Download In situ hybridization

In situ hybridization
Next we will be examining the wildtype expression patterns for the single-minded (sim) and snail (sna) genes in
Drosophila w1118 embryos that are 2-6 hours old. To detect the sim or sna mRNA in the embryonic tissue, we
will use in situ hybridization which is the best technique for looking directly at mRNA levels in the tissue and
thus transcriptional regulation in cells. **Note- for practical reasons, time and money, you will be performing
the stain on the w1118 embryos while we make the antisense RNA probe for you from a successfully cloned
recombinant plasmid.
The general procedure starts with a labeled “antisense” RNA probe. This is a single stranded RNA molecule
that is complementary to the mRNA that we wish to detect. To make a labeled probe, we used the plasmids that
you made and synthesized the RNA in vitro in the presence of all the rNTPs, except that the UTP used has a
small digoxigenin (dig) molecule attached.
Figure 1. Structure of digoxigenin-labeled UTP. Used in the synthesis of the antisense RNA probe to allow detection of mRNA
within a cell in vivo through the technique called in situ hybridization.
The embryos are collected at specific time points, in our case from 2 to 6 hours, and then fixed in formaldehyde
which crosslinks proteins. This will preserve the cells in the tissue, freezing them as they are at that exact
moment, in essence like taking a picture of all the cells in the embryo. We then incubate the embryos in a
special hybridization buffer in the presence of our labeled probe. The embryos have been treated with detergent
to open up holes in the membrane large enough to allow our probes to enter the cell. This incubation is done
under optimal conditions for basepairing and requires a lot of time (1-2 days). We then must extensively wash
the embryos to remove unbound probe (antisense RNA that has not basepaired with our mRNA) which would
otherwise create high background staining in the sample.
Once hybridized and washed, we can detect the labeled antisense RNA probe with an anti-digoxigenin antibody
that has been convalently linked with the enzyme alkaline phosphatase (AP). After incubating the antibody
with our samples, we will detect the mRNA by adding the substrate BCIP (bromo-chloro-indolyl-phosphate,
also called X-phos – think of the parallel with X-Gal, bromo-chloro-indolyl-galactoside) and NBT (nitro blue
tetrazolium). The cleavage of BCIP in the presence of NBT leads to a blue precipitate where the enzyme is
located. Therefore any cell expressing eve mRNA will bind the eve antisense RNA probe labeled with dig, will
bind the anti-dig AP antibody, which will then turn blue once BCIP and NBT are added. An example of the
usefulness of this technique is shown below. This is an image of a double ISH on a Drosophila embryo using a
probe against snail mRNA (brown, a mesodermally expressed gene) and rhomboid mRNA (blue, a
neurectodermally expressed gene). For a visual representation of the ISH technique see the appendix.
Figure 2. In situ hybridization on a Drosophila embryo using two probes to detect snail (brown) mRNA and rhomboid (blue) mRNA.
Procedure
Week 1, Day 1:
A. Preparing the embryos for hybridization The embryos have already been dissected, fixed in formaldehyde and stored in PBT (phosphate-buffered saline
with 0.1% Tween-20 detergent) but the in situ hybridization buffer (ISHB) is very different. It contains
formamide and we have to gradually assimilate the embryos to the buffer.
1. Label your tubes of embryos on the top and side with your group name/initials. Everyone will perform an
ISH on 2-6 hr w1118 embryos.
2. Carefully drain your embryos by removing the buffer they are in with a plastic pipet. PLEASE NOTE:
EMBRYOS SETTLE QUITE NICELY TO THE BOTTOM OF THE TUBE, BUT ARE EASILY
DISTURBED. If you disturb them, allow them to settle again. Try to remove as much liquid as possible
without removing any embryos. You will probably need to leave about 50 µl of liquid above the embryos in the
tube.
3. You will add different mixtures of PBT and ISHB, rock the embryos gently on the nutator for 3 minutes, and
then change to the next mixture by draining and adding. The idea is to eventually get the embryos into 100%
ISHB. Be sure to give the embryos enough time to settle in between washes to ensure that you do not throw
them away into the waste. *** As you progress through the washes, be aware that the embryos change from
opaque white to translucent in appearance and settle more slowly as the percentage of ISHB increases.
1) 25% ISHB, 75% PBT (750 µl)
2) 50% ISHB, 50% PBT (750 µl)
3) 75% ISHB, 25% PBT (750 µl)
4) 100% ISHB (750 µl)
5) 100% ISHB + carrier (750 µl, leave in this solution for the prehybridization)
B. Prehybridization.
Prehybridization is an incubation to allow the blocking agents in ISHB to bind to any sites on the tissue that our
probe might stick to non-specifically. These agents include non-specific DNA and RNA (carrier), heparin, and
BSA. All of these are either “sticky” or charged molecules that should help prevent background staining.
1. Incubate your embryos in the 55°C water bath for 30 minutes.
2. In the meantime, prepare your probe by heating it to 80°C for 5 minutes and then placing it immediately on
ice until the next step. The probe is labeled antisense RNA, made by in vitro transcription from the class
recombinant eve or twist plasmids, which has been diluted in ISHB. This treatment removes any secondary
structure (internal basepairing) in the probe, assuring that it will be single stranded.
C. Hybridization.
1. Remove your tube to the rack at your bench and allow the embryos to settle.
2. Carefully drain your embryos as before and then add 300 µl of the probe to each sample (1:100 sim or sna
antisense RNA in 100% ISHB+carrier).
