Southern Transfer and Hybridization
Labeling Probe and Southern Hybridization (Part 3)
Molecular Biology Lab #8
Background:
The process of labeling and hybridization of Southern blots is performed in a series of
steps over several days. The first step is called prehybridization. During this step the
membrane containing the DNA is pretreated with a buffer containing blocking reagents
such as albumin or salmon sperm DNA. These treatments block any nonspecific ‘hot
spots’ on the membrane that might bind to probe nonspecifically. Often, prehybridization
is performed for 30 min using the same buffer that will be used subsequently for
hybridization. If prehybridization is inadequate, the blot may have high background with
nonspecific probe binding.
Probe labeling is the next step. There are several methods for labeling probes, but all
methods require that the DNA molecule first be denatured to separate the 2
complementary strands. This is usually performed by boiling the probe DNA for 5 min
then rapidly cooling. Rapid cooling helps to prevent renaturation of the complementary
strands.
The most sensitive method of detection uses probes that are labeled with radioactive 32P
by random priming or nick translation. These methods add a 32P labeled nucleotide (often
deoxy CTP) throughout the probe DNA molecule. The labeling reaction is then passed
through a column that binds unincorporated nucleotides and allows the radioactive DNA
to elute. This probe is very hot (1x108 cpm/g DNA) and can be used to detect single
copy genes with ease. The radioactive DNA that binds specifically to the probe is
detected by placing the membrane next to X-ray film, or analyzing the membrane in a
phosphoimager. The use of 32P is also associated with potential hazards of external
contamination. 32P is a high-energy beta emitter and the researcher must take precautions
to shield the body from radiation. Usually this is done by working behind a small
Plexiglass shield. All of the experimental waste material must be carefully retained and
then disposed, and careful records must be kept. All sources of 32P must be kept under
lock and key in the lab (i.e. lab doors are always locked when using 32P) which can be an
inconvenience. 32P is rather expensive and has a short half-life. In fact, probes must be
used within several days or they decay so much that they are no longer useful.
Another method for labeling probes is by utilizing chemiluminescent detection methods.
These are slightly less sensitive than 32P and are not widely used for Southern or
Northern blotting (RNA detection) when sensitivity is important. However, there is little
or no hazard associated with chemiluminscent probes and they have achieved wide use in
Western blot applications. This method works by directly labeling the probe DNA with
an alkaline phosphatase enzyme. This is achieved by first denaturing the probe DNA and
then adding the enzyme along with a cross linking reagent. The alkaline phosphatase
labeled DNA that specifically binds to the complimentary DNA on the blot is detected by
placing the washed blot in a special substrate solution that alkaline phosphatase can
dephosphorylate. This reaction is associated with the release of chemiluminescence
which can be detected with X-ray film or a phosphoimager. On the positive side, there is
no biological hazard. You can leave the lab doors open, drink coke in the lab, and throw
the waste in the regular trash. The probes also have a long half-life and they can be stored
for weeks to months.
A third method of detection is colorimetric. This method is very similar to the
chemiluminescent protocol, except that the sensitivity is much lower. The probe DNA is
labeled with an alkaline phosphatase enzyme by cross-linking and the labeled DNA
hybridizes specifically to the complementary gene on your membrane. The color
detection method uses a substrate for the alkaline phosphatase enzyme that becomes
insoluble and turns blue when it is cleaved. This method is inherently less sensitive than
chemiluminescence or radioactivity. However, it has the advantage that it requires no
darkroom or film development reagents.
The third step is hybridization of the labeled probe to the membrane. This is usually
performed in a hybridization oven, which carefully regulates temperature and allows the
blot to turn constantly so that it is continually bathed in new hybridization solution. There
is definitely an optimal temperature for performing hybridization reactions. If the
temperature is too low (low stringency) the DNA strands can join rather easily and they
often are able to join even if the complimentary strands don’t match completely. For
example, at low stringency, a probe for the beta actin gene might cross hybridize with a
gene for alpha actin. The alpha actin gene from humans might hybridize to the alpha actin
gene from a dog. The match doesn’t need to be perfect. This can be an advantage if one is
searching for new members of a gene family in the same species (maybe you want to use
low or relaxed stringency to look for new actin genes). It is also useful if you are
searching for a gene in another species that has not yet been cloned (trying to find the
actin gene in the East African Yellow Bellied Snail Darter or some other obscure species.
