Lab Protocols
queries to Nigel Pringle
1. In situ hybridization with 35S-labelled RNA probes
All solutions and equipment need to be RNase free. This involves either baking glassware at 250oC for 24 hours or treating solutions and plastic containers with diethylpyrocarbonate (DEPC). (Add 0.5ml of DEPC to 1000ml of solution, shake vigorously and autoclave.) In the following protocol it is assumed that all solutions and equipment used will be RNase free, unless stated otherwise.
Slide preparation
1. Scratch a numbering system on to each glass microscope slide using a diamond scriber. This will help later when you are in the darkroom, trying to figure out which side the slides are on.
2. Place slides in a metal rack and soak in hot soapy water for an hour (fairy liquid is OK).
3. Wash thoroughly under running tap water for an hour.
4. Soak slides in distilled water (DH2O) for 10 minutes, repeat this a further 2x before baking slides at 250oC for 24 hours.
5. Coat slides in 2% APES (3-Aminopropyltriethoxysilane, from Sigma) diluted in industrial methylated spirits (IMS) for 30 seconds. This coating helps cryostat sections adhere firmly to the slides during the in situ procedure.
6. Wash in 9% IMS for 30 seconds.
7. Wash in DH2O for 30seconds.
8. Dry slides and store in a dust free container. These slides will keep for months.
Tissue fixation
1. Place fresh tissue in 4% paraformaldehyde in PBS and leave at 4oC for 24 hours. PBS:- 8.0g NaCl, 0.2g KCl, 0.1g CaCl2, 0.1g MgCl2, 1.15g Na2HPO4, 0.25g KH2PO4. Make up to 1 litre with DH2O, pH7.0
2. Place tissue in 0.5M sucrose in PBS and leave at 4oC for a further 24 hours.
3. Embed tissues in OCT embedding compound (BDH) in aluminium foil containers and freeze slowly on dry ice. Tissue blocks can be stored at -70oC until required.
4. Cut frozen sections (10mm nominal thickness) and collect on APES-coated slides.
5. Dry slides (with sections) briefly on a hotplate at 50oC for 2 minutes. Then air-dry in a dust-free environment for up to 2 hours.
6. Fix sections in 4% paraformaldehyde in PBS for 15 minutes.
7. Wash in PBS for 5 minutes. At this point you can a) dehydrate the sections through ascending concentrations of ethanol (30, 60, 80, 95, 100%, 1 min each), dry and store at -70oC for at least 6 months (rehydrate before use), or b) continue with the in situ hybridisation procedure.
In situ hybridization
1. Take slides from PBS, wash and place in proteinase K buffer for 5 minutes., following fixation; or if they have been stored at -70oC rehydrate and place in Proteinase K buffer for 5 minutes. Proteinase K buffer: 50 mM Tris-HCl 5 mM EDTA pH 7.5
2. Incubate for 30 minutes at 37oC with RNase free proteinase K at1mg/ml in proteinase K buffer. The digestion time might have to be adjusted for different tissues.
3. Wash in 0.2% (w/v) glycine in PBS for 30 seconds to stop the digest.
4. Wash 2 x 30 seconds in PBS.
5. Fix in 4% paraformaldehyde in PBS for 15 minutes.
6. Wash in PBS for 2 minutes.
7. Place slides in 0.1M triethanolamine (pH 8) for 5 minutes.
8. Replace with 0.1M triethanolamine (pH 8.0) containing freshly added acetic anhydride (1/400 dilution; add the acetic anhydride to the triethanolamine, shake vigorously for a few seconds and use immediately). Incubate for 10 minutes, then repeat. This acetylation step is very important for reducing nonspecific binding of the probe.
9. Wash in PBS for 3 minutes.
10. Dehydrate through ascending ethanol concentrations (30, 60, 80, 95, 100%, one minute in each). Air dry thoroughly.
11. Incubate the sections with radioactive probes overnight at 55oC. The 35S-labelled probes are usually kept as a 10 x stock at 1ng/ml per 1kb probe length (e.g. 1.5ng/ml for a 1.5kb probe). Heat the probes at 80oC for 5 minutes in hybridization buffer, chill briefly on ice and add an appropriate volume (20ml for a 22 x 22mm coverlip, 100ml for a 22 x 60mm coverslip over each section. Cover the sections with a glass coverslip, taking care not to trap bubbles over the sections. Place in a humid container* and incubate at 55oC overnight. * We use a lunch box with wet filter papers in the bottom; it is important to include 50% formamide in the wetting solution to prevent evaporation of formamide from the hybridization solution from around the edges of the coverslip. Also, make sure that the slides are absolutely level in order to get uniform hybridization signals.
Hybridization buffer:- 0.3M NaCl, 10mM Tris-HCl pH6.8, 5mM EDTA, 10% (w/v) dextran sulphate, 0.1mg/ml yeast tRNA, 1 x Denhardt's, 10mM dithiothreitol (DTT) 50% (v/v) formamide. We have found that there is no need to deionize the formamide as long as the bottle is recently opened. However, if you prefer, deionize the formamide by stirring 10g Amberlite monobed resin MB-1 (BDH) with 100ml of formamide for 30minutes at room temperature. Remove the resin beads and store at -20oC.
100 X Denhardts: 2% (w/v) Ficoll 400, 2% (w/v) polyvinyl pyrrolidone (PVP), 2% (w/v)BSA (Sigma fraction V) Filter through a 0.45mm filter and store at -20oC.
12. * For the following washing steps, solutions and containers do not have to be RNase free.
