C, with gentle agitation.
5. Drain excess wash solution, but do not allow the slide to dry. Add 40 µl DAPI in
antifade counterstain. Coverslip with 22 × 50–mm glass and seal with clear nail
polish. Store slide up to several months at −20◦ C.
Fluorescence fading will depend upon the frequency of viewing.
6. Examine using a fluorescent microscope equipped with the appropriate filters.
Rapid Wash of Directly Labeled Probes
This protocol describes the use of high temperatures and stringent SSC washes for the
removal of unbound probe from target DNA. This protocol is generally effective for directly labeled DNA, particularly from commercial sources. It is important to consider
the differing stringency requirements of each probe being used when planning this protocol since excessive temperature can strip bound probe from the target. This protocol is
applicable to cytogenetic slides or paraffin sections.
ALTERNATE
PROTOCOL 9
Additional Materials (also see Basic Protocol 4)
2× SSC (UNIT 18.6)
0.3% (v/v) NP-40 in 2× SSC, room temperature and 72◦ C
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.17.
1. Carefully remove rubber cement from the hybridized slide. Place the slide and coverslip in a Coplin jar containing 2× SSC to gently remove coverslip.
2. Once the coverslip has fallen off, incubate the slide 1 to 2 min in a Coplin jar containing
0.3% NP-40 in 2× SSC, 72◦ C.
3. Remove slide and wash 1 min in a Coplin jar containing 0.3% NP-40/2× SSC, room
temperature.
4. Drain solution from slide and add DAPI in antifade counterstain. Add a 22 × 50–mm
glass coverslip and seal with clear nail polish.
5. Examine the slide with a fluorescent microscope equipped with the appropriate filters.
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ALTERNATE
PROTOCOL 10
Post-Hybridization Washes for Specimens Hybridized with PNA Probes
This protocol describes a procedure for removing unbound PNA probe from cytogenetic
specimens or from paraffin sections.
Materials
Slides hybridized with PNA probes (see Alternate Protocol 5 or 7)
70% (v/v) formamide/10 mM Tris·Cl (pH 7.0 to 7.5)/0.1% (w/v) BSA (see recipe)
0.1 M Tris·Cl (pH 7.0 to 7.5)/0.15 M NaCl/0.08% (v/v) Tween 20 (store up to
several weeks at room temperature)
70%, 90%, and 100% ethanol
DAPI in antifade solution (see recipe)
Fluorescent microscope and appropriate filters
NOTE: A list of all Coplin jars used in this protocol is given in Table 22.4.18.
1. Remove coverslip from slide hybridized with PNA probes and wash 15 min in a
Coplin jar containing 70% formamide/10 mM Tris·Cl (pH 7.0 to 7.5)/0.1% BSA.
Repeat once.
2. Wash 5 min each in three consecutive Coplin jars containing 0.1 M Tris·Cl (pH 7.0
to 7.5)/0.15 M NaCl/0.08% Tween 20.
3. Dehydrate slide by incubating 5 min each in Coplin jars containing 70%, 90%, and
100% ethanol. Air dry (5 to 10 min).
4. Apply DAPI in antifade and coverslip.
5. Examine the slide using a fluorescent microscope and appropriate filters.
6. Store slides up to several months at −20◦ C
INTERPRETATION OF FISH FINDINGS
This section discusses interpretation of FISH experiments and relevant troubleshooting
measures. Each laboratory may adopt a different means of assessing positive and negative
results as is applicable to the experiment and the question being asked. The guidelines
below can generally be applied to both interphase and metaphase analysis.
Evaluating FISH
FISH encompasses four main components: the slide and its preparation (see Target Slide
Preparation), the DNA probe and its preparation (see Labeling DNA Probes for FISH),
Table 22.4.18 Coplin Jars Needed for Post-Hybridization Washes for Specimens Hybridized with PNA Probesa
No. Coplin jars
Fluorescence
In Situ
Hybridization
(FISH)
Contents
Temperature
2
70% formamide/10 mM Tris·Cl
(pH 7.0–7.5)/0.1% BSA
Room temperature
3
0.1 M Tris·Cl (pH 7.0–7.5)/0.15 M
NaCl/0.08% Tween 20
Room temperature
1
70% ethanol
Room temperature
1
90% ethanol
Room temperature
1
100% ethanol
Room temperature
a See Alternate Protocol 10.
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denaturation and hybridization of the probe to the slide (see Hybridization), and finally,
post-hybridization washes and detection (see Post-Hybridization Washes and Detection).
Each of these components is, in itself, multistepped. The discussion below outlines many
of the problems encountered in FISH experiments, including factors influencing signal
strength, background, and preserving optimal chromosome morphology. Other sources
for FISH optimization and parameters can be found in Beatty et al. (2002), Henegariu
et al. (2001), Schwarzacher and Heslop-Harrison (2000), and van de Rijke et al. (1996).
Signal strength
Slide age. In the author’s experience, optimal results have been obtained from slides not
older than 2 months. As the slides age further (3 to 6 months), they become harder to
denature. Conversely, very old preparations (>1 year) often have partially degraded DNA
that may reduce or preclude effective FISH experiments. In such situations, preparations
tend to become very sensitive to denaturation. Previously G-banded slides (UNIT 22.3) have
an even shorter life span and should be processed within 2 weeks (refer to Table 22.4.5).
Cytoplasmic debris. The presence of cytoplasm on the slide may inhibit binding and contribute to overall background. A more aggressive protease pretreatment may be required
to reduce cytoplasmic protreinacious noise. This can be accomplished by increasing the
time of digestion or the amount of protease added (maintain the same digestion time).
One must also consider whether the slide is made from a dropped suspension or FISHed
to a slide made from an in situ culture, since the latter slides tend to possess more cytoplasmic and cellular debris. A high background obscures true signals. If there is minimal
background but the signal is weak in different parts of the slide, then there may be a
gradient or patchiness of cellular protein in areas of the slide with a higher density of
fixed cells.
Excessive slide pretreatment. Excessive enzymatic treatment may damage the target DNA,
making it less efficient for hybridization with the probe. This is usually indicated by weak
uptake of the counterstain and the presence of bright centromeres.
Denaturation time. As the slide ages (see above), the chromosomes become harder to
separate into single strands. Slides used within 1 to 2 weeks of preparation should be denatured for 1.5 to 2 min. Slides that are older may require times that range from 2 to 3 min.
Under-denaturaton results in insufficient strand separation of the target DNA, decreasing
the effective hybridization efficiency. Over-denaturation of the target DNA causes DNA
damage and reduces the amount of target DNA that is able to hybridize with the probe;
it also results in reduced hybridization efficiency. Over-denaturation usually also results
in poor uptake and banding using the DNA counterstain and very bright centromeres.
Sealing of coverslip. It is critical that the coverslip be adequately sealed with rubber
cement during incubations to prevent any moisture from entering the hybridized area,
thus diluting the probe. Do not use contact cement or other ultra-adhesive glues.
Proper temperature for washes and incubations. Check that the temperature in the oven,
hot plate unit, or water bath is correct for hybridization and other incubations. High temperatures during post-hybridization washes may remove bound probe with weak sequence
homology (see Critical Parameters), thus decreasing the amount of DNA available for
signal detection or antibody binding. Avoid taking shortcuts during incubation times with
blocking reagents or antibody detection reagents. Also be sure that the reagents do not
dry up on the slide; this makes washing more difficult.
Detergent type and concentration. Increasing the detergent content or choice of detergent
may influence signal strength. Tween 20 is generally less harsh than NP-40. Increasing
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the detergent concentration in combination with increasing temperature will remove
more probe and/or bound antibodies. Be sure that the concentration and temperatures
are correct.
SSC concentration. Decreasing SSC concentration increases stringency. Thus 0.1× SSC
destabilizes double-stranded DNA more readily than 2× SSC. Double-stranded DNA
stability is also affected by the extent of sequence homology between the probe and
target DNAs. Ensure that the proper SSC concentrations are used during washes.
Proper filter sets. For fluorescence, make sure that the proper filter sets are used for image
acquisition. The use of filters with incorrect spectral characteristics for the fluorochrome
being used can severely impair the ability to detect the correct signals or they may increase
the amount of autofluorescence. Also refer to UNIT 4.2 and APPENDIX 1E.
Amplification of signal. When using an indirectly labeled DNA probe, signal amplification
may be required. If the signal is amplified but still weak, check the concentrations and shelf
lives of antibodies used. If the antibodies are in good order, relabel the probe and check
the labeling efficiency (see Critical Parameters). For directly labeled probes, consider the
labeling efficiency of the probe.
Probe characteristics
In-house probes. If probes are labeled in-house (see Labeling DNA Probes for FISH),
strict controls must be undertaken to ensure that proper hapten or fluorochrome incorporation has been obtained to produce a high-quality DNA probe. Haptens may be assessed
using the dot blot method of incorporation. Directly labeled probes can be assessed by
removing a small aliquot of probe, placing it on a slide, and viewing it by fluorescence
microscopy.
Cot-1 suppression. Excessive Cot-1 suppression may be the cause of reduced signal.
Smaller DNA probes or cDNAs contain fewer repeat elements compared to larger inserts.
Normally, 5 to 10 µg Cot-1 is sufficient per 100 ng labeled DNA; adjust as required.
Commercial probes. Usually commercial suppliers properly process the product with the
necessary quality controls. Check that the probes were properly stored and used before
the expiration date. For some experiments it is possible to optimize procedures using
concentrations of commercial probes at levels slightly below those recommended by the
supplier. This can prove very cost effective if a specific commercial probe is going to be
used routinely.
Probe concentration. The probe is usually used in excess of the target DNA; however,
make sure that sufficient probe volume has been added to adequately cover the area of
interest on the slide (see Table 22.4.11).
Preannealing. Preannealing at 37◦ C is usually carried out for 1 hr following denaturation.
This step is performed to permit the annealing of repetitive sequences in the DNA probe
with unlabeled Cot-1 DNA to prevent cross-hybridization. If there are few repetitive
elements in the probe, or the probe is a cDNA or small insert, this step may be omitted.
The probe is simply denatured and applied to the denatured slide.
Background
Fluorescence
In Situ
Hybridization
(FISH)
Cytoplasmic debris. This is the most common cause of background. Increase the incubation time during protein digestion or maintain the same time but change the concentration
of the enzyme.