On the other hand, you can raise the temperature too high (high stringency) so that it is
very difficult for any DNA to hybridize. This is because the temperature is too close to
the DNA melting temperature so the hybrids that form are easily denatured again. Under
these circumstances, only those sequences that are perfectly matched can form stable
hybrids.
How do you choose the right temperature? It depends on many factors including the GC
content of the DNA (GC rich DNA melts differently than AT rich DNA), salt
concentration (high salt means lower stringency), or the presence of formamide (this
lowers the melting temp of DNA). You can also control for stringency of hybridization at
the next step, which is washing the unbound probe away from the blot. If you wash at
high temperature or low salt concentration, you will remove any hybrids that are not
perfectly matched. If you wash at low temperature and/or in high salt, you are leaving
many imperfect hybrids on your membrane. Usually, washing proceeds with a low
stringency wash at first to remove most of the unbound probe. This is followed by a
higher stringency wash. One advantage of using radioactive probes, is that you can easily
monitor how hot your blot is by simply checking it with the Geiger counter. If it were too
hot you would use a more stringent wash. With chemiluminescent or colorimetric
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detection, you will not know whether you have too much background until you actually
develop he blot.
Objectives:
The objective of this lab is to provide experience in labeling DNA probes and using the
probe in a hybridization reaction. The lecture will cover basic aspects of probe labeling
and hybridization.
Materials:
AlkPhos Direct Labeling and Detection Kit
Hybridization buffer
Primary wash buffer (2M urea, 0.1% SDS, 50 mM sodium phosphate at pH 7, 150 mM
Sodium chloride, 1 mM magnesium chloride, 0.2 % blocking reagent)
10X Secondary wash buffer (1M Tris pH 10, 2M sodium chloride)
Alkaline Phosphate Conjugate Substrate Kit
Hybridization oven set at 55ºC
Hybridization bottles and nylon mesh
37ºC water bath
55ºC water bath
Shaker platform
Plastic dishes for washing blots
Pipetters and yellow tips
Ice water bath
Eppendorf microcentrifuge
Previously prepared blot with RNA sample
Eppendorf tubes
Boiling water bath
Dark room with developer and fixer solutions
Film cassettes
X-ray film
Plastic wrap
Methods:
Prehybridization:
1. Preheat the required volume of hybridization buffer to 55ºC in the hybridization
oven. Heat enough buffer for 0.25 ml/cm2 of membrane. Also, preheat the glass
hybridization bottle containing 15 ml of deionized water.
2. Rehydrate the nitrocellulose membrane in water for 5-10 min.
3. Place the blot on a sheet of nylon mesh that is slightly larger (1-2 mm on each
side). Make sure that the DNA side of the blot is facing up. Carefully, roll the blot
and mesh into a roll and slip the roll into the glass hybridization bottle. The DNA
side should face into the hybridization chamber.
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4. Place the cap on the tube and hold the tube horizontally. Turn the tube slowly
until the membrane unrolls inside the tube and is applied to the walls of the tube.
Inspect the tube and membrane carefully to make sure that there are no air
bubbles between the tube wall and the membrane. If air bubbles are present, pull
the filter out and start again. Any air bubbles will lead to high background.
5. Once the filter is applied to the wall of the tube, pour out the water and add about
15 ml of hybridization buffer. Cap the tube and place it into the clips in the
hybridization oven. Turn on the speed control such that the bottle turns slowly
through the oven (one revolution every 10 sec). Check to see that the bottle is
even and that the hybridization fluid covers the bottom of the bottle.
6. Allow the blots to prehybridize (before adding the probe) for approximately 30
min. This step is important to block any nonspecific reactive sites on the blot.
