IN SITU HYBRIDIZATION
OF NON-RADIOLABELED RNA PROBES TO SECTIONED TISSUE.
In situ hybridization is a useful technique for
determining spatial patterns of gene expression. In this lab, we will carry out hybridizations of digoxigenin
labeled probes to plant material that has been fixed, embedded in paraffin,
sectioned and placed on microscope slides.
We don’t have enough time to carry out the entire protocol - it takes a
week just to fix and embed tissue - so we will perform only the actual
hybridization. You will be provided
with slides that have sectioned siliques on them. The siliques, and the embryos within them, are at various stages
of development. You will also be
provided with probes for transcripts that are localized to specific subdomains
of the developing embryo. (Protocols
for tissue fixation, embedding, sectioning and probe synthesis are included for
your general information. These follow
the protocols for the hybridization procedure.)
The hybridization
experiment takes two days. On the first
day, the tissue sections are subjected to a variety of pretreatments and the
hybridizations are set up. On the
second day, the tissue sections are washed and the probe is visualized. The digoxigenin labeled hybrids are detected
by applying anti-digoxigenin antibodies to the tissues. This antibody is conjugated to alkaline
phosphatase. Alkaline phosphatase
carries out a reaction that yields a colored (blue in this case)
precipitate. The color develops over
the course of one to three days.
This protocol is the
one used in the Barton lab. It has been
modified by Jeff Long from protocols used in Sarah Hake’s, Vivian Irish’s and
Elliot Meyerowitz’s labs. Useful
references for in situ hybridization to plant tissues are:
Jackson, D. (1991) In-situ
hybridization in plants.
in Molecular Plant Pathology: A Practical
Approach Eds. Bowles, D.J., Gurr,
S.J and McPherson, M. Oxford University
Press.
In situ hybridization : a practical approach (1992), edited by D.G. Wilkinson, Oxford
University Press.
I. TISSUE FIXATION AND EMBEDDING
This protocol uses a
paraformaldehyde fixative. FAA (50%
ethanol, 5% acetic acid, 3.7% formaldehyde) is an alternative fixative. (If FAA is used, ethanol series for
dehydration begins at 50%.)
Day 1- fix tissue
Make up the amount of
1X PBS you will need and adjust the pH to 11 with NaOH. Heat the solution to 60 to 70°C. Add paraformaldehyde. Do this in a fume hood. The paraformaldehyde should dissolve within
a few minutes. Place solution on ice
and when it has cooled to about 4°C, adjust the pH with H2SO4.
Dispense the
paraformaldehyde solution into glass scintillation vials. Place vials on ice in a vacuum
desiccator. Add freshly harvested
tissue to the vials. Close the
desiccator and apply a vacuum to the samples until the paraformaldehyde
solution starts to bubble. Hold the
vacuum for 15 minutes and then release it slowly. Repeat this procedure until the tissue sinks. Replace the paraformaldehyde solution with
fresh paraformaldehyde solution and shake gently overnight at 4°C.
Day 2 - dehydrate tissue
All steps are done at
4°C with gentle shaking or rotating. A
Labquake rotator works well.
|
Solution |
time |
number
of changes |
|
1 X PBS |
30 minutes |
two |
|
30% ethanol |
60 minutes |
one |
|
40% ethanol |
60 minutes |
one |
|
50% ethanol |
60 minutes |
one |
|
60% ethanol |
60 minutes |
one |
|
**70% ethanol |
60 minutes |
one |
|
85% ethanol |
60 minutes |
one |
|
95% ethanol + eosin* |
overnight |
one |
*The amount of eosin
added is not exact. Add enough to give
the ethanol a nice bright orange color.
The eosin is added to dye the tissue so that it can be easily seen when
embedded in paraffin.
**Tissue can be stored
in 70% ethanol at 4°C for at least several months.
Day 3 - Finish dehydration and begin embedding
All steps are done at
room temperature with gentle shaking or rotating.
|
Solution |
time |
number
of changes |
|
100% ethanol + eosin |
30 minutes |
two |
|
100% ethanol + eosin |
60 minutes |
two |
|
25% Histoclear, 75% ethanol |
60 minutes |
one |
|
50% Histoclear, 50% ethanol |
60 minutes |
one |
|
75% Histoclear, 25% ethanol |
60 minutes |
one |
|
100% Histoclear |
60 minutes |
two |
|
100% Histoclear + 1/4 volume Paraplast Plus
(paraffin) chips |
overnight (no shaking) |
|
Day 4 - continue embedding
Place the vials at 42°C
until the Paraplast chips melt completely.