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Bacterial/yeast contamination. Microorganisms that contaminate cultures used to make
the slide, or reagents used to prepare the slide or for hybridization are deposited onto the
slide. If the contamination is minor, then it is only necessary to analyze the slide in areas
that contain fewer deposits of fungi or bacteria. If the contamination is heavy, the slide
or reagent must be prepared again.
Coverslips. Coverslips should be clean and dust free. If cells are to be grown in situ, use
only glass coverslips as plastic coverslips autofluoresce.
Residual oils. Slides that have been previously visualized using immersion oil (e.g., Gbanded slides) should be cleaned with xylene. Residual oils will prevent hybridization
and cause background problems.
Dust and other particles. Particles may be deposited on the slide during various transfers
into solutions. Be sure that solutions are well mixed and filtered as needed. Some solutions
may form precipitates that may bind to the slide.
In-house probes. Ensure that nonincorporated conjugated nucleotides are removed from
the final probe preparation.
Commercial probes. These probes are usually not the cause of background, especially
when the same probe on another slide preparation has not produced background; however,
note the lot number for future reference. It is also very helpful to introduce a new batch
of a commercial probe into routine use before the previous lot is exhausted in case the
supplier has had production difficulties
Unlabeled DNA: carrier and Cot-1. An excess of unlabeled DNA is added to the probe
mixture. Although it is unlabeled, it can contribute to background by trapping antibodies.
To reduce this problem, ensure that the resuspended probe pellet is fully dissolved in
sufficient hybridization buffer. If the pellet does not completely dissolve, add more buffer.
This will not significantly alter the reaction.
Denaturation time. The probe is usually heat denatured a minimum of 5 min. This
should be sufficient time to denature the probe and dissolve any remaining DNA.
The denaturation time may be increased to 10 min without any damage to the DNA
probe.
Post-hybridization washes. Make sure that the correct temperature has been maintained
for washes and incubations. Agitation during the washes can help to remove unbound
probe and antibodies. Increasing the stringency of the washes by either increasing the
temperature or altering the amount of SSC in the washes can also help considerably
if background is encountered. Avoid drying the slide with any residual blocking or
detection reagents.
Detergent concentration. Increasing the detergent concentration in post-hybridization
washes in combination with increasing temperature removes more probe and/or bound
antibodies. Be sure that the concentration and temperatures are correct.
Fading signals
In-house probes. Ensure that stocks of directly conjugated dNTPs are properly stored
in the dark and used well before the expiration date. Unlike indirectly labeled probes,
directly labeled DNAs have a much shorter shelf life.
Commercial probes. Although these probes are supposed to be quality controlled, these
too are subject to the same concerns as in-house probes (see above). During denaturation
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and hybridization, keep lights dimmed and hybridize in a light-proof container for directly
labeled probes.
Directly-labeled DNA probes. When directly labeled probes are used, slides should be
washed with the lights dimmed to prevent unnecessary exposure to extraneous light.
Mounting medium. Check that the DAPI/antifade mounting solution is used before its
expiration date. Normally, the medium is clear with a slight pink tinge; medium that
is degrading will turn increasingly amber. Expired antifade medium causes rapid signal
degradation and displays a red glow when viewed under the microscope.
Image acquisition. Extended exposure of the hybridized slide to UV sources leads to
signal quenching. When not in use, store slides at −20◦ C. During visualization, keep the
light source off when not in use.
Poor chromosome morphology
Slide age. See Signal Strength (above) for information concerning this parameter.
Cytoplasmic debris. See Signal Strength (above) for information concerning this
parameter.
Excessive slide pretreatment. Excessive enzymatic treatment may damage the target DNA,
making it less efficient for hybridization with the probe
Previously G-banded slides. G-banding (UNIT 22.3) causes DNA to become sensitive to
subsequent denaturation. For this reason, denaturation times are greatly reduced. This
also has an effect on the quality of hybridization and signal strength.
Over-denaturation. The cause of poor chromosome morphology is usually overdenaturation, which causes the DNA to be destroyed. This can be sample specific and/or
slide-age related.
Denaturation temperature. Check that the temperature for denaturation is accurate. The
final internal temperature of the Coplin jar should be 72◦ C. Add 1◦ C for each slide that is
denatured in the jar. Avoid denaturing more than five slides in one jar at one time. Note
that plastic Coplin jars have a greater differential temperature between bath and internal
temperatures as compared to glass.
Probe cross-hybridization
Clone identity. When using in-house FISH probes for research applications, ensure that
the clone identity and the insert are correct for the experiment. Some genes have highly
homologous sequences elsewhere in the genome and may cross-hybridize to other family
members.
DNA contamination. Consider whether stock DNA was contaminated with DNA from
another clone. Increase the hybridization temperature if contamination is suspected. Typically, the hybridization temperature is 37◦ C. If cross-hybridization occurs, try increasing
the hybridization temperature to 42◦ C.
Fluorescence
In Situ
Hybridization
(FISH)
Stringency of washes. Some probes may cross-hybridize, particularly centromere probes.
Usually cross-hybridization signals are significantly weaker than true signals. More stringent washes can be undertaken by decreasing the SSC concentration, increasing wash
temperature, or adding additional washes.
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Evaluation of a New DNA Probe
The most critical factor in optimizing a new FISH probe which was labeled in-house
is first determining whether the probe of interest maps to the correct location. Many
vectors (e.g., plasmids, cosmids, BACs) have traveled from laboratory to laboratory
and have changed hands many times. These DNAs, as well as cloned DNA obtained
from reputable repositories, can be mislabeled or misidentified. After a cloned DNA has
been extracted and labeled, the first FISH experiment should be carried out on normal
lymphocyte spreads and should address the following questions.
Is a signal visible on a pair of chromosomes?
If there is a weak signal, refer to the section above.
Does the DNA probe map to the right chromosomal location? Does it map to only
one location?
It helps to have someone who can identify the chromosomes by DAPI banding. Most
people are not as proficient as trained cytogeneticists, thus it may be useful to carry
out a two-color experiment using a commercially available centromere probe for the
chromosome of interest. Refer to UNIT 22.3 for general conventions for identification of
chromosomes according to ISCN nomenclature. Some DNA sequences/genes may belong
to families with similar sequences. It is possible that hybridization to these similar regions
may occur. Usually the true signals are stronger than the cross-hybridized ones. If this is
the case, wash conditions can be adjusted or another clone should be obtained.
Does a positive control probe work properly?
It is useful to perform parallel experiments with a probe known to give good signals in
the same experiment. This will help to identify any problems that are not related to the
newly labeled probe (see Table 22.4.15 for interpretation of such control experiments).
For example, in panel iii of Figure 22.4.4A, two mouse BAC probes were labeled (one
green and the other red) and assessed. A separate slide with a previously labeled BAC
known to give good signal and no background was prepared as a control at the same
time (not shown). The newly labeled BAC probes shown in panel iii of Figure 22.4.4A
mapped to the proper location with weak signal intensity and high background. Analysis
of this experiment is as follows.
The BAC probes map to the correct location. Because the signal intensity was weak
consider (1) starting DNA quality and quantity, (2) labeling efficiency, (3) fidelity of DNA
polymerase I and DNase I, (4) labeling time and temperature, (5) effective removal of
nonincorporated nucleotides, and (6) insufficient Cot-1 suppression. Because the control
probe displayed no background problems, one can assume that the hybridization and
wash conditions were sufficient.
Evaluation of Hybridization Efficiency
The minimum number of cells or metaphase spreads required to obtain a given result
reflects the clinical context of the finding, the limitations of the patient material available
for study, and the question being asked. With tissues or cells that are hard to obtain, a
single abnormal metaphase may be significant. For example, in some situations, limited
FISH data may be supported by results obtained using PCR and/or Southern blot analysis.
Prior to enumerating or analyzing FISH results on a test sample, it is important to carefully assess the overall quality, uniformity, and effectiveness of hybridization as discussed
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Figure 22.4.4
Legend at right.
Fluorescence
In Situ
Hybridization
(FISH)
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above. Each hybridized slide should be evaluated for the specificity of the hybridization,
the probe signal intensity, and the signal-to-background noise, to determine if the hybridization was optimum for the given analyses. There should be minimal background or
nuclear fluorescent noise. At least 85% of all nuclei in the target area should be easily innumerable. For some applications (such as the detection of mosaicism or minimal residual
disease), more rigorous analytical sensitivities and hybridization efficiencies are required.
1. Perform the FISH protocol appropriate for the type of patient and control slides in the
experiment.
2. Prior to determining the hybridization efficiency, quickly scan the whole slide, noting
the general signal-to-noise levels in different regions of the slide and any areas with
high background or unusually weak signals. It may be useful to mark the underside
surface of the slide in these areas with a diamond pen. If the signal intensity far exceeds
background levels, proceed with estimating the hybridization efficiency.
3. Pick several representative areas of the slide and score at least 200 nuclei from the
areas selected, following the selection criteria described below in Selecting Cells for
FISH Microscopy. Keep a running log of the number of cells scored and the observed
signal counts for the patient and control slides.
4. For all slides, add up the number of cells with no signal. In general, hybridization
is considered to be adequate if >85% of the cells scored have one or more signals.
Lower hybridization efficiencies may be encountered with smaller probes (generated
Figure 22.4.4 (at Left) Composite figure illustrating FISH hybridization. (A) Assessment of a
DNA probe. (i) Cross-hybridization of probe (arrowhead) to chromosome 1. True signals are located on chromosome 22. (ii) Changes in stringency washes and post-hybridization removes
cross-hybridization. (iii) High background associated with a insufficient repetitive sequence (Cot1) suppression in this mouse metaphase spread. Weak signals (arrows) indicate poor labeling
efficiency. (B) Interphase FISH analysis. (i) A cytogenetic specimen from a short-term ovarian
primary culture hybridized with PNA probes for centromeres 7 (green) and 8 (red). When scoring
interphase nuclei, it is especially important to focus through the cells since signals may be present
at different z planes. (ii) Schematic indicating the nuclei that are acceptable for scoring: A, cell acceptable for scoring; B, cell is likely acceptable for scoring, but requires careful attention; ?, cell has
qualities that make it questionable for scoring; X, cell is not acceptable for scoring. (iii) Interphase
analysis using PNA probes specific for centromeres 7(green) and 8 (red) on a paraffin section
from a prostate carcinoma. Arrowheads indicate cells containing changes in ploidy. (C) Analysis
of translocation probes. (i) and (ii) Results of consecutive hybridization of a G-banded metaphase
spread with the Vysis BCR/ABL translocation probe. Two fusion signals (yellow) are produced from
the hybridization of BCR/ABL fusion on the Philadelphia chromosomes (Ph) on chromosome 22
and the reciprocal ABL/BCR on the derivative chromosome 9. Separate green and red signals
from the normal chromosomes 9 and 22 are also seen. (iii) Interphase pattern from this specimen.
