Mapping Cloned DNA on Metaphase
Chromosomes Using Fluorescence in situ Hybridization
Fluorescence in situ
hybridization (FISH) provides a
quick means of placing almost any recombinant DNA
clone onto a physical map. Clones can be mapped onto
banded metaphase chromosomes for regional localization,
to assign new markers to a chromosomal
segment, to validate clones selected using previously
assigned markers, or to characterize chromosomal
In genome mapping, unrecognized clone chimerism
causes considerable problems with contig assembly
and can result in much wasted effort when trying
to extend or link contigs. For example, as many as
30-40% of YAC clones may contain coligated
sequences from different genomic regions (Selleri et al.
1992). The shift to large-insert, single-copy plasmids
such as PACs and BACs has substantially reduced the
problem of clone chimerism. However, large-scale
genome sequencing projects are revealing significant
levels of regional homology that can confound
mapping analyses at all levels (Cheung et al.
Bailey et al.
, 2002). Preliminary FISH screening of
clones is a simple and useful precaution.
DNA clones can also be ordered by FISH at increasing
levels of resolution. Clones separated by 1-2Mb
can be ordered by pairwise, two-color FISH on metaphase
chromosomes (Trask et al.
, 1991; Inazawa et al.
1994). Hybridization of sets of three clones to interphase
nuclei provides mapping information down to
about 50-100kb (Lawrence et al.
, 1990; Trask et al.
1991), while the highest resolution is for overlapping
clones on linear DNA molecules (Raap et al.
II. MATERIALS AND
DNase I (D-4527), DNA polymerase I (D-9380),
bovine serum albumin (BSA, B-4287), dextran sulfate
(D-8906), Denhardt's solution (D-2532), dithiothreitol
(DTT, D-9779), SDS or sodium lauryl sulfate (L-4390),
mouse antidigoxin FITC (F-3523), goat anti-mouse-
FITC (F-0257), rabbit anti-mouse-FITC (F-7506), and
DAPI (D-9542) are from Sigma. Deoxyribonucleotides
solutions of dATP, dCTP, dGTP, and dTTP)
are from Amersham Biosciences (Cat. No. 27-2035-01).
Human Cot-1 DNA (Cat. No. 15279011) is from Invitrogen.
Formamide is from Fluka (Cat. No. 47670).
Molecular biology grade mixed bed resin AG 501-X8
is from Bio-Rad (Cat. No. 143-6424). Biotin-16-dUTP
(Cat. No. 1093070) and digoxigenin-11-dUTP (Cat. No.
1573152) are from Roche Diagnostics.
Berliner Glas microscope slides are from H. V. Skan
Ltd., Solihull. Glass coverslips (No. 1-22 × 22mm, 22
× 32mm, 22 × 50mm) and Nescofilm are from Fisher
Scientific. Rubber cement can be obtained from art
suppliers or from cycle shops (e.g., Halfords). Plastic
syringes are from Becton-Dickinson, and 0.2-µm
syringe filters are from Nalgene (Cat. No. 190-2520).
Plastic slide boxes (50 slide capacity, Cat. No.
406028600), self-indicating silica gel (Cat. No. 300624V), Tween 20 (Cat. No. 66368 4B), glycerol (Cat.
No. 101186M), 96% ethanol, and 99% ethanol are from
BDH (VWR International).
Fine forceps suitable for handling slides and coverslips
comfortably, without mishap (13mm, Cat. No.
E12), glass Coplin jars (Cat. No. E94), Hellendahl jars
(Cat. No. E95), slide racks (Cat. No. E89.03, E89.055),
and diamond marking pencils (Cat. No. E17) are from
Raymond A. Lamb.
Cy3-streptavidin (Cat. No. PA43001) is from Amersham
Biosciences. Texas red avidin DCS (Cat. No. A-
2016), biotinylated antiavidin (Cat. No. BA-0300), and
Vectashield (Cat. No. H-1000) are from Vector Laboratories
Ltd. Filter paper discs, grade 4 (Cat. No. 1004
240), are from Whatman. Nonfat milk powder (e.g.,
Marvel) is widely available. Similarly, nail varnish
from any inexpensive source should be adequate.