To remove coverslips wash in 4 x SSC at room temp (usually in glass coplin jars) until the coverslips have come off. Incubate in 4 x SSC at room temperature for a further hour. This step is considered very important for low backgrounds on sections (see Du Pont Biotech Update on in situ hybridisation; vol 4, No.5, 1989).
20 X SSC: 3M NaCl 0.3M Na3Citrate pH7.5
13. Incubate in wash buffer at 65oC for 30 minutes. This wash is at a temperature close to the Tm of the RNA/RNA hybrids. As a rough working guide, if you are using homologous probes - e.g. rat RNA probes on rat sections - this is ~65oC. However for non-homologous probes - e.g. human RNA probes on rat sections - the temperature should be lower, say around 55oC.
Wash buffer: 0.3M NaCl, 10mM Tris-HCl pH6.8, 5mM EDTA, 10mM DTT, 50% (v/v) formamide (see above, step 11).
14. Incubate in RNase A buffer 2 x 5 minutes: 0.5M NaCl, 10mM Tris-HCl, 0.1mM EDTA pH7.5
15. Incubate in RNase A buffer at 37oC containing RNAse A at 20mg/ml for 30 minutes.
16. Incubate in RNase A buffer for a further 15 minutes at room temperature.
17. Incubate in wash buffer at 65oC for a further 30 minutes.
18. Incubate in 2 x SSC at 45oC for 30 minutes.
19. Incubate in 0.1 x SSC at 45oC for 30 minutes.
20. Dehydrate through ascending ethanols (30, 60, 80, 95%, 100%; 1 minute each) and air dry. The slides are now ready to be coated with photographic emulsion.
Autoradiography
* It is not necessary to use RNase free reagents for autoradiography.
1. Work under an appropriate safelight (e.g. Ilford 902S filter). Melt ~5ml of Ilford K5 emulsion in a narrow slide holder at 40-43oC. When this has melted add 5mls of DH2O containing 100ml of glycerol. Mix gently by inverting the tube. Do not shake as this will make the emulsion mixture froth. I usually use what are called "slide postal boxes", available from A.R.Horwell. These are convenient as they only use small quantities of emulsion; 5ml of emulsion + 5ml of H2O is usually sufficient to coat over 25 slides.
2. Dip a test slide, remove and wipe excess emulsion from the back of the slides. Hold up to safelight and check there are no bubbles. If there are, continue dipping until they have been removed. Now dip your processed slides with sections. Be careful to wipe the emulsion from the side that does not have the section on it! (Use scratched numbering on slides to determine the right side).
3. Allow the emulsion to set by placing the slides face up on foil covered glass plates on ice for 10 minutes.
4. Remove from ice, lay the slides flat and allow the emulsion to dry in complete darkness. This usually takes around an hour.
5. Place slides in a slide rack inside a light-proof box containing some silica gel, and store at 4oC until ready to develop.
Developing slides
Slides should allowed to expose in the dark overnight or for up to several weeks, depending on the strength of the signal. Often 1 week is good.
1. Remove slides from 4oC and allow to warm to room temperature for at least an hour before opening the box; this prevents condensation from forming on the photographic emulsion. The developing, stop and fix must be done in a darkroom under a safelight.
2. Develop in Ilford D19 developer for temperature and developing times are critical!! 23oC: 1 minute 20 seconds 22oC: 1 minute 40 seconds 21oC: 1 minute 50 seconds 20oC: 2 minutes 19oC: 2 minutes 15 seconds 18oC: 2 minutes 30 seconds 17oC: 2 minutes 40 seconds 16oC: 3 minutes
3. Stop the developing with 1% acetic acid for 1 minute.
4. Fix in 30% (w/v) sodium thiosulphate for 5 minutes.
5. Wash in distilled water for at least 20 minutes at room temperature. This can be done with the light on.
6. Staining sections. I use hematoxylin (Gill's no.3, Sigma) diluted 1:1 with H2O for 10 seconds at room temperature. Filter the stain (0.45mm) and discard afterwards. Differentiate the stain under running tap water for a few minutes. Beware of over-staining, which can mask the in situ hybridization signal.
7. Dehydrate through ascending ethanols (30, 60, 80, 95, 100%; 1 minute each).
8. Clear in xylene 2 x 1 minute and mount under a xylene based mountant such as Xam or Depex. Slides can now be viewed under bright field and dark field illumination.
Preparation of 35S-labelled RNA probes
Several points are important. 1) if you are making your own ribonucleotide stocks make sure they have been neutralized to pH7.0. It is easier to buy rNTPS ready-prepared in solution. 2) Add the components of the reaction in the order shown. 3) The mixture should be kept at room temperature during the addition of the components, since DNA can precipitate in the cold in the presence of spermidine.
This protocol produces high activity probes (usually ~2 x 108 DPM/mg of RNA) and 100-200ng RNA.
1. Assemble in vitro transcription reaction: 4.0ml of 5 x transcription buffer (200mM Tris-HCl pH7.6, 30mM MgCl2, 10mM spermidine, 50mM NaCl), 2.0ml of 100mM DTT, 20 units RNasin ribonuclease inhibitor, 4.0ml each of 2.5mM ATP, GTP and CTP (prepare just before use by mixing 1 volume of DH2O with 1 volume each of 10mM ATP, GTP and CTP stock solutions), 2.4ml of 100mM UTP, 1.0ml linear template DNA (in DH2O or TE at 1mg/ml), 5.0ml alpha-