(D) Example of a chromosomal inversion on chromosome 11. An inversion was identified involving
the terminal portion of chromosome 11 by gross cytogenetic analysis. Clones 200 kb apart and in
the 11p15.5 region, containing the IGF2 gene (green) and H19 (red) were hybridized to the patient
specimen. The normal chromosome 11 shows the red and green signal hybridizing on top of each
other at 11p15.5. The inverted 11 shows the clear spit of signal along the abnormal chromosome
11 indicating the breakpoint lies within the 200 kb between IGF2 and H19. (E) Gene amplification
of MYCN in neuroblastoma specimens. (i) Double minute chromosomes (dmns) containing hundreds of copies of the MYCN gene. This is in contrast to amplification of MYCN in (ii) as a large
block of signal called a homogeneously staining region (HSR). Interphase nuclei nearby show the
typical hybridization pattern for an HSR. (F) Use of subtelomeric and pan-centromeric PNA probes.
(i) Hybridization of a prostate cell line with PNA subtelomeric probes. (ii) Hybridization of another
prostate cell line with subtelomeric and pan-centromeric PNA probes. Loss of telomeric sequences
are indicated by the arrow while the presence of multicentric chromosomes are indicated by arrow
heads. This black and white facsimile of the figure is intended only as a placeholder; for full-color
version of figure, see color plates.
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for example in a research laboratory). Extreme caution must be exercised when using
probes with lower hybridization efficiencies or elevated background signal to provide
clinical information.
Selecting Cells for FISH Microscopy
Generally, look at all areas of the slide and analyze regions with uniformity in signal
strength. Compare the intensity of the background signals to the intensity of the signals
in the nuclei or metaphases of interest. The FISH signal intensity should consistently be
greater than the background intensity in the regions of the slide chosen for analysis. If
the background signals are equivalent to signals in the nuclei, then the counts will be
skewed and the results biased. The following provides a guide to selecting targets for
analysis.
1. Ideally analyze cells from all areas of the slide. Systematically select representative areas from different regions of the slide. Any regions that have unacceptable background
or weak signals identified in prescreening evaluations should be excluded.
2. Select nuclei or metaphase cells that do not touch or overlap (Fig. 22.4.4B).
3. Select intact nuclei that have smooth well-rounded borders. Partially ruptured nuclear
membranes may have lost informative chromatin. Similarly select metaphase spreads
that have no evidence of preparation artifacts or breakage.
4. Select cells that are not surrounded by cytoplasmic material and that are without
evidence of potential drying artifacts such as rings or clumped cells.
5. Do not evaluate interphase nuclei with signals located on the extreme periphery of the
nucleus.
6. Do not score regions of the slide containing nuclei that have no signals. Absence of
signals may represent uneven or patchy hybridization, resulting in some areas of the
slide having very weak or absent signals.
Analysis of Interphase Nuclei
Interpretation of interphase FISH is very much dependent on statistical analyses and has
inherent technical challenges. For instance, the presence of signal is dependent upon the
probe and its fluorescent label successfully entering the cell and hybridizing to the target
DNA. Detection of the correct number of signals can be complicated by signals overlapping or splitting. Any background hybridization whatsoever leads to major complications
in interpretation. It is not uncommon to find monosomy or trisomy in nuclei that reflect
technical artifact or false positive background signals. Therefore, the accuracy of interphase FISH analysis is dependent upon recognizing these technical issues, correcting for
them, and standardizing the scoring criteria accordingly. Interphase analysis is typically
used for enumeration of chromosomes using centromere probes, detection of gene amplification or deletion (see below), and detection of the presence of translocations, so that
many cells should be scored. Metaphase FISH analysis, although more informative, is
more difficult, especially when there are few metaphase spreads present.
1. Select nuclei in which signals generally have the same intensity.
2. Focus up and down in the z axis to accommodate spatial configurations of probe signals
within the nucleus (Fig. 22.4.4B).
Fluorescence
In Situ
Hybridization
(FISH)
3. Signals that are more intense in some nuclei than the specific signal indicate the
presence of regional background. Care must be taken when analyzing any sample
with this type of background noise.
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Figure 22.4.5 Schematic representation of scoring criteria. Shown in the illustration are typical hybridization signal
configurations. These images pertain to a single probe, such as a centromere- or locus-specific probe. Signals must be
more than one signal width apart to be considered one signal. Signals joined by a string of hybridization are also considered
as one signal.
4. Do not count lower level nonspecific hybridization signals. These signals can usually
be recognized by their lower intensity and different shape.
5. Some nuclei will have passed through the S phase of the cell cycle and may be present
as G2-paired signals (i.e., two smaller signals in very close proximity). These paired
signals represent a single chromosome already divided into chromatids and should be
counted as one signal (Fig. 22.4.5).
6. Count two signals connected by a strand of fluorescence as one signal. Sometimes centromere or long genomic probes will generate signals that are not spherical. Typically,
FISH signals appear as separate fluorescent dots on each chromatid of a metaphase
chromosome when the target size is 100 to 250 kb. Similarly, in interphase nuclei,
such probes will also generate discrete easy-to-interpret signals. Larger probes can
appear as fused signals straddling both chromatids, and in interphase nuclei, these
probes can generate signals that present more diffuse or dispersed hybridization spots
in the chromatin of interphase nuclei. Knowledge of the probe size and anticipated
configuration in both metaphase spreads, and interphase nuclei is essential. As long
as the signal is continuous it should be scored as one signal.
7. Count only nuclei in which a definite enumeration can be made. Do not analyze or
enumerate inconclusive cells.
8. Use two people to score 200 consecutive nuclei from each sample such that each
person scoring will analyze ∼100 nuclei from a given sample. The slides should be
coded and scored independently by two analysts. Any discrepancies may mean the
established scoring criteria for the FISH assay are not being adhered to rigidly.
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Analytical sensitivity of interphase FISH assays
Once a particular probe set is made available as a routine FISH test for a clinical service, it is important that the laboratory perform assay validation and create a database
establishing reportable range and general laboratory experience with each probe. The
analytical sensitivity assay measures the success of a given FISH test in a particular
laboratory environment and on a given tissue type. Since there are known differences
in the cell populations and tissue types, it is important to use the appropriate positive
control tissue for the assessment of analytical sensitivity and for the establishment of the
database. Analytical sensitivity analyses are performed by scoring 200 interphase nuclei
representing at least five normal, preferably male, individuals. (Pooling samples on one
slide is acceptable.) The nuclei are scored for the percentage of nuclei exhibiting the
appropriate number of distinct signals.
For constitutional studies using FISH, the recommended analytical sensitivity for probes
intended for nonmosaic detection is 90%, while for probes intended to detect mosaicism, it is 95%. Similarly, for detection of minimal residual disease, sensitivities
≥95% are helpful. Databases for each probe should be established so that it will
then possible to determine the mean and standard deviation of results from a series of normal samples processed and analyzed in the same manner as clinical samples. False positive rates can then be calculated and used for final scoring reports.
More discussion on this issue is available from the following sources: VYSIS guidelines for single (http://www.vysis.com/tech sup fishproto quality single.asp) and dual
probes (http://www.vysis.com/tech sup fishproto quality dual.asp), and scoring criteria
for preimplantation genetic diagnosis of numerical abnormalities for chromosomes X, Y,
13, 16, 18, and 21 (Munne et al., 1998).
Statistical considerations concerning interphase FISH analysis of paraffin sections
Due to truncation of the nuclei during sectioning, loss of signal from areas of the nucleus
excluded from the target slide will be encountered when enumerating signals after FISH
has been performed on paraffin sections. The criteria for determining the significance
of loss or gain of signals in interphase nuclei will depend on a number of parameters
(e.g., nuclear diameter, age of patient, type of tissue). Readers are referred to some of the
scientific literature where suggested cutoff values are adopted from the available literature
(Qian et al., 1996). In the authors’ experience with FISH analysis of prostate cancer,
chromosomal gains can been identified when more than ∼10% of the nuclei exhibit more
than two signals. Panel iii of Figure 22.4.4B, shows an example of a prostate section
hybridized with centromere probes for centromeres 7 (green) and 8 (red). Chromosomal
losses have been identified when more than 50% of the nuclei exhibit a reduction of
signal number, and tetraploidy has been assumed when all chromosomes investigated
show signal gains up to four. For some classes of tumor, extreme polyploidy together
with complex patterns of chromosomal rearrangement means that it is not realistic to
select a suitable control chromosomal region in which two signals are expected. In such
situations, it may be helpful to perform flow cytometric analysis of DNA content in
parallel with interphase FISH analysis.