DNA concentrations are determined using a Hoefer
TKO minifluorimeter. The slide drying bench (Cat. No.
E18.1) is from Raymond A. Lamb. Also needed are at
least two water baths (to be set at 37 and 65°C), a
microcentrifuge for both 0.5- and 1.5-ml tubes, a 37°C oven, and facilities for performing agarose gel electrophoresis,
as well as countup/countdown timers
(e.g., Smiths), thermometers, micropipettes and sterile
tips, domestic air-tight plastic freezer boxes, and lintfree
tissues (Kimwipes). Incubation chambers are
prepared using 245 × 245-mm2
bioassay dishes (Nunc,
Cat. No. 240835). Racks to accommodate a maximum
of 16 slides are made by trimming four plastic 10-ml
pipettes to fit and fixing them in place with rubber
We use a Zeiss Axioskop epifluorescence microscope
fitted with a 100-W mercury arc lamp, triple
bandpass/dichroic filter blocks for both rhodamine (or
Cy3) and Texas red (Chroma Technologies, sets 82000
and 83000), and a motorized excitation filter wheel
(Ludl) and a Photometrics KAF1400 cooled CCD
camera controlled by SmartCapture imaging software.
Similar imaging systems can be obtained from Applied
Imaging. SmartCapture software is supplied by Digital
A. Probe Labeling
- 10× nick translation buffer: 0.5 M Tris-HCl, pH 7.5,
0.1M MgSO4, 1mM DTT, and 500 µg/ml BSA. To make
10 ml, mix 5 ml 1M Tris-HCl (pH 7.5), 1ml 1M MgSO4,
10µl 1M DTT, 500µl 10mg/ml BSA, and make up to
10ml with sterile deionized water. Store 1-ml aliquots
- DNase I stock solution: Resuspend 10,000 units in
1ml 0.3 M NaCl and add 1ml sterile glycerol. Store at
- DNase I working solution: To make 1ml, mix 100
µl 10x nick translation buffer with 400 µl sterile deionized
water and 500µl sterile glycerol; then add 1µl
DNase I stock solution. Mix thoroughly and store at
- 0.5 mM dNTPs: To make 1200µl, mix 2 µl each of
100mM dATP, dCTP, and dGTP and then add 1194µl sterile deionized water. Store 50-µl aliquots at -20°C.
- 1mM biotin-16-dUTP or digoxigenin-11-dUTP
- DNA polymerase I (10 units/µl)
- 0.5M EDTA, pH 8.0: To make 100ml, mix 18.61 g
EDTA in 80ml deionized water, adjust to pH 8.0 with
NaOH (the salt will not dissolve until near pH 8), and
make up to volume with deionized water. Sterilize by
- 1M Tris-Cl, pH 7.4: To make 100ml, dissolve
12.11 g Tris base in 80ml deionized water, adjust to pH
7.4 with HCl, and make up to volume with deionized
water. Sterilize by autoclaving.
- 3M sodium acetate, pH 7.0: To make 100ml,
dissolve 24.6g sodium acetate (anhydrous) in 80ml
deionized water, adjust to pH 7.0 with glacial acetic
acid, and make up to 100ml with deionized water.
Sterilize by autoclaving. Use small aliquots as working
stock, replacing frequently.
- Absolute ethanol: Store in a sterile 50-ml Falcon
tube at -20°C.
- 70% ethanol: Store in a sterile 50-ml Falcon tube
- TE buffer (10mM Tris-Cl, 1mM EDTA): To make
100ml, mix 1ml of 1M Tris-Cl, pH 7.4, and 0.2ml of
0.5M EDTA and make up to 100ml with deionized
water. This is best made prior to sterilizing the stock
solutions of Tris and EDTA. Sterilize by autoclaving.
B. in situ Hybridization
- Prepare a water bath at 14°C. A robust polystyrene
box (or ice bucket) with a lid is a suitable container
(although some are not reliably water tight).