Lack of adequate prehybridization can lead to high background due to nonspecific
binding of probe to the membrane.
Preparation of probe:
7. Prepare the labeled hybridization probe. Dilute 20 l of cross linker solution with
80 l of the water supplied with the kit. This working concentration should be
kept on ice.
8. Dilute HPV-16 DNA to a concentration of 10 ng/l using the water supplied with
the kit.
9. Place 10 l of the diluted DNA sample into an eppendorf tube and denature the
DNA by heating for 5 min in a boiling water bath.
10. Immediately cool the DNA on ice for 5 min. Briefly spin the sample in a
microcentrifuge to collect the contents at the bottom of the tube.
11. Add 10l of reaction buffer to the cooled DNA and mix thoroughly but gently. Be
sure to keep the tube on ice.
12. Add 2 l of labeling reagent and mix thoroughly but gently.
13. Add 10 l of the cross linker working solution. Mix briefly and spin to collect the
contents at the bottom of the tube.
14. Incubate the reaction at 37ºC for 30 min. The probe can be used immediately or
kept on ice for up to 2 hours.
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Hybridization reaction:
15. Add labeled probe to the buffer used for prehybridization. Use about 5-10 ng of
probe per ml of hybridization buffer. Avoid placing the probe directly on the blot.
Remove a small amount of and mix with the probe before returning this mixture
to the bulk of the hybridization buffer.
16. Hybridize at 55ºC for 2 days in the hybridization oven.
Post hybridization washes:
17. Preheat the primary wash buffer to 55ºC (do not overheat). Use this in excess at a
volume of 2-5 ml per cm2 of membrane.
18. Carefully remove the roller bottle from the oven. This is easier if you briefly
switch off the motor for rotation, and then turn it on again after the bottle is
removed. Pour out the hybridization buffer from the roller bottle and add the
preheated primary wash buffer (fill the tube halfway).
19. Place the tube with wash buffer back in the hybridization oven and wash for 10
min at 55ºC.
20. After 10 min, remove the roller bottle and pour out contents. Add more
prewarmed primary hybridization buffer and allow washing in the oven for
another 10 min at 55ºC.
21. Carefully remove the membrane and mesh from the hybridization bottle using
forceps. Place the membrane in a plastic wash dish with 100-200 ml of secondary
wash buffer. Wash with gentle agitation on a shaker platform for 10 min at room
temperature. Several blots may be washed in the same secondary wash buffer
provided that there is enough volume to allow them to move freely.
22. Pour off the wash buffer and add 100-200 ml of fresh secondary wash buffer.
Shake gently for an additional 10 min at room temperature.
Chemiluminescent signal detection:
23. Allow the membrane to drain and briefly dab any excess fluid away with a paper
towel
24. Add 3ml of chemiluminescent substrate to your blot and allow it to saturate the
membrane. This can be done by placing the blot and substrate in an empty yellow
tip box and rocking back and forth.
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25. Dab excess substrate from blot using a paper towel and wrap the membrane
carefully in plastic wrap. Tape the wrapped membrane to the inside of an X-ray
film cassette.
26. Bring the cassette and an unopened X-ray film into the darkroom. Turn on the
safelight and shut the door. Check to see that no light is coming in from outside.
27. Open the X-ray film and carefully place a sheet into the cassette so that it covers
the blot. Close the cassette cover and allow the film to be exposed for 1 hour.
Make sure that the cassette snaps closed (listen for the click).
28. After exposure, return to the darkroom and close the door. Wear latex gloves.
Turn on the safelight and open the cassette. Place the film in the developer for
approximately 30 sec with periodic agitation. You should see the image of the
blot appear.
29. Transfer the developed film to water to remove excess developer and agitate for 2
to 3 min.
30. Place the film in fixer and leave for 2 to 3 min. After this, you can turn on the
light. After 30 min, you can place the film in water and wash for 1 to 2 hours.
Most modern molecular biology labs have an automated film processor that
automatically develops films in 1 to 2 min.
31. Clean up!
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