Fluorescence
In Situ
Hybridization
(FISH)
Analysis of Translocation and Inversion Probes
If the probes based on green and red fluorescence used for FISH are close to specific
translocation breakpoints on different chromosomes, they will appear joined as a result
of the translocation, generating a yellow color fusion signal. Commercial probes are
now available for many of the common translocations in cancers (Table 22.4.19). One
such probe from Vysis detects the Philadelphia chromosome (Ph) resulting from the
22.4.34
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Table 22.4.19 Commonly Used Commercial Probes for Detection of Translocations in Sarcomas
and Hematological Malignancies
Neoplasm
Chromosomal
location
Probe
Scoring method
CML/pediatric ALL
9q34/22q
BCR/ABL
Color fusion observed in
metaphase and interphase
Various leukemias
11q23
MLL
Split signal, metaphase
Various leukemias
5q31
EGFR1
Loss of signal,
metaphase/interphase
Various leukemias
7q31
DSS486
Loss of signal,
metaphase/interphase
AML M4 EO
Inv(16)
CBFB
Split signal, metaphase
Various leukemias
20q13.2
ZNF217, D20S183
Loss of signal,
metaphase/interphase
Various hematologic
malignancies
8q24 /14q32
MYCC/IgH
Color fusion observed in
metaphase and interphase
AML-M1
12p13/21q22;
8q22/21q22
TEL/AML1;
AML1/ETO
Color fusion observed in
metaphase and interphase
AML-M3
15q22/17q21.1
PML/RARA
Color fusion observed in
metaphase and interphase
Ewings sarcoma
t(11;22) (q24;q12)
FLI1/EWS
Color fusion observed in
metaphase and interphase
Rhabdomyosarcoma
t(2;13)(q35;q14)
PAX/FKR
Color fusion observed in
metaphase and interphase
translocation between ABL on chromosome 9 and BCR on chromosome 22. Shown in
Figure 22.4.4C is an example of combined G-banding and FISH analysis using the Vysis
BCR/ABL probe set. Panel i of Figure 22.4.4C shows the G-banded metaphase spread to
which the BCR/ABL probe was subsequently hybridized. In panel iii of Figure 22.4.4C,
detection of a Ph chromosome in interphase nuclei of leukemia cells is achieved by the
presence of two double-fusion (D-FISH) signals. All nuclei positive for the translocation
contain one red signal (BCR gene), one green signal (ABL gene), and two intermediate
fusion yellow signals because the 9;22 chromosome translocation generates two fusions,
one on the 9q+ and a second on the 22q−. The following general guidelines may be
helpful for performing this type of assay.
1. Green and red signals that are juxtaposed but not overlapping should be scored as
ambiguous.
2. Do not score nuclei that are missing a green or red signal. This assay is looking for
the presence or absence of a fusion signal, not the absence of a green or red signal.
3. Atypical signal patterns have been reported and are now considered to be clinically
important (Kolomietz et al., 2001).
Table 22.4.19 lists some of the commonly used FISH assays in hematological cancers
as well as sarcomas. In addition to the scientific literature, readers are referred to the
suppliers web sites (see Internet Resources and SUPPLIERS APPENDIX), which will provide
the most up-to-date listing of currently available probes and the preferred scoring method.
Inversions are related to translocations such that a break and rejoining occurs within the
resident chromosome. Probes are available to detect common inversions present in AMLs
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(see Table 22.4.19). Mapping of breakpoints is common in many research cytogenetic
laboratories and involves chromosome walking. To identify the locus containing the
breakpoint, rough cytogenetic analysis locates the band region. Clones are then obtained
spanning the putative breakpoint. These clones are then FISHed to the specimen of
interest. Such is the case in Figure 22.4.4D. An inversion was identified involving the
11p15.5 region. Clones 200 kb apart, containing the IGF2 gene (green) and H19 (red)
were hybridized to the patient specimen. The normal chromosome 11 shows the red and
green signal hybridizing on top of each other at 11p15.5. The inverted 11 shows the clear
split of signal along the abnormal chromosome 11 indicating the breakpoint lies within
the 200-kb distance between 1GF2 and H19.
Use of FISH Probes in Assessing Solid Tumors and Gene Amplification
Gene amplification is one of the mechanisms by which cancer cells achieve over expression of some classes of oncogenes, which involves an increase in the relative number of
copies of a gene per cell. This can range from one or two additional copies per cell to
extreme examples where over a thousand copies per cell have been reported. Gene amplification can occur in association with the over-expression of oncogenes, thus conferring
a selective growth advantage or mechanism of acquired resistance to chemotherapeutic
agents leading to poor prognosis. Gene amplification is highly suited to FISH analytical
approaches that have the added benefit of excellent sensitivity and the ability to address
cellular heterogeneity.
Neuroblastoma is characterized by the frequent occurrence of a highly amplified oncogene, MYCN. It has been known for many years that the presence of this aberration is
strongly associated with poor outcome. More aggressive management is usually required
when MYCN is found to be amplified. Similarly, breast cancer can be accompanied by
an amplified oncogene HER2/Neu, and presence or absence of this aberration may determine which of different treatment regimens are followed. Examples of metaphase and
interphase FISH assays for gene amplification are shown in Figure 22.4.4E. In this figure, a DNA probe containing the MYCN gene was hybridized to a cytogenetic specimen
from a neuroblastoma patient. Panel i of Figure 22.4.4E illustrates double minute (dmns)
chromosomes containing hundreds of copies of the MYCN gene as extrachromosomal
bodies. This is in contrast to another patient where amplification of the MYCN gene occurs as a homogeneously staining region (HSRs) inserted in a chromosome other than the
resident site of MYCN (normally at 2p24). Interphase nuclei in this image show a large
patch of hybridization signal characteristic of HSRs. Some of the commonly detected
aberrations observed in solid tumors which are amenable to FISH analysis are presented
in Table 22.4.20.
Fluorescence
In Situ
Hybridization
(FISH)
Analysis of Telomere Probes
Telomeres are located at the ends of chromosomes and are characterized as (T2 AG3 )
repeat sequences and their associated proteins (Poon et al., 1999). Maintained by the
ribonucleoprotein complex, telomerase, they function to protect chromosomes from endto-end fusions. In most normal tissues, telomerase is expressed at very low levels. As such,
each round of DNA replication results in the gradual shortening of the telomeres. The
shortening of telomeres is associated with replicative cell senescence. The up-regulation
of telomerase extends the proliferative lifespan of a normal cell. In abnormal cells, expression of telomerase is associated with the maintenance of telomere length or telomere
lengthening. Conventional Southern blot analysis gives average telomere length but fails
to yield information on individual chromosome ends. It also underestimates the size
and number of short telomeres. Although DNA probes for these sequences are available
commercially, the use of commercial PNA probes for such sequences, as well as for centromeric and pancentromeric sequences, has enabled researchers to determine the overall
22.4.36
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Table 22.4.20 FISH Probes Used For Assessing Solid Tumors and Gene Amplification
Neoplasm
Chromosomal location
Probe
Scoring method
Neuroblastoma 2p24; 2p23-24
MYCN
Interphase, metaphase;
amplification
Ewings
sarcoma
t(11;22) (q24;q12)
FLI1/EWS
Color fusion observed
in metaphase and
interphase
Breast cancer
17q11.2-q12
HER2/neu
Interphase
amplification
Glioblastoma
7p10-p21
EGF-R
Interphase
amplification
Bladder cancer Centromere regions of
Centromere 7, 13, 9 Interphase enumeration
chromosomes 3, 7,17 and
9p21 region of chromosome 9
telomere lengths of individual cells (interphase nuclei) and chromosomes (metaphases)
using quantitative digital imaging, as described by Poon (1999), with greater specificity
and accuracy. Both fixed cytogenetic cells (Poon et al., 1999) and paraffin-embedded samples (Vukovic et al., 2003) have been used for telomere analysis. Panel i of Figure 22.4.4F
illustrates subtelomeric PNA probes hybridized to a prostate cell line metaphase spread.
Digital imaging and signal intensity ratios determine the relative telomere length. In some
cases, loss of telomere signals can also be identified as in panel ii of Figure 22.4.4F (arrow). In this figure, a prostate cell line was hybridized with subtelomeric PNA probe as
well as a PNA pan-centromeric probe. The pancentromeric probe confirmed the presence
of multicentric chromosomal structures indicative of chromosomal instability.
For those who wish to engage in telomere studies, access to digital imaging and analysis software capable of determining telomere length is suggested. Like all other FISH
experiments, the background should be minimal. Telomere signals are relatively small
and located at the ends of chromosomes. Background, such as antibody speckling can
greatly affect the sensitivity of the analysis. The proper controls must also be established.
Since telomere length is dependent on the number of cell divisions, age- and sex-matched
controls should be included in all experiments.
REAGENTS AND SOLUTIONS
Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX
2A; for suppliers, see SUPPLIERS APPENDIX.
Agarose gel, 2% (w/v)
Dissolve 2 g molecular biology–grade agarose in 1× TBE buffer (APPENDIX 2A) by
either microwaving or placing on a hot plate, and allow to cool slightly. Add 5 µl of
10 mg/ml ethidium bromide in a well vented chemical hood. Pour into casting trays
(APPENDIX 3A) and allow to solidify. Store up to 1 week at 4◦ C covered with foil.
AP-labeled antibody mixture
Dilute anti-biotin or anti-digoxigenin conjugated to alkaline phosphatase (AP; Invitrogen) to a final concentration of 0.75 U/ml in 100 mM Tris·Cl (pH 7.5)/15 mM
NaCl (APPENDIX 2A). Prepare fresh for each experiment.
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Antibodies
Prepare antibodies stocks according to the manufacturer’s instructions. Tertiary
antibodies are conjugated to fluorchromes. Store up to several months as frozen
aliquots. Prepare working solutions of primary and secondary antibodies according
to the manufacturer’s instructions using the guidelines for final antibody listed in
Table 22.4.14. Diluted antibodies should be kept at 4◦ C in light-proof containers
for 2 to 3 months.
Minor adjustments in concentration will need to be made to account for background
and signal intensity.
DAPI, 100 µg/ml
Dilute all of the powder from a 1-mg bottle of 4 ,6-diamidino-2-phenylindole (DAPI;
Sigma) in 10 ml buffer to produce a 100 µg/ml stock solution. Store in small aliquots
up to 1 year at −20◦ C.
CAUTION: DAPI is a potential carcinogen and should be handed with care.
DAPI in antifade
Combine the following in order:
5.0 ml 1× PBS (APPENDIX 2A)
500.0 µl 100 µg/ml DAPI stock (see recipe; 1 µg/ml final)
0.5 g p-phenylenediaminie (Sigma; 10 mg/ml final; dissolve well)
45.0 ml glycerol (90% v/v final)
Transfer to a 50.0-ml conical tube, wrap with aluminum foil (product is light sensitive), and place on a rotator 30 min to ensure proper mixing. Store in 1-ml aliquots
up to 1 year at −20◦ C.
The resulting solution is very viscous
This reagent is available commercially as Vectashield (Vector Laboratories).
DNase I, 3 mg/ml
3.0 mg DNase I powder
500.0 µl glycerol (50% v/v final)
50.0 µl 1 M Tris·Cl, pH 7.5 (50 mM final; APPENDIX 2A)
5.0 µl 1 M MgCl2 (5 mM final)
1.0 µl 1 M 2-mercaptoethanol (1 mM final)
1.0 µl 10 mg/ml BSA (10 µg/ml final)
Adjust volume to 1.0 ml with H2 O
Store in 50-µl aliquots up to 1 year at −20◦ C.