- Place approximately 1 µg of each DNA sample to
be labeled in 1.5-ml microfuge tubes and make up the
volumes to 10µl with sterile deionized water. Stand
- Prepare 15µl labeling master mix for each
sample, allowing a little extra for dispensing. For 4 µg
DNA, take 10µl nick translation buffer, add 34µl sterile
deionized water, 7.5µl dNTPs, 2.5µl 1mM biotin-16-dUTP, 4µl DNase I working solution, and 2µl 10 U/µl
DNA polymerase I. Mix thoroughly and pulse microfuge
to collect all the solution in the base of the tube.
Stand on ice.
- Add 15 µl of master mix to each DNA sample and
mix by pipetting several times.
- Incubate the samples at 14°C for 40-60min. The
exact length of time must be determined for each new
preparation of DNase I working solution.
- Transfer the tubes to ice and add 2.5µl 0.5M EDTA to each, mixing quickly with the pipette.
- Add 2.5 µl 3M sodium acetate and 1ml cold 100%
ethanol to each tube. Mix by inversion.
- Incubate at -20°C overnight.
- Microfuge the tubes for 10min at maximum
speed. Remove the supernatants carefully so as not to
disturb the visible pellets.
- Carefully add 1ml cold 70% ethanol to each
tube. Microfuge again for 10 min. Carefully aspirate off
the supernatant from the transparent pellets and air
dry. Do not overdry.
- Add 10µl TE buffer to each tube and stand on
ice for 15 min. Flick mix to resuspend.
- Run 2-µl aliquots in a 1% agarose gel to assess
the probe fragment length. A smear between 200 and
800bp is satisfactory.
- Store the probes at -20°C.
- 50% dextran sulfate: To make 50ml, weigh 25g
dextran sulfate into a graduated 100-ml pyrex bottle,
add 20ml deionized water, and heat in a 65°C water
bath until dissolved. Make up to volume with deionized
water and sterilize by autoclaving. Dispense
10-ml aliquots into sterile 50-ml Falcon tubes and store
- Deionized formamide: Add 5 g Bio-Rad mixed bed
resin AG 501-X8 to 100ml formamide and stir in a
fume hood for 60min. Allow the beads to settle and
decant the formamide. Store aliquots at -20°C.
- 20× SSC (3 M NaCl, 0.3 M sodium citrate): To make
1 liter, dissolve 175.2g sodium chloride and 88.2g
trisodium citrate in deionized water and make up to
- 10% SDS: To make 50ml, dissolve 5g sodium
dodecyl sulfate in 50ml deionized water. Sterilize by
filtering through a 0.2-µm filter.
- 0.5M Na2HPO4: To make 100 ml, dissolve 7.098 g
Na2HPO4 in deionized water and make up to volume.
Sterilize working aliquots by filtration.
- 0.5M NaH2PO4: To make 100ml, dissolve 7.8g
NaH2PO4.2H2O in deionized water and make up to
volume. Sterilize working aliquots by filtration.
- Hybridization buffer: To make 50 ml, thaw a Falcon
tube containing a 10-ml aliquot of 50% dextran sulfate,
add 25 ml deionized formamide, 5 ml 20× SSC, 1ml 50x
Denhardt's solution, 4ml 0.5M sodium phosphate
buffer, pH 7.0 (2.308 ml Na2HPO4, 1.692ml NaH2PO4),
0.5 ml10% SDS, and 4.5 ml sterile deionized water. Mix
very thoroughly by inversion, preferably on a rotator
for 10min. Dispense into 1-ml aliquots and store at
-20°C. This should be stable for at least 1 year. Mix
newly thawed aliquots thoroughly by vortexing.
- Cot-1 DNA: Supplied at 1mg/ml.
- 70% formamide: To make 100 ml, mix 70 ml formamide
and 30ml 2× SSC. Store at 4°C between uses
and replace weekly. (Deionizing the formamide is not
- 70% ethanol in a Hellendahl jar: Store at -20°C. Replace weekly.
- Ethanol series: Hellendahl jars containing 70%
ethanol (2×), 90% ethanol (2×), and 100% ethanol (1×).