3. Gently tap the tube to mix the contents and place the tube back into the 55°C water bath overnight or up to a
few days.
Day 2-4:
D. Washing the hybridized samples.
At some point during 1-4 days after you have started the hybridization, come in to wash your hybridizing
samples. We have to reverse the process to get them back into 100% PBT. This should take you about 1 1/2
hours.
1. Remove your tube from the 55°C water bath and drain the embryos. Remember to give your embryos time
to settle properly.
2. Add 750 µl of fresh 100% ISHB and rock the tube for 5 minutes. Then drain the embryos again.
3. Repeat step 2 twice more (3 washes total).
4. Add different mixtures of PBT and ISHB, rock the embryos gently on the nutator for 3 minutes unless
otherwise indicated, and then change to the next mixture. The idea is to slowly get the embryos back into
100% PBT.
1) 75% ISHB, 25% PBT (750 µl)
2) 50% ISHB, 50% PBT (750 µl)
3) 25% ISHB, 75% PBT (750 µl)
4) 100% PBT (1 ml) 3 times without rocking, just let the embryos settle
5) 100% PBT (1 ml) 3 times, rock for 5 minutes on the nutator
6) 100% PBT (1 ml)
E. Detecting the probe with a sheep anti-digoxigenin antibody coupled to alkaline phosphatase.
To detect the hybridized RNA we will use an antibody that can bind to the label on the RNA probe. The probe
is labeled with digoxigenin and we want to detect this. Roche Biochemicals has developed an antibody that will
recognize digoxigenin and they have covalently linked this antibody to the enzyme alkaline phosphatase (α-dig
AP antibody).
1. Drain your embryos once more and add 500 µl of antibody, labeled α-dig AP antibody, (which has been
diluted 1:1000 in PBT with a 2% BSA and stored in the fridge) to each sample. Gently invert the tube and
then place it in the labeled rack back in the refrigerator. We will do the last step the following week and
observe the staining.
[End of Week 1 of ISH]
Week 2, Day 8:
E. Staining the embryos.
We are now set to stain.
1. Drain your embryos and wash them six times with 1 ml of PBT, gently rocking on the nutator for 3 minutes
each time.
2. Drain the embryos once more and add 1 ml of Staining Buffer (SB). This is the buffer without the
substrates (just to get the embryos in the proper conditions for the enzyme to work). Set up the scope: take
out a dissecting microscope and look at your embryos by angling the tube under the microscope initially.
You want to be able to focus close up on the embryos as you are rotating the tube around. This will allow
you to see the embryos as clearly as possible, without having to look through the solution which tends to
distort the view.
3. Drain the embryos yet again and add 500 µl of Alkaline Phosphatase Staining Buffer (APSB, staining buffer
with the substrates) to each sample. View the embryos initially and then every few minutes once the stain is
added. We’ll decide together when to stop the staining, but essentially when a decent amount of purple
staining is seen through your collection of embryos the color development should be stopped. If there is no
staining by about 30 minutes something has gone wrong, did you really add the APSB solution???
F. Stopping the staining and preparing a slide.
1. Once the reaction has preceded far enough, stop it by washing 5 times with 1 ml of PBT (no incubation
required, just mix and allow embryos to settle).
2. After the last wash, drain the embryos and add 300 µl of 70% glycerol to each sample. Using a transfer
pipette, gently pipet up and down (don’t introduce air bubbles) to resuspend the embryos. Pick up as many
embryos as you can (don’t worry about getting them all – you can always prepare another slide) and place
them in a drop on a microscope slide.
3. Try to remove as much excess 70% glycerol from your drop as you can (use a P200 with tip or the edge of a
Kimwipe to soak up the excess) without removing the embryos and then place a coverslip on top of the
slide. Try to avoid squishing your embryos. Observe your stained embryos with a compound microscope
and take pictures of representative stages with the camera through the ocular. Record the numbers of the
pictures you took and provide me with this information along with whether they were stained with the eve
or the twist probe.
Appendix:
Principle of in situ hybridization
1. A cell with nucleus, showing a single mRNA from the gene of interest. Although we show only one
mRNA molecule, there are many copies of mRNA from this one gene in the cell and they represent a small
fraction of the total mRNA.
2. Hybridization of antisense RNA probe labeled with digoxigenin. The antisense RNA is capable of basepairing with the mRNA of interest (and not with mRNAs from other genes). After the hybridization,
unbound antisense RNA probe is washed off. The hybridized probe is bound so strongly, it is resistant to
washing.
3. Recognition of antisense RNA probe by an anti-digoxigenin antibody. The antibody has been designed
to specifically bind to digoxigenin, which was incorporated into our antisense RNA probe. The enzyme
alkaline phosphatase (AP) was chemically attached to the antibody.
4. Detection of bound anti-dig antibody coupled to alkaline phosphatase. To visualize our mRNA, we add
BCIP, a substrate for AP, and NBT (which enhances the color). Both of these are soluble but when AP
cleaves the phosphate group off of BCIP, the resulting molecule turns blue and precipitates. This stains the
local area (in this case, the cytoplasm) blueish purple.
5. An example of six large cells within a tissue that have been detected by ISH. Note that the center of
each cell is not stained (the large nucleus) whereas the surrounding cytoplasm is blueish purple. This is a
much higher magnification that we will be looking at. Under lower levels of magnification, you may or
may not be able to make out individual cells, but you will be able to see patterns.