Add another 1/4 volume
of chips and wait until completely melted.
Move vials to 60°C.
After several hours,
replace the wax/Histoclear solution with freshly melted wax. Leave overnight.
Day 5 - more embedding
Replace old wax with
fresh wax twice.
Day 6 - more embedding
Replace old wax with
fresh wax twice.
Day 7 - more embedding
Replace old wax with
fresh wax twice.
Day 8 - place tissue in molds
Place tissue in
molds. We use aluminum weigh boats
(Fisher Scientific). Be careful to
arrange tissue in the weigh boats so that it will be easy to cut the wax block
into appropriate sized pieces and so that it will be easy to orient the tissue
for sectioning. Store paraffin blocks
with embedded tissue at 4°C.
II. SECTIONING
Use ProbeOn Plus slides
(Fisher Scientific). They are
pre-cleaned, charged, and easily sandwiched together for later steps.
Place slide on a slide
warmer that has been pre-warmed to 42°C.
Apply several drops of DEPC-treated water to the slide.
Cut six to eight
micrometer thick sections on a microtome.
Float the ribbons of tissues on top of the water droplet on the
slide. Within a few minutes, the
ribbons should flatten out. Paint
brushes can be used to manipulate the ribbons.
Wick off excess water
from the slide with a Kimwipe. Excess
water can cause the tissue to bubble.
Incubate the slides on
the slide warmer overnight so that tissue adheres tightly to the slide. Slides can be stored with desiccant for
several weeks at 4°C.
III. PROBE SYNTHESIS
Transcription
Template: Transcripts are made from plasmids carrying
promoters for T3, T7 or SP6 polymerases.
The plasmid is linearized first.
Be sure to digest to completion.
Do not use enzymes that leave a 3’ overhang. Phenol/chloroform extract to get rid of any RNAses. Resuspend the DNA in DEPC-treated water to a
concentration of 0.5 micrograms/microliter.
Transcription
Reaction:
|
template DNA
(0.5micrograms/microliter) |
4 microliters |
|
5X transcription
buffer (Stratagene) |
5 microliters |
|
5X nucleotides (2.5 mM each; UTP is 1/2 “cold” UTP
and 1/2 dig-UTP
(Boehringer, cat#1209256) |
5 microliters |
|
RNASIN (Promega) (RNAse inhibitor) |
to 1 unit/microliter |
|
RNA polymerase |
to 0.4
units/microliter |
|
water |
to total |
|
Total Volume |
25 microliters |
Incubate the reaction
at 37°C for 30 to 60 minutes.
To check that you have
synthesized transcripts, run 1 microliter of transcription reaction on a
minigel (add ethidium bromide to gel) at about 100 V for about 15 minutes. The
RNA degrades quickly so do not run it for too long. Treat the gel box and comb with 0.2 N NaOH for 30 minutes before
use. Probe will appear as a band below
the template band. Use the relative
intensity of the two bands to estimate the amount of probe synthesized. If, for instance, the transcript band and
the template band are of equal intensity, estimate that you have synthesized 2
micrograms of RNA.
Add 75 microliters
water and 1 microliter of 100mg/ml tRNA and 5 units of RNAse-free DNAse to
transcription reaction. Incubate for 10
minutes at 37°C.
Precipitate RNA probe
by adding an equal volume of 4M NH4OAc and 2 volumes of 100%
ethanol. Place at -20°C for twenty
minutes. Centrifuge in a microfuge to bring down the precipitated RNA. Rinse the pellet with 70% ethanol. Air dry pellet or place tube in speed vac with
no heat.
Carbonate hydrolysis
The probe is hydrolyzed
into fragments between 75 and 150 base pairs long. To calculate the length of time to let the reaction go, use the
following formula:
T (time) = (Li-Lf)/K
Li Lf
where Li is
the initial length of probe, Lf is the final length of probe and
K=0.11 kb/minute.
Resuspend the RNA
pellet in 100 microliters water. Add
100 microliters 2X carbonate buffer (80 mM NaHCO3, 120 mM Na2CO3). Incubate at 60°C for the time
calculated. Neutralize with 10 microliters
10% acetic acid.
Precipitate with 1/10
volume 3 M NaOAc (pH 5.2) and 2 volumes ethanol. Place tube at -20°C for twenty minutes, centrifuge and rinse
pellet with 70% ethanol. Resuspend in
50% formamide at a concentration of 1 microliter/slide (see below).