Fluorescence
In Situ
Hybridization
(FISH)
DNase I dilution buffer
250.0 µl 1 M Tris·Cl, pH 7.0 (50 mM final; APPENDIX 2A)
25.0 µl 1 M MgCl2 (5 mM final)
5.0 µl 1 M 2-mercaptoethanol (1 mM final)
20.0 µl 10 mg/ml BSA (4 µg/ml final)
4.7 ml H2 O
Store in 1-ml aliquots up to 6 months at −20◦ C or
In 1 ml aliquots up to 1 month at 4◦ C.
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dNTP mixture, 1 and 2 mM
For a 1 mM mixture:
1.0 µl 100 mM dATP
1.0 µl 100 mM dCTP
1.0 µl 100 mM dGTP
97.0 µl H2 O, sterile
Store in 100-µl aliquots up to several months at −20◦ C
For a 2 mM mixture: Double the volume of each 100 mM dNTP added (i.e., use
2 µl of 100 mM dNTP) and reduce the amount of water to 94 µl. Store up to several
months.
The total volume of either solution is 100 µl.
Fluorochrome/dTTP mixture
The amount of fluor-dUTP will vary. Thus, the recipes below are guidelines. Refer to the manufacturer’s suggested final concentrations. All labeled nucleotides
should be stored in 20 to 30-µl aliquots up to 6 months at −20◦ C in light-proof
containers.
FITC-dUTP/dTTP:
3.5 µl 1 mM dTTP (0.6 mM final)
1.75µl 1 mM fluorescein (FITC)-dUTP (0.3 mM final; Roche)
The total volume, 5.25 µl, is appropriate for one labeling reaction and can be scaled
up as needed.
Rhodamine-dUPT/dTTP mixture:
3.5 µl 1 mM dTTP (0.6 mM final)
1.75 µl 1 mM rhodamine-dUTP (0.3 mM final; Roche)
The total volume, 5.25 µl, is appropriate for one labeling reaction and can be scaled
up as needed.
Texas Red-dUPT/dTTP mixture:
3.5 µl 1 mM dTTP (0.7 mM final)
1.0 µl 1 mM Texas Red-dUTP (0.2 mM final; Roche)
The total volume, 4.5 µl, is appropriate for one labeling reaction and can be scaled
up as needed.
Cy3-dUPT/dTTP mixture:
3.5 µl 1 mM dTTP (0.7 mM final)
1.25 µl 1 mM Cy3-dUTP (0.2 mM final; Amersham Biosciences)
The total volume, 4.75 µl, is appropriate for one labeling reaction and can be scaled
up as needed.
Cy5-dUPT/dTTP mixture:
3.5 µl 1 mM dTTP (0.7 mM final)
1.25 µl 1 mM Cy5-dUTP (0.2 mM final; Amersham Biosciences)
The total volume, 4.75 µl, is appropriate for one labeling reaction and can be scaled
up as needed.
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Formamide, 70%/Tris·Cl (pH 7.0 to 7.5), 10 mM/BSA, 0.1% (w/v)
Dissolve 100 mg BSA in 29 ml water. To this solution, add 70 ml formamide (Invitrogen) and 1 ml of 1 M Tris·Cl, pH 7.0 to 7.5 (APPENDIX 2A). Store up to several weeks.
CAUTION: Formamide is a carcinogen and should be handled with care. Discard
according to institution’s biohazard rules.
Total volume is 100.0 ml; however, the solution can be made in larger quantities
and stored up to several weeks at 4◦ C.
Hapten/dTTP mixture
The amount of hapten-dUPT will vary. Thus, the recipes below are guidelines. Refer
to the manufacturer’s suggested final concentrations. Store up to several months or
a year at −20◦ C.
Biotin-dTTP/dUTP mixture:
3.5 µl 1 mM dTTP (0.6 mM final)
1.75 µl 1 mM biotin-16dUTP (0.3 mM final; Invitrogen)
The total volume, 25 µl, is appropriate for one labeling reaction and can be scaled
up as needed.
Digoxigenin-dTTP/dUTP mixture:
3.5 µl 1 mM dTTP (0.6 mM final)
1.75µl 1 mM dig-11dUTP (0.3 mM final; Roche)
The total volume, 5.25 µl, is appropriate for one labeling reaction and can be scaled
up as needed.
Hybridization buffer
500.0 µl high grade formamide (50% v/v final; Invitrogen)
100.0 µl 20× SSC (2× final; UNIT 18.6)
100.0 µl dextran sulfate (10% final)
300.0 µl H2 O
Store in 100-µl aliquots up to several months at 4◦ C
Total volume is 1.0 ml.
Hybridization buffer can also be purchased from DAKO.
Loading dye, 5×
0.125 g bromphenol blue (0.25% final)
15.0 ml glycerol (30% final)
35.0 ml H2 O
Store in 1.0-ml aliquots up to several months at 4◦ C
Total volume is 50.0 ml.
Fluorescence
In Situ
Hybridization
(FISH)
NBT/BCIP
22.5 µl 75 mg/ml NBT (Invitrogen)
17.5µl 50 mg/ml BCIP (Invitrogen)
4.96 ml 100 mM Tris·Cl (pH 9.5; APPENDIX 2A)/100 mM NaCl/50 mM MgCl2
Prepare fresh for each experiment.
Volumes can be scaled up or down as required.
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Nick translation buffer, 10×
1.0 µl 10 mg/ml BSA (0.1 µg/µl final)
10.0 µl 1 M 2-mercaptoethanol (0.1 M final)
50.0 µl 1 M Tris·Cl (0.5 M final)
5.0 µl 1 M MgCl2 (50 mM final)
34.0 µl H2 O
Store in 100-µl aliquots up to several months at 4◦ C
Total volume is 100.0 µl.
PBS, 1×/MgCl2 , 0.05 M
950.0 ml 1× PBS
50.0 ml 1 M MgCl2
Store at room temperature until ready for use.
The total volume is 1 liter.
Pepsin, 0.5% (w/v) in NaCl, 0.85%
Dissolve 500 mg pepsin in 100.0 ml of 0.85% (w/v) sodium chloride. Adjust pH to
1.5 with 12 N HCl. Prepare fresh for each experiment.
PI, 100 µg/ml
Dissolve the powder from an entire 10-mg bottle of propidium iodide (PI; Sigma) in
10 ml water to a final concentration of 100 µg/ml. Store in small aliquots indefinitely
at −20◦ C.
PI in antifade
Combine in the following order:
5.0 ml 1(PBS; APPENDIX 2A)
150.0 µl 100 µg/ml PI (0.3 µg/ml final; see recipe)
0.5 g p-phenylenediaminie (10 mg/ml final; Sigma; dissolve well)
45.0 ml glycerol (90% final)
Transfer to a 50.0-ml conical tube, wrap in aluminum foil (the product is light
sensitive), and place on a rotator 30 min to ensure proper mixing. Store in 1-ml
aliquots up to several months at −20◦ C.
The resulting solution (50 ml total) is very viscous.
This solution is available commercially available from Vectashield (Vector Laboratories).
RNase I, 100 µg/ml
Prepare a final 100 µg/ml solution of RNase I in 2× SSC (UNIT 18.6) fresh for each
experiment using any molecular-grade RNase enzyme.
Sonicated salmon sperm DNA standards
Prepare 50.0 ng/µl salmon sperm DNA standard by combining 5.0 µl of a 10 µg/µl
stock (Invitrogen), 300 µl of 5× loading dye (e.g., UNIT 18.6), and 695 µl water.
Prepare a 25.0 ng/µl standard by combining 500 µl of the 50 ng/µl standard, 100 µl
of 5× loading dye, and 400 µl water. Prepare a 12 ng/µl standard by combining 500
µl of the 25.0 ng/µl standard, 100 µl of 5× loading dye, and 400 µl water. Store all
standards up to several months at 4◦ C or indefinitely at −20◦ C.
The average size of the commercial salmon sperm is ∼500 to 2.0 kb
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Triethanolamine (pH 8.0), 0.1 M
660.0 µl triethanolamine (final 0.1 M; Sigma)
50.0 ml H2 O
Prepare fresh for each experiment.
Total volume is 50.0 ml.
COMMENTARY
Background Information
Fluorescence
In Situ
Hybridization
(FISH)
Applications of FISH
As early as 1988 (Lichter et al., 1988), FISH
was used to visualize labeled DNA probes hybridized to chromosome and interphase nuclei preparations. The improvement of cloned
DNA sources, antibodies, fluorochromes, microscopy and imaging equipment, and software has permitted a variety of scientific investigations. FISH analysis is routinely used
in clinical cytogenetic laboratories for detection of syndromes in prenatal assessments,
including Prader-Willi/Angelman Syndrome,
DiGeorge Syndrome, and Cri-du-chat. Cancer cytogenetics laboratories use FISH to confirm and monitor hematological malignancies such as CML and AML (Table 22.4.19
and Table 22.4.20). Gene amplification studies
for neuroblastoma (MYCN amplification), and
breast cancer (HER2/NEU amplification) are
also routinely carried out. Mapping of newly
identified genes have relied heavily on FISH
(Squire et al., 1993; Lichter et al., 1988) both
in humans and other species (Boyle et al.,
1990, Giguere et al., 1995). Metaphase spreads
may not always be easy to obtain, thus interphase cells offer a means of obtaining information, albeit via an indirect method. This
has been useful for gauging chromosome instability (Speicher et al., 1995; Ghadimi et al.,
1999; Al-Romaih et al., 2003; Vukovic et al.,
2003) and determining normal versus abnormal content, as well as gene amplification or
deletion. FISH applied to paraffin-embedded
sections appeared in the early 1990s (Thompson et al., 1994) and was applied to both thick
(>5 µm) and thin (5 µm) paraffin sections. Application of FISH to paraffin-embedded sections provides a tremendous opportunity to
correlate the histopathological classification to
the genomic changes detected. Furthermore, it
enables the investigator to study concepts of
cellular heterogeneity, tumor focality, and
metastasis (Squire et al., 1996). Paraffin FISH
analysis requires patient and careful technical expertise. Proper controls must be used
when interpreting final results. Depending on
the type of information sought, paraffin FISH
is an acceptable form of data collection when
fresh tissues for cytogenetic suspensions are
not available; however, paraffin FISH will not
provide chromosome-based information.