- Coverslips: 22 × 22-mm, 22 × 50-mm coverslips
immersed in 100% ethanol in an air-tight plastic
C. Visualization of Hybridization
- Prepare a Coplin jar or Hellendahl jar containing
70% formamide. Place in a water bath and turn the
temperature to 65°C. (The glass jars will not tolerate
sudden temperature changes and should always be
allowed to warm gradually.)
- Set a second water bath to 37°C.
- Prepare the probe hybridization mixes by adding
10µl hybridization buffer to 0.5-ml microfuge tubes
(dispensing will be aided by first warming the
hybridization buffer). Stand the tubes on ice. Add I or
2µl Cot-1 DNA to each tube, depending on whether
one or two probes are to be added. Add 0.5µl of
biotinylated probe and/or 0.5µl of another digoxigenin-
labeled probe. Mix thoroughly by flicking the
tubes and pulse microfuge the solutions to the bottom
of the tubes.
- Denature the probe mixes at 65°C for 10min,
ensuring that the tubes are fully sealed and not likely
to take up water.
- Transfer the probe mixes to the 37°C water bath
to preanneal for at least 20min (to several hours).
- Check that the 70% formamide has reached 65°C
using a thermometer reserved for the purpose.
- Carefully immerse slides, paired back to back,
into the 70% formamide at 5-s intervals. Start a countup
timer as the first pair of slides is immersed. Take care to hold the slides clear of the steam from the water
bath while preparing to place them in the formamide.
- While the slides are denaturing, remove the
Hellendahl jar of 70% ethanol from the freezer.
- Denature the slides for 2min. As the denaturation
time elapses, remove the pairs of slides from the
formamide and drain briefly against the inside of the
jar. Immerse the slides, agitating briefly, in the cold
- After 60s in the cold 70% ethanol, transfer the
slides with occasional agitation through an ethanol
series at room temperature: 60 s each in successive jars
of 70, 70, 90, 90, and 100% ethanol.
- Separate the slide pairs and carefully wipe the
backs of the slides dry. Air dry the slides standing in a
rack on a slide-warming bench at 37-40°C.
- Prepare the required number of 22 × 22-mm
coverslips by removing them from the 100% ethanol
and lightly polishing them dry with a lint-free tissue.
Place the coverslips on a clean tissue on or by the slidewarming
- Place the dry slides flat on the slide-warming
bench. Make sure that all are labeled and numbered
- After the probes have preannealed at 37°C for
at least 20min, transfer the first hybridization mix to
its target slide. As the volumes can be difficult to gauge
accurately, set the micropipette for 12 µl and be careful
to avoid bubbles when drawing up and discharging
the mix onto the slide.
- Using fine forceps, gently lower a clean coverslip
over the mix. If bubbles are present, these can
usually be disrupted by surface tension by allowing
the hybridization mix to spread out under the upper
half of the coverslip while the lower edge of the
coverslip is still supported by the tip of the forceps.
It is usually not worth trying to remove every
last bubble as this may damage the chromosome
- Repeat these steps for each remaining probe
mix. If there are a large number to be processed, it may
be preferable to station the slide bench next to the 37°C
water bath and remove each mix from the water bath
as required. Alternatively, the hybridization mixes can
be placed on ice, but this can make the mixes more
viscous and difficult to pipette.
- Seal the edges of the coverslips with rubber
cement. If the hybridization mix has not reached the
edges of a coverslip because the coverslip cannot lie
flat, it will be necessary to prevent the rubber cement
from being drawn under the coverslip. If the coverslip
cannot be flattened by gentle pressure over the high
point, fill in the gap with extra hybridization buffer
- Place on a tray and hybridize overnight in a
- 50% formamide: To make 200 ml, mix 100 ml formamide
and 100ml 2× SSC. Store at 4°C between uses
and replace weekly.
- 2× SSC: To make 1 liter, take 100ml 20x SSC and
make up to volume with deionized water.
- 4 × TNFM: To make 1 liter, take 200ml 20x SSC,
add 700ml deionized water, 500 µl Tween 20, and 50 g
nonfat milk powder and mix vigorously, then make up
to 1 liter with deionized water. Filter through several
layers of Whatman No. 4 filter paper. (The solution
should be a slightly translucent yellow-green color. If
it remains cloudy try another brand of nonfat milk
- Immunochemical solutions: Allow 100µl per slide
plus 50-100µl excess. Protect from strong light.