The probe is used at a
final concentration of 0.5 ng/ microliter/ kb probe complexity. Hybridizations are done in 100 microliters
per slide. So if your probe is 1 kb,
you will need 0.5 ng x 100 microliters x 1 kb = 50 ng of probe per slide. If your probe is 2 kb, you will need 100 ng
of probe per slide, etc.
Experiment with
different amounts of probe (try up to 5X higher and lower) to find the
concentration that gives the best signal.
DAY ONE
A. Section
Pretreatment
The paraffin is removed
from the sections with Histoclear (National Diagnostics) or Citra-Solv (Fisher)
and the sections are rehydrated through an ethanol series.
(A note about
solutions: All solutions should be made RNAse
free. Most of the solutions are not
diethylpyrocarbonate (DEPC)-treated but everything is autoclaved before
use. Plastic containers are treated
with 0.1 M NaOH overnight and rinsed with sterile water. Be careful to handle all tissue and
solutions with gloves to avoid contamination with RNAses.)
1. Deparaffinize and rehydrate sections. All steps are at room temperature unless
otherwise noted. Place slides in rack. Move slide rack through the following
solutions:
|
Solution |
time |
number
of changes |
|
100% Histoclear |
10 minutes |
two |
|
100% ethanol |
1-2 minutes |
two |
|
95% ethanol |
1-2 minutes |
one |
|
90% ethanol |
1-2 minutes |
one |
|
80% ethanol |
1-2 minutes |
one |
|
60 % ethanol |
1-2 minutes |
one |
|
30 % ethanol |
1-2 minutes |
one |
|
water |
1-2 minutes |
one |
|
2XSSC |
15-20 minutes |
one |
2. Protease.
Tissue sections are proteased to increase the accessibility of the
target RNA. Treat sections with
proteinase K (1 microgram/ml in 100 mM Tris (pH 8) and 50mM EDTA) at 37°C for
30 minutes. To make protease solution,
prewarm Tris/EDTA solution to 37°C;
then add the proteinase K (from a 10mg/ml frozen stock) immediately
before adding slides.
3. “Refix” tissue. This prevents disintegration of the tissue following protease
treatment.
|
Solution |
time |
number
of changes |
|
2mg/ml glycine in 1 X PBS |
2 minutes |
one |
|
1 X PBS |
2 minutes |
two |
|
4% paraformaldehyde, pH 7 (made fresh*) |
10 minutes |
one |
|
1 X PBS |
5 minutes |
two |
*See section on
fixation for paraformaldehyde recipe.
4. Acetic anhydride treatment. In this step, positively charged amino groups
are acetylated. This reduces
non-specific binding of the probe.
Place slides in a
solution of 0.1 M triethanolamine (pH 8, made fresh) and acetic anhydride for
10 minutes with stirring. (To do this,
elevate the slide rack above a stir bar).
Dispense the acetic anhydride (4 mls for 800 mls of triethanolamine)
into the triethanolamine and mix for a few minutes before putting the slide
rack in.
5. Wash and dehydrate.
|
solution |
time |
number
of changes |
|
1X PBS |
5 minutes |
two |
|
30% ethanol |
30 seconds |
one |
|
60% ethanol |
30 seconds |
one |
|
80% ethanol |
30 seconds |
one |
|
90% ethanol |
30 seconds |
one |
|
95% ethanol |
30 seconds |
one |
|
100% ethanol |
30 seconds |
two |
Store slides in an
airtight container with a small amount of 100% ethanol in the bottom for up to
several hours at 4°C.
B.
Hybridization.
Organize the slides to
be probed. Slides will be sandwiched
together as pairs. Probe is applied to
each individual pair. Decide which pairs
will be probed with sense strand probes (controls) and which will be probed with
antisense strand probes.
1. Make hybridization
solution (enough for 5 slide pairs):
100 microliters 10X in
situ salts
400 microliters
deionized formamide
200 microliters 50%
dextran sulfate
20 microliters 50 X
Denhardt’s
10 microliters 100
mg/ml tRNA
70 microliters
DEPC-treated water
total volume: 800
microliters
The hybridization
solution is very viscous from the dextran sulfate. It can be warmed before using and/or you can make more than you
need to make up for losses in dispensing.
Air dry the slides on a
clean paper towel. They must be
completely dry.
2. For each pair of slides, the desired amount of probe (see section
on probe synthesis for suggested amount) should be added to 50% formamide so
that the volume is 40 microliters. Heat
this solution to 80°C for 2 minutes.
Ice, spin down and keep on ice.