Preparation of probes
The development of reliable cloning strategies in the 1980s facilitated the genomic
analysis and sequencing of specific DNA fragments. Mapping of these genes to their chromosomal locations was previously laborious
and infrequently reliable. The emergence of
FISH in the early 1990s paved the way for an
effective and direct means of mapping specific DNA fragments to their chromosomal
locations.
Types of probes. The creation of comprehensive genomic libraries as a result of the
human genome project provides renewable
resources for FISH probes. At present,
bacterial artificial chromosomes (BACs) are
the most popular cloned forms of genomic
DNA used for FISH probes. Although yields
are generally considered low in comparison
to plasmids and cosmids (predecessors of
BACs), BAC inserts are larger (200 kb) and
can produce a stronger FISH signal compared
to cosmids and plasmids, which are considerably smaller in insert size (i.e., ranging from
2 to 30 kb). P1 artificial chromosomes (PACs)
and yeast artificial chromosomes (YACs) have
also been used in the past and are still used
in many research laboratories. Other probes,
such as RNA and oligonucleotide probes are
options and are reviewed in Speel (1999), and
Schwarzacher and Heslop-Harrison (2000).
More recently, peptide nucleic acid (PNA)
probes (Lansdorp, 1996), have been used
for FISH experiments with great success
(Martens et al., 1998; Vukovic et al., 2003.).
Using the same nucleotide bases as DNA
probes, PNA probes do not have a phosphate
or deoxyribose sugar backbone. Specific
fluorochromes, haptens, or enzymes can be
chemically attached to the bases and used in
the same fashion as DNA probes. PNA probes
are sequence specific. Thus, sequence information must accompany a PNA custom probe
22.4.42
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Current Protocols in Cell Biology
order to companies such as Applied Biosystems
(http://www.appliedbiosystems.com).
There are several advantages of PNA probes.
For instance, PNA has a neutral backbone,
which provides stronger binding and greater
specificity of interaction. Also, the uncharged
PNA structure creates stronger binding independent of salt concentration, allowing more
robust hybridization applications. Finally, the
fact that PNA oligomers have resistance to
nucleases and proteases, permitting more robust in situ experiments, is also an advantage.
The drawback of PNA probes are in the cost
of their fabrication. The typical molecular
(cytogenetic) laboratory cannot generate
these probes in-house. The cost-effectiveness
of PNA probes, therefore, can only be
appreciated if the laboratory is carrying out a
particular hybridization frequently, as in the
case of a clinical cytogenetic laboratory.
Methods for preparing probes. By far the
most commonly used strategy for labeling
DNA probes is by nick translation. Nick translation involves the simultaneous actions of two
enzymes: DNase I and DNA polymerase I.
DNase I randomly nicks the DNA fragment
in each strand of the double-stranded DNA
molecule. DNA polymerase I (derived from
E. coli), with its three activities—i.e., exonuclease function (removes bases in the 5 to
3 direction), polymerase function (adds nucleotides from the 3 nick site), and the 3 to
5 proof reading function—incorporates the label, whether indirectly using a hapten (i.e., biotin or digoxigenin) or directly (i.e., fluorescein or rhodamine). Nick translation can be
applied to all cloned DNA sources without the
need to remove vector sequences. Polymerase
chain reaction (PCR) is another commonly
used method of labeling DNA probes from
cloned sources, as well as from microdissected
genomic DNA or flow-sorted chromosomes.
PCR employs multiple rounds of template
denaturation, primer annealing, and template
replication, facilitated by Taq DNA polymerase. Primers are variable and may include
sequence-specific primers to amplify those targeted fragments, vector sequence primers to
amplify and label cloned DNA, and/or universal primers. Compared to nick translation,
which requires micrograms of starting DNA,
PCR requires only nanograms of template.
Furthermore, nick translation is most efficient
when the starting DNA is of high molecular
weight with little or no degraded material. PCR
labeling is a little more forgiving of DNA quality; thus, suboptimal DNA may still be used
when not acceptable for nick translation. De-
pending on the template, several rounds of
amplification may be required before labeling, as in the case of in-house production of
chromosome painting probes. Other labeling
strategies, including random priming and endlabeling, are also options and are described in
Schwarzacher and Heslop-Harrison (2000).
For both nick translation and PCR labeling,
commercial kits and enzyme mixes are readily
available and optimized. Usually, commercial
kits combine DNase I and DNA polymerase I
as an enzyme mix. The drawback of combining them is the fact that one enzyme (usually
the DNA polymerase I) will lose its enzymatic
activity before the other. Commercial kits can
also be costly. Making an in-house kit allows
the investigator to alter enzyme concentrations
as necessary to better optimize the reaction.
This alleviates the combined-enzyme problem
with commercial kits. In-house kits are also
expensive and they do require rigorous internal quality control and troubleshooting from
time to time.
See Critical Parameters for discussion
of slide preparation, hybridization, posthybridization washes and detection, and interpretation of FISH findings.
Critical Parameters
Preparation of probes
DNA quality. Ensure that the starting DNA
to be labeled has a high molecular weight
with no degraded DNA fragments or protein.
Contaminating RNA will contribute slightly
to the spectrophotometric quantification of the
DNA and cause a miscalculation of the actual starting amount of DNA. DNA should be
assessed using an accurate spectrophotometer
and by gel electrophoresis. Compare the concentration of DNA as determined using a spectrophotometer and gel with a DNA of known
concentration. The DNA used as a standard for
concentration can be another undigested clone
or genomic DNA.
The starting DNA should be dissolved in
water rather than Tris/EDTA (TE) buffer. TE
is normally used to inhibit any DNases that
may be present in the water. By that reasoning,
TE buffer has been known to inhibit the enzymatic activity of the DNase during the nick
translation reaction. Furthermore, it may inhibit the polymerase activity for nick translation and PCR labeling.
Reagents. Ensure that reagents are used
prior to their expiration dates. The most critical
reagents in the labeling reactions are:
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1. DNA polymerase I/Taq DNA polymerase. Failure of polymerase activity in the
reaction will lead to inefficient incorporation
of the labeled nucleotide into the DNA, resulting in a very weak FISH signal.
2. Conjugated nucleotides. Nucleotides
conjugated to a fluorochrome are light sensitive. Repeated and prolonged exposure to
direct light will quench the fluorescent signal. Store these reagents in small aliquots at
−20◦ C in light-proof containers. Restrict exposure to light during the labeling procedure
and store labeled DNA at −20◦ C in lightproof containers. Unlike nonfluorescently labeled probes, these directly-labeled DNAs will
have a shorter shelf life. The concentration of
nucleotides can vary from as much as 1 mM to
as low as 0.2 mM in different nick translation
protocols. The labeled nucleotide should always be in at least a 2- fold excess of the corresponding unlabeled nucleotide. Furthermore,
the labeled nucleotide can be interchangeable
between hapten-dUTP or hapten-dATP. If the
investigator chooses to use Biotin 14-dATP
rather than the dUPT form, the nucleotide mixture should be adjusted accordingly.
3. 2-Mercaptoethanol. Buffers containing
2-mercaptoethanol should be replaced every
3 to 4 months, particularly if not stored at
−20◦ C.
4. DNase I. Although the activity of DNase
I is usually potent enough that it may be used
for up to a year when stored at −20◦ C, the specific activity may decrease over time. This is
evident when the fragment sizes are larger than
expected when the standard DNase I concentration and incubation times are used. Simply
increase the amount of DNase I used in the reaction and keep the same period of incubation,
or keep the same amount of DNase I used in the
reaction and increase the time of incubation.
5. PCR factors. Treat PCR labeling like
any other PCR experiment and consider the
following factors: amount of template, pH,
magnesium ion concentration, and primer
sequence.
Equipment. Small volumes are used for
both labeling strategies. Pipets should be calibrated and serviced often. Use an accurate
spectrophotometer and water bath that can
effectively maintain a constant temperature,
if a thermocycler is not available for nick
translation.
Fluorescence
In Situ
Hybridization
(FISH)
Preparation of slides
FISH involves the hybridization of the labeled and denatured DNA probe to its denatured DNA target located on the slide. Each
component requires care to obtain the most effective results. Pretreatment of the slide with
a protease before denaturation permits the removal of any cellular and cytoplasmic debris. In the case of paraffin-embedded sections,
more aggressive treatment is required to remove residual wax as well as to allow sufficient protease treatment to permit access of
the probe to the intended target DNA.
Protease treatment. Treatment with pepsin
has been featured in this protocol because its
action is relatively mild as compared to other
proteases such as proteinase K; however, many
laboratories use proteinase K alone or in combination with pepsin, particularly for the digestion of paraffin-embedded sections, to obtain the proper digestion conditions for FISH
(see Table 22.4.9). The incubation times of any
protease treatment will vary according to the
type of tissue (e.g., paraffin-embedded), quality of the slide (i.e., quantity of cytoplasmic debris/cellular debris), as well as the lot or batch
of pepsin used. It is suggested that incubation
times be increased rather than the concentration of pepsin used in the digest. The temperature at which the incubations occur is also
critical. Higher temperatures facilitate faster
digestions as opposed to those at room temperature or below. While variations in the pretreatment protocol can be made, the starting
quality of the fixed tissue is the most critical factor. Ideally, the tissue should be fixed
shortly after removal from the host.
Paraffin-embedded samples. In the case of
paraffin-embedded tissues, formalin is the fixative of choice for routine use in most clinical
laboratories; however, superior results for most
research needs may be obtained if tissue sections are fixed using ethanol or paraformaldehyde prior to embedding. Buffered formalin
solutions have also been implemented to provide better recovery of DNA, RNA, and protein
for future studies. The time spent in fixation is
equally as important as the type of fixative. The
longer the time spent in fixative, the more difficult it may be to denature the DNA to a singlestranded state. This is due to the cross-linking
action of the fixative to proteins that enables the
cells to maintain their morphology. Sections
for FISH should be cut fresh from the original block, preferably a few sections from the
exposed face. This will help to ensure that the
DNA has not been exposed to DNases, or oxidative damage from the handling of the block
or from the local environment. Several sections
should be taken so that different pretreatment
conditions can be tested. It is suggested that
the investigator take the time to contact the
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histology laboratory preparing the specimens
to document the fixation procedure for troubleshooting later.