- For biotinylated probes, make 2 aliquots of
4µg/ml streptavidin-Cy3 and one aliquot of
4 µg/ml biotinylated antiavidin.
- For digoxigenin-labeled probes, make a 1:500-1:1000 dilution of mouse antidigoxin-FITC
and a 1:250 dilution of rabbit anti-mouse-FITC.
- For dual-color detection, use avidin-Texas
red instead of streptavidin-Cy3. Combine
the biotinylated antiavidin with the mouse
antidigoxin-FITC, and the second avidin-
Texas red with the rabbit anti-mouse-FITC.
- 4 × T: To make 200 ml, mix 40 ml 20x SSC, 160 ml
deionized water, and 100µl Tween 20.
- DAPI staining solution: Prepare a Hellendahl jar
with 75ml of 2× SSC and 6µl 1mg/ml DAPI. Protect
from light by wrapping in aluminium foil.
A. Probe Labeling
- Place three Coplin jars or Hellendahl jars containing
2× SSC and two jars of 50% formamide into a
water bath and warm to 42°C.
- Place a jar of 4 × TNFM to warm in a 37°C oven.
- Prepare a humidified chamber by placing damp
tissues in the bottom of an incubation dish and warm
in a 37°C oven.
- Prepare immunochemical solutions diluted in 4
x TNFM according to the haptens used. Stand at room
temperature for 10min and then microfuge for 10min
to pellet any protein complexes that might contribute
to nonspecific background.
- Remove the rubber cement from the slides using
fine forceps and soak off the coverslips in the first jar
of 2× SSC at 42°C, allowing approximately 5 min.
- Transfer the slides to the first jar of 50% formamide
for 5 min.
- Repeat this incubation in the second jar of 50%
formamide and then in each of the remaining jars of
2× SSC, agitating the slides briefly after each transfer.
- Transfer the slides to the prewarmed jar of 4 x
TNFM and incubate for 5-10min at 37°C.
- Remove the slides one by one from the 4 × TNFM
and drain briefly, wipe the back, and blot excess liquid
from the top and bottom edges. Place the slides on a
rack in the prepared humidified chamber and apply
100µl of the first immunochemical solution (streptavidin-
Cy3 or mouse antidigoxin-FITC or avidin-Texas
red). Overlay with a 25 × 50-mm strip of Nescofilm.
Note: The slides must not dry out at any point during
- Incubate the slides at 37°C for 20-30min.
- Meanwhile, discard the three 2× SSC wash solutions
and replace with 4 × TNFM, allowing the jars to
warm to 42°C again.
- Discard the Nescofilm strips, drain the slides,
and rinse them in the three changes of 4 × TNFM at
42°C for 5 min each.
- Repeat step 9 using the second antibody layer
(biotinylated antiavidin, rabbit anti-mouse-FITC, or
biotinylated antiavidin plus mouse antidigoxin-FITC).
- Repeat steps 10-12.
- Repeat step 9 using the final immunochemical
layer (streptavidin-Cy3 or avidin-Texas red plus rabbit
anti-mouse-FITC). If using only digoxigenin-labeled
probes, proceed to step 17.
- Repeat steps 10-12.
- Wash twice in 4 × T at room temperature.
- Stain the slides in DAPI for 2-3 min. Transfer the
slides to a jar containing 2× SSC, rinse briefly, and pour
off the 2× SSC (holding the slides in place with a
gloved finger or forceps across the top of the jar). Rinse
briefly with deionized water and pour off.
- Dehydrate the slides by passing through an
ethanol series, gently agitating 30-60 s in each of 70, 70,
90, 90, and 100% ethanol. Air dry.
- Lightly polish 22 × 50-mm coverslips and lay
them out on flat absorbant tissue. Apply 25-30µl
antifade solution to each coverslip.
- Invert each slide over a coverslip, rest the
bottom edge on the tissue, and gently lower until the
slide touches the antifade droplet. Allow the coverslip
to lift up to the slide before laying it flat. When the
antifade has spread fully, gently blot any excess. Seal
the edges of the coverslips with nail varnish. The slides
can now be stored in the dark at 4°C for many weeks.