Add 160 microliters of hybridization solution to the 40 microliters of
probe solution. Mix without generating
air bubbles.
3. Apply probe/hybridization solution mixture
to slides. 200 microliters of the
probe/hyb solution needs to be added to each slide pair. One technique is to apply 100 microliters to
each slide, spreading it out over the entire slide with the side of a pipette
tip so that all of the tissue is exposed to probe. Then slowly sandwich the slides together, avoiding air
bubbles. A second technique is to put
all of the probe in the middle of one slide and slowly massage the other slide
on top until the two slides are together.
Elevate the slide
sandwiches above wet paper towels in an airtight plastic
container. Plastic serological pipettes
can be used to raise the slides above the paper towels. Place the container at 50 to 55°C overnight.
DAY TWO
A. Wash
slides
Warm 0.2X SSC to 55°C
and NTE to 37°C.
Dip pairs of slides
into prewarmed 0.2X SSC to separate and rinse them before placing in slide
rack. Carry out all washes with gentle
agitation.
|
solution |
time |
temp |
number
of changes |
|
0.2X SSC |
60 minutes |
55°C |
two |
|
NTE |
5 minutes |
37°C |
two |
|
RNAse (20 micrograms/ml in NTE) |
30 minutes |
37° |
one |
|
NTE |
5 minutes |
37°C |
two |
|
0.2X SSC |
60 minutes |
55° |
one |
|
1 X PBS |
5 minutes |
room temperature |
one |
B. Block and apply antibody
1. Place slides face up on bottom of plastic
container. Wash 45 minutes with 1%
Boehringer block (cat. #1096176) (made fresh following manufacturers
directions) in 100 mM Tris, pH 7.5, 150 mM NaCl. Use just enough of the solution to cover the slides and place on
a rocking platform at room temperature.
2. Replace block solution with 1% BSA in 100 mM
Tris (pH7.5), 150 mM NaCl, 0.3% Triton X-100 for 45 minutes.
3. Dilute the anti-digoxigenin antibody
(Boehringer, cat#1093274) 1 to 1250 in the BSA/Tris/Triton solution from step
2. Make a puddle of antibody solution
in a plastic weigh boat. Make slide
“sandwiches” (two slides, tissue sides together). Dip the long edge of the sandwich in the antibody solution and
allow capillary action to pull the solution up into the gap between the slides. Drain the solution onto a Kimwipe and
repeat, avoiding air bubbles.
Place a layer of wet
paper towels in an airtight plastic container.
Elevate the slides above the towels (for instance on a few disposable
serological pipettes) and allow to sit at room temperature for 2 hours.
C. Wash
slides
1. Drain the slides on Kimwipes and
separate. Place the slides face up in a
single layer on the bottom of a plastic container. Wash 4 times at room temperature, 15 minutes each time, with the
BSA/Tris/NaCl/Triton solution.
2. Wash once for ten minutes with 100mM Tris
(pH 9.5), 100 mM NaCl, 50 mM MgCl2.
Then dip each slide in fresh Tris 9.5/NaCl/MgCl2 solution to ensure that
all of the detergent is washed off.
D. Apply
substrate
Pour 10 mls of the
NBT/BCIP premix (Western Blue from Promega, cat #S3841) into a plastic weigh
boat. Sandwich slides together, place
edge of sandwich in solution and allow capillary action to draw the solution up
between the slides. Drain on a Kimwipe
and repeat. Place the slides in an
airtight plastic container, elevated above wet paper towels. Place the container in total darkness for
one to three days.
DAY 3, 4 OR 5
Take down and mount slides
1. You should peek at the slides periodically
to monitor the color reaction. When
sufficient signal has developed, drain the solution onto a Kimwipe. Lower the slides into a bath of TE to
separate them and to stop the reaction.
2. Dehydrate the slides - in this ethanol
series it is important to keep the time in ethanol to a minimum (less than five
seconds per solution) since the color product is soluble in ethanol.
|
solution |
number
of changes |
|
30% ethanol |
one |
|
50% ethanol |
one |
|
70% ethanol |
one |
|
85% ethanol |
one |
|
95% ethanol |
one |
|
100% ethanol |
two |
|
100% Histoclear |
two |
3. Drain and mount the slides in Cytoseal 60
(Stephens Scientific).
Recommended Stock Solutions for in situ hybridization
|
Solution |
Composition |
Remarks |
|
10XPBS, pH 7 |
1.3 M NaCl 70 mM Na2HPO4 30 mM NaH2PO4 |
|