Denaturation. Denaturation is also a critical
parameter for successful FISH. Too much denaturation of the target DNA causes excessive
DNA damage, yielding poor banding and/or
hybridization. Too little denaturation results in
ineffective hybridization due to limited access
to the target DNA. The extent of denaturation
will depend on several factors.
1. Quality of the slide. Namely, the presence
of cytoplasmic and/or cellular debris.
2. Age of the slide. Whether using a fresh
slide, a fresh slide with artificial aging, a
slide that is 1 week or 1 month old, and so
forth.
3. Previous assays. Namely if banding has
previously been performed on the slide.
4. Tissue type. Whether fibrous, bony, or
muscle tissue, or the like, is being used.
5. Age and method of fixation. See above.
6. Section thickness. Normally 5-µm-thick
sections are used.
Hybridization
Optimal hybridization is influenced by
many factors that will affect the target DNA
and the probe itself. These factors are sources
for inefficiencies and should be carefully monitored and documented such that effective troubleshooting can be carried out.
Stringency. The hybridization of a DNA
probe to its DNA target is dependent on the
extent of sequence homology, the complexity
of the DNA sequence, and the chemical factors
that influence the success of maximal binding
efficiency. Stringency refers to the percentage
of matches and mismatches between the probe
and target such that the hybridized double helix
remains stable. FISH experiments are typically
run at 70% to 90% stringency, with 70% representing a lower relative stringency than 90%.
This is critical when determining the feasibility of carrying out cross-species FISH experiments (i.e., human probes on mouse targets
or vice versa), where sequence homology may
be considerably lower in parts of the probe.
The reagents present in hybridization buffers,
as well as the temperature significantly influence stringency.
Temperature. DNA strand melting and annealing are influenced by temperature. Since
reagents such as formamide are present in the
hybridization buffer, high temperatures (i.e.,
80◦ to 90◦ C) are not required for DNA denaturation. This is the reason denaturation of
the DNA can be accomplished at 72◦ C in
the presence of formamide. Increasing temperatures also increases stringency, such that
a 1◦ C alteration increases the stringency by
1%. Thus, increasing the hybridization temperature from 37◦ to 42◦ C will increase the
stringency of the hybridization step, requiring the probe to possess higher sequence
homology with its target. This may be particularly useful if there is a chance of crosshybridization, which is typically the case with
locus-specific probes of genes that belong to
a class of genes/sequences that are very similar. Pseudogenes may also cross-hybridize.
Lowering the hybridization temperature under
37◦ C will result in lower stringency conditions
permitting the hybridization of the probe to
targets with greater mismatch. This also has
advantages and disadvantages. At low stringency, mismatched spurious hybridization
events will be more common and contribute
to more experimental noise at the detection
step.
Probe size. The rate of hybridization is also
influenced by probe size. Generally, longer
fragments (>500 bp) will require a longer time
to hybridize than shorter ones. Once again, the
hybridization is affected by the extent of sequence homology.
PNA probes. In the case of PNA probes,
the specificity of the probe, as a required function of their fabrication through oligosynthesis, eliminates some of the complexities experienced by DNA probes. Since the PNA
probe has an uncharged backbone, there is no
charge repulsion when the probe hybridizes to
the DNA, thus creating a stronger bond. Furthermore, PNA/DNA bonds will melt 15◦ C
above DNA/DNA bonds, almost independently of salt concentration. The specificity
of the PNA/DNA bond lies in the increased
destabilization of the PNA/DNA bond when a
mismatch occurs.
Post-hybridization washes and detection
Stringency. The factors that affect hybridization also influence the conditions involved in post-hybridization washes and antibody incubations. Incubation at 37◦ C under
a range of 50% to 60% formamide in 2× SSC
sets a stringency of ∼70% to 80%. This permits hybridization of sequences with a 20%
to 30% mismatch and maintains duplex stabilization. Increasing hybridization temperatures raises the stringency, requiring greater sequence homology for effective hybridization.
Conversely, decreasing temperatures permits
hybridization with greater mismatch. Decreasing salt ion strength (i.e., SSC) also increases
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Table 22.4.21 Stringency of DNA/DNA Hybridization and Post-Hybridization Washesa
Stringency as a function of % formamide
SSC
concentration
Temperatureb
60
55
50
45
40
35
30
5×
37◦ C
76
73
70
67
64
61
58
2×
◦
37 C
83
80
77
74
71
68
65
1×
◦
37 C
88
85
82
79
76
73
70
0.75×
37◦ C
90
87
84
81
78
75
72
0.5×
37◦ C
93
90
87
84
81
78
75
0.2×
37◦ C
100
96
93
90
87
84
81
0.1×
37◦ C
105
101
98
95
92
89
86
5×
42◦ C
81
78
75
72
69
66
63
2×
42◦ C
88
85
82
79
76
73
70
1×
◦
42 C
93
90
87
84
81
78
75
◦
42 C
95
92
89
86
83
80
77
0.5×
◦
42 C
98
95
92
89
86
83
80
0.2×
◦
42 C
105
101
98
95
92
89
86
0.1×
◦
110
106
103
100
97
94
91
0.75×
42 C
a This Table describes the changes in stringency for probes with 43% GC content at 300 bp (modified from Schwarzacher
and Heslop-Harrison, 2000). Increasing the formamide content or temperature, or decreasing the SSC strength will
increase stringency. A combination of these three factors can be used to modulate the stringency.
b Hybridization of probe to DNA typically occurs under 50% formamide in 2× SSC at 37◦ C, giving a 77% stringency.
Washing of unbound probe following hybridization typically occurs under 50% formamide in 2× SSC at 42◦ C, giving
an 82% stringency.
Fluorescence
In Situ
Hybridization
(FISH)
stringency. The combination of temperature
and salt concentration can affect hybridization
kinetics considerably. The same holds true for
removal of unbound probe.
Typically, post-hybridization washes are
carried out at 42◦ to 45◦ C. This temperature, coupled with 50% formamide and 2×
SSC, permits 90% stringency for probes with
a 43% GC content, at a probe size of 300 bp
(Schwarzacher and Heslop-Harrison, 2000).
Table 22.4.21 describes the stringency conditions based on these parameters.
Detergents. Some suppliers of commercial probes may suggest higher temperature
washes of 72◦ C in a low ion (high stringency)
SSC wash containing a detergent. This too is
based on high sequence homology and the relative GC sequence content of the probe. This
rapid wash method eliminates the need for formamide. Caution should be exercised when using high temperatures, since there is the risk
of partial denaturation of the hybridized probe
and target. This will result in loss of signal.
A rapid wash should only be used with probes
that possess high sequence homology and high
labeling efficiency. Commercial probes tend to
fit these criteria.
The subsequent SSC washes and washes
with detergents serve to remove any remaining unbound probe and antibodies. As before, decreasing the SSC content will create a
more stringent wash. The choice of detergent
is subjective, largely based on the preference
and experience of the investigator. The most
commonly used detergents include Tween 20,
NP-40, IPEGAL, SDS, and Triton X-100 (reviewed in Speel, 1999). In general, the actions
of these detergents are comparable when used
at the standard 0.1% in 4× SSC; however,
SDS, Triton X-100, and NP-40 are generally
harsher than Tween 20. If high background persists, the number of washes with the detergent
can be increased or the percentage of detergent
and SSC strength can be altered.
Antibodies. The choice of antibody systems is also fairly standard. Many companies
provide antibodies raised in different animals
and conjugated to different fluorochromes so
that a variety of probes may be used and detected at one time. Some antibodies may require a higher concentration; thus, it is important that the signal strength be monitored
carefully for background and the antibody concentration adjusted as required. As discussed
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above, caution should be exercised when using various antibody combinations for detecting more than one probe: ensure that there is
no cross-reactivity between antibodies raised
in different species.
Counterstains. Finally the use of a counterstain helps to visualize the cells or spreads.
DAPI stains DNA blue upon fluorescence and
is most commonly used. DAPI, however, may
obscure a green signal if using a dual filter
(i.e., FITC/DAPI), so a red counterstain, such
as propidium iodide (PI), may be more appropriate. If the color of probe detected is red,
do not use PI as a counterstain. The signals
are protected from rapid quenching through
the reagents used in the mounting medium.
These counterstains in mounting (antifade)
medium can be purchased commercially. If the
counterstain is prepared in the laboratory, be
sure to use high quality glycerol, since some
lower quality glycerols can cause autofluorescence. The concentration of counterstain may
require adjustment. Higher counterstain concentrations may obscure FISH signals. If this
is the case, dilute the counterstain in antifade
by adding more glycerol and PBS in amounts
that will be consistent with the final concentrations.
Storage. All slides should be stored at
−20◦ C for long-term storage (i.e., up to 1 year)
and at 4◦ C for short-term storage (i.e., up to
6 months).
Troubleshooting
Preparation of probes
See Table 22.4.22 for a troubleshooting
guide to labeling.
Preparation of slides
Cytogenetic specimens. The influence of
pretreatment and denaturation cannot be readily assessed until the entire FISH protocol has
been carried out and the slides are visualized
by fluorescence microscopy (Henegariu et al.,
2001). The slides, however, may be visualized
by phase-contrast microscopy for cytogenetic
preparations after the dehydration step following pepsin treatment to determine the extent
of digestion. The slides may be returned for
digestion if the cytoplasmic debris does not
appear to have been removed. Similarly, for
treatment of paraffin sections, following visualization for autofluorescence, the slides may
be returned for digestion and reassessed.
If there are no cells remaining on the slide
following pepsin treatment, then the treatment
was too harsh. If many cells have lifted off,
the treatment was too harsh. Adjust the time or
pepsin concentration as required.
Paraffin sections. The diameter of nuclei in
various tissue types will vary between 4 and
8 µm. The estimated diameter for the tissue
of interest should be ascertained from the literature and sections of the same thickness or
2 to 4 µm greater should be cut, so that it is
likely that the entire nucleas is included in the
section.
Pepsin digestion works for most tissue
types. Occasionally, the tissue will exhibit persistent green autofluorescence of nuclei and
no signal (see Table 22.4.19). Variation in histology fixation procedures and quality control
performed by the hospital pathology laboratory necessitates careful assessment of treatment times and/or concentration of pretreatment (also see Table 22.4.9). Certain tissue
sections are more resistant to effects of protein digestion than others. This may be due
to the tissue type or amount of protein crosslinking elicited by excessive formalin fixation. If surplus fresh tissue can be obtained,
then ethanol or paraformaldehyde fixation will
cause less cross-linking and provide better results for most research applications. It should
also be noted that normal tissues are often more
refractory to protein digestion than tumor material derived from the same tissue type. If there
are a limited number of slides remaining from
the patient sample of interest it is sometimes
helpful to use a superfluous slide from another
sample of the same age and cell type to derive the optimal digestion parameters with respect to aging and storage. More comprehensive guidelines for improved paraffin results
are summarized by Hyytinen et al. (1994).
Anticipated Results
Preparation of probes
Successful labeling for FISH will yield
fragment sizes between 200 and 500 bp. Fragments up to 1 kb are acceptable, but larger fragments will contribute to background. Investigators should expect that the actual amount
of DNA aliquoted onto the gel for visualization should be fairly close to the expected
amount loaded onto the gel. The labeling reaction is fairly tolerant to variations in concentration from half the actual amount to two-times
greater than the actual amount.
Once the labeling conditions have been
established and tested in control FISH experiments, the investigator can be confident that
subsequent labeling experiments will proceed
without incident. Minor adjustments must
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Table 22.4.22 Troubleshooting Guide to FISH Probe Labelinga
Problem
Cause
Solution
DNA failed to digest
DNase I not present
Ensure that DNase I was added to the
labeling reaction.
Loss of DNase I activity If DNase I was added, check the expiration
date or change to a fresh aliquot.
Fragments too small
TE buffer used to
dissolve DNA (EDTA
inactivates DNase I)
If TE buffer was used to resuspend the DNA,
reprecipitate the DNA, resuspend in water,
quantify by spectrophotometry, and relabel.
Too much DNase I
added
Pipetting errors are the leading cause of
many experimental problems. Use accurate
pipettors.
DNA starting
Refer back to spectrophotometer readings
concentration inaccurate and gel images of undigested DNA to
determine whether there were any
discrepancies. Contaminating RNA and
proteins may contribute to an inaccurate
reading, such that the actual amount of
starting DNA was less than expected, thus
changing the labeling (digestion) kinetics for
a specified amount of DNA over a specific
time at a specific DNase I concentration.
Relabel and increase the amount of starting
DNA as required
Fragments too long
Fluorescence
In Situ
Hybridization
(FISH)
Starting DNA was
degraded
If the starting DNA already contains
degraded fragments, they will simply
continue to degrade into relatively smaller
fragments. Refer back to the gel of the
undigested DNA and determine the
percentage of degraded fragments present.
Compensate for the fragments that will be
digested away and increase the amount of
DNA required to relabel. Consider PCR
labeling strategies or obtain a better DNA
sample.
Labeling time too long
Be sure that the correct incubation time was
used.
Unstable water bath
temperature
The labeling reaction has been optimized for
a specific temperature. Increasing the
temperature will facilitate faster nicking,
while decreasing the temperature will slow it.
Insufficient DNase I
Use accurate pipettors (see above).
DNA starting
As described above, check the
concentration inaccurate spectrophotometer reading and gel image. If
too much DNA was added to the labeling
reaction, the same amount of DNase I used to
nick a given amount of DNA in a specific
time will increase when there is more DNA
present. The labeling of this DNA can be
salvaged by continuing to incubate for 20 to
30 min (or as required). Be sure to spike the
reaction with fresh enzymes.
continued
22.4.48
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Table 22.4.22 Troubleshooting Guide to FISH Probe Labelinga , continued
Problem
Cause
Solution
Starting DNA was
dissolved in TE buffer
Check whether the starting DNA was
dissolved in TE as this will inhibit the
nicking of DNA. The labeling of this DNA
can be salvaged by continuing to incubate as
required. Be sure to spike the reaction with
fresh enzyme.
Decreasing activity of
DNase I
Over time, the specific activity of DNase I
will decrease. If this is the case, increase the
amount of DNase I used in the reaction,
while keeping the same incubation period.
Otherwise, keep the same DNase I
concentration and increase the labeling time.
No fragments present Absence of DNA added Make sure that DNA was added to the
to reaction
reaction
Insufficient DNA added As discussed above, check to see that the
concentration of DNA is accurate and
pipettors are accurate.
Starting DNA degraded If a significant portion of the starting DNA
was degraded to begin with, as shown by gel
electrophoresis, this will cause an increase in
fragmentation speed, with the smaller
fragments running off the gel. A new DNA
specimen should be used, or the investigator
should consider PCR methods.
Too much DNase I
added
Expected amount of
DNA starting
DNA too low/too high concentration inaccurate
(but fragmented to the
right size)
PCR failed
Check that the proper DNase I concentration
and/or volume was used in the labeling
reaction.
Assess using a concentration standard. If the
actual amount of DNA is less than expected,
as determined by gel electrophoresis, then
the starting amount of DNA was insufficient
or caused by some of the factors described
above. If the investigator chooses to relabel,
increase the amount of starting DNA as
required. If the actual amount of DNA is
greater than expected, again as determined by
gel electrophoresis, then the starting amount
of DNA was incorrect. The investigator may
choose to proceed and use this probe for
FISH, but the overall incorporation of label
into the DNA will be lower since the labeling
reaction contains sufficient dNTPs for a
specified DNA quantity. If the investigator
chooses to relabel, decrease the amount of
starting DNA material as required.
Amount of template,
Refer to APPENDIX 3F and APPENDIX 2A.
pH, or MgCl2
concentration incorrect,
or quality of template
poor
a The guide assumes that the labeling conditions have generally been optimized but occasionally anomalies occur.
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22.4.49
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be made when using new reagents. Once a
probe has been FISHed, the signal should be
clearly visible with little or no background
or cross-hybridization. The investigator should
be aware that other factors, such as hybridization efficiency, slide quality, antibody quality,
stringency of post-hybridization washes can
also affect the signal strength.
Preparation of slides
Optimal pretreatment of cytogenetic or
paraffin-embedded slides should produce
slides that are relatively free of cytoplasm/cellular debris, free of paraffin, and have
adequately denatured DNA for FISH. The
samples should not be autofluorescent.
Hybridization
Due to the multistepped nature of FISH, it is
difficult to determine the success of hybridization until the entire assay is completed. Successful hybridization will depend on adequate
denaturation of the target DNA on the slide,
as well as adequate denaturation of the probe.
Proper suppression of repetitive sequence elements with Cot-1 DNA can increase the
specificity of hybridization. The specificity of
hybridization is also regulated by stringency
factors described above (see Background information; also see Critical Parameters). Successful hybridization will allow the identification of chromosomal location of the probe
sequence.
Post-hybridization washes and detection
The protocols described provide a basic
framework for adjusting post-hybridization
washes and detections. Under ideal conditions, the stringency of the post-hybridization
washes is sufficient to adequately remove any
unbound/nonhybridized probe from the target
DNA. The specificity of the probe and the suppression of repetitive elements during preannealing will also decrease the number of crosshybridization signals. The concentration of
detergent used in the washes is also optimized
to remove any unbound antibodies, although
minor modifications, as required, should be
made when using a different brand, lot, or type
of antibody. The result, upon visualization with
a microscope, is a clean preparation showing
minimal background and an easily visualized
signal(s) on the DNA target.
Time Considerations
Fluorescence
In Situ
Hybridization
(FISH)
Preparation of probes
The most time consuming portion of labeling for FISH is in the optimization of the la-
beling procedure if using an in-house protocol.
As discussed previously, commercial kits for
labeling are available and give relatively consistent results.
Preparation of slides
The preparatory work for preparing cytogenetic slides requires minimal time and the protocol is fairly robust. Optimization for paraffinembedded sections requires more careful time
and trial-and-error. The preparation of cytogenic slides for FISH can be accomplished in
∼1 hr. Once optimized conditions have been
obtained for paraffin-embedded tissues, it too
can be accomplished in 1 to 2 hr. However,
initial optimization may require several experimental attempts.
Hybridization
The hybridization step of FISH typically
occurs overnight (i.e.,. DNA probes), ∼18 to
24 hr at 37◦ C. The time may be lengthened to
48 or 72 hr (as in the case with weekends), with
no adverse affects. Hybridization with PNA
probes require less time, at least 1 hr at room
temperature. Caution, however should be taken
that the slides are adequately sealed such that
the probe in hybridization solution does not
evaporate. Maintaining a slightly dampened
slide container will help alleviate this problem.
Post-hybridization washes and detection
The time allotted for post-hybridization
washes and detection will vary depending on
the type of DNA probe used in the experiment.
Directly labeled probes (DNA or PNA probes)
require very little bench time since no antibody detection is required; typically they take
up to 30 minutes. Probes are simply washed using high stringency washes as described in the
methods. Indirectly labeled probes require incubations with antibodies (primary, secondary
and/or tertiary), with the final incubation with
a fluorochrome-conjugated antibody, as well
as detergent washes and/or blocking steps between each antibody incubation. If the signal requires amplification, the bench time increases as each antibody incubation should last
at least 30 minutes. A wash protocol using an
indirectly labeled probe with signal amplification will range from 3.5 to 4 hours.
Acknowledgments
The authors would like to thank Elena
Kolomeitz, Jana Karaskova, Paula Marrano,
Ajay Pandita, Bisera Vukovic, and Zong
Mei Zhang from the Department of Cellular and Molecular Biology, University Health
22.4.50
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Network, Ontario Cancer Institute in Toronto,
Ontario, Canada.
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Internet Resources
http://www.appliedbiosystems.com
The Applied Biosystems website. This company supplies PNA probes and custom orders PNA probes.
http://www.cytocell.co.uk
The Cytocell website. This is a commercial supplier
of DNA FISH probes
Cell Biology of
Chromosomes
and Nuclei
22.4.51
Current Protocols in Cell Biology
Supplement 23
http://www.biochem.roche.com
The Roche Molecular Biochemicals website. This
company is a supplier of antibody and labeling
reagents.
http://www.vysis.com
The Vysis website. This company is a supplier of
commercial DNA FISH probes.
Contributed by Jane Bayani and
Jeremy A. Squire
University of Toronto
Ontario, Canada
Fluorescence
In Situ
Hybridization
(FISH)
22.4.52
Supplement 23
Current Protocols in Cell Biology