- View the slides using an epifluorescence microscope
equipped with the excitation and emission filters
appropriate for the fluorochromes used.
Whole clone DNA preparations are usually the best
material for nick translation. Bacterial clone DNA isolated
by standard alkaline lysis should be satisfactory.
Most matrix-binding protocols, as used by commercial
kits, do not give very high yields for larger insert
clones such as cosmids, PACs, or BACs (often only
enough for one or two labeling reactions from 10 ml of
bacterial culture) so be sure that sufficient DNA (of a
concentration of at least 100µg/ml) is obtained. Total
yeast DNA gives good results for YACs, although
the yeast genome may contribute excess ribosomal
sequences that may not be fully suppressed during
Different grades of DNase 1 have different levels of
activity. New working solutions should be titrated to
determine the best incubation time. A 50-µl reaction
containing 2µg DNA and 2µl DNase I working solution
in 1× nick translation buffer can be sampled at five
intervals (e.g., at 20, 30, 40, 50, and 60min) and the
DNA fragments compared by electrophoresis. Even
less expensive products, such as DN-25 (Sigma), are
suitable for use after titration, allowing for the relative
number of units per milligram of protein in the
working solution. Enzyme activity is also affected by
the amount of Ca2+
present in the reaction. Usually, sufficient
is present in the original DNase I stock;
however, higher grades of enzyme may need additional
to maintain active enzyme conformation
during the nick translation reaction.
Approximately 70-80% of the original DNA is
usually recovered after ethanol precipitation, giving
a final probe concentration of 70-80ng/µl. Probe concentrations
can be verified by DNA fluorimetry if
desired. Ethanol precipitation is not absolutely
required for probe preparation and can be omitted
after labeling repetitive DNA clones, such as chromosome-
specific satellite DNAs, as these are routinely
used at much lower hybridization concentrations
(0.5-1ng/µl) than "unique-sequence" clones (2.5-
5ng/µl). Ethanol precipitation enables the labeled
probe to be concentrated so that it can be used without
further precipitation prior to hybridization. This
ensures more consistent results from one experiment
Large insert clones can be mapped efficiently using
direct labeling with fluorochrome-conjugated dUTPs.
This avoids the time-consuming immunochemical
detection steps described here, but may not be as effective
in revealing smaller signals at secondary chromosomal
The hybridization buffer can be dispensed more
accurately after it is warmed to 65°C. The components
may separate during freezing and the warmed solution
should be mixed thoroughly before use. Probes
will last several years if handled aseptically and stored
at -20°C. They should be thawed and kept on ice when
in use and returned to the freezer without delay (do
not store in frost-free freezers). Routine metaphase
mapping throughput can be increased by using pairs
of biotin- and digoxigenin-labeled probes on two separate
spots of metaphase cells on a single slide. The
slide can be mounted in antifade under a single large
|FIGURE 1 (A) Two-color hybridization of two cosmid
mapping to chromosome 22. One cosmid was
detected with avidin-Texas red (red) and
the second cosmid was
labeled with digoxigenin and
visualized using FITC-conjugated
antibodies (green). (B)
Black-and-white inverted image of the
in A. The chromosomes appear essentially G
although the 9qh region is dark in this example.
The 4 × TNFM washes and the immunochemical
incubations do not need to be performed at 37-42°C as
the results are usually satisfactory at room temperature.
However, it is preferable to protect the slides from
bright light, and in a busy laboratory the slides can
often be tucked away more safely and neatly by
leaving them in an incubator or water bath.
If the clones to be mapped all localize to a small
region, it may not always be possible to use a doubleprobe
protocol as interpretation can be complicated
by poor-quality probe. When performing dual-color
analysis, it is easier to use fluorochromes that are spectrally
well separated. Cy3 is a very stable, bright
orange fluorochrome, but it can be difficult to use with
FITC, as the Cy3 signals are often visible through FITC
filters. This is a particular problem when imaging with
black-and-white CCD cameras as strong Cy3 signals
can be confused with FITC signals. Texas red is preferred
for combinations with FITC (Fig. 1A).
The DAPI staining reveals a clear banding pattern,
which permits chromosome identification, provided
the slides are not overdenatured. If a digital image of
the DAPI-stained metaphase is converted into a blackand-
white inverted image, the chromosomes appear
similar to a conventional G-banded metaphase
- If no signal is obtained, one possibility may be
insufficient DNA. It is not unusual for spectrophotometric
determinations of DNA concentration to be as
much as 5-10 times higher than the real figure. Alternatively,
the original DNA may have deteriorated.
These problems are monitored easily by running
approximately 100-200ng of the original DNA next to
the 2-µl aliquot of the precipitated nick-translated
product in a standard 1% agarose gel. This will also
reveal whether bacterial clones retain the expected size
insert. When using labeled whole yeast DNA, it may
be necessary to increase the amount of probe used per
hybridization to ensure that there is sufficient YAC probe present. Usually 80-100 ng of whole yeast probe
will ensure clear YAC signals. Using additional
amounts of probe may also assist the mapping of
smaller insert clones.
Another major problem is poor hybridization
efficiency because of poor-quality slide preparation.
Probe accessibility is critical. Good-quality metaphase
preparations, with well-spread chromosomes and a
minimum of residual cytoplasm, are far more useful
than any amount of protease pretreatment. RNase A
digestion is not normally necessary at all.
It is always possible that the clone you are trying to
map will never hybridize to your metaphase spreads.
For example, human monochromosomal libraries
made from material flow sorted from somatic cell
hybrids may contain a small proportion of nonhuman
clones. Try hybridizing a larger amount of probe
(100-200 ng) without any Cot-1 DNA and no preannealing
at 37°C. The abundant Alu short, interspersed
repeat sequences are distributed throughout the
human genome and should be present in most
genomic clones. In the absence of preannealing, Alu
signals should be distributed all over the chromosomes,
concentrated in the G-light chromosome R
It also appears that about 25-30% of cDNA clones
cannot be localized by FISH (Korenberg et al., 1995).
This may be due to the relatively short genomic
extent of the target gene. Mapping cDNAs is a much
more demanding process and every step must be
rigorously optimized. Protease digestion around the
chromosomal DNA may be useful here, but it must be
carefully controlled to avoid loss of chromosome morphology
(Fan et al., 1990). More sensitive detection
systems, such as the use of tyramides, may also be
useful (Raap et al., 1995).
- If the cloned DNA does not cut well after nick
translation, the large probe fragments will result in
excessive background hybridization, which is seen as
large, bright spots of signal all over the chromosomes
and nuclei. Large spots of signal that appear to float
above the spreads are nearly always indicative of
excessive probe fragment length. Because DNase I
digestion is affected by contaminants in the DNA, it
may be necessary to repurify the sample before repeating
nick translation. DNA that digests with restriction
enzymes should be satisfactory. Try increasing the
duration of the next nick translation reaction by at least
10 min. Check the concentration of your DNA also; the
labeling reaction may have contained far too much
If the probe length is in the acceptable size range, it
may be that one of the immunochemicals has deteriorated.
These should always be kept in small aliquots
and handled as aseptically as possible. Depending on
the manufacturers' recommendations, it should be
possible to store most reagents at -20°C with a single
working aliquot stored at 4°C. Avoid repeated
freeze-thawing and, for this reason, do not store
reagents in frost-free freezers. Using immunochemicals
at too high a concentration can result in generalized
nonspecific background over the slide. Similarly,
precipitation of immunochemical complexes will
occur if the slides are allowed to dry out during the
Certain clones may require increased amounts of
Cot-1 DNA to suppress apparently nonspecific
hybridization to other chromosomal regions. Another
potential source of nonspecific background may be
residual material in the fixed metaphase suspensions.
This may be reduced by washing the cells in several
changes of methanol/acetic acid fixative before
preparing more slides. We routinely postfix slides in a
Coplin jar of fixative, air dry, and then dehydrate
through a fresh ethanol series before a final fixation in
My thanks to Dr. Nigel Carter for useful discussions
over the manuscript.
BAC Resource Consortium (Cheung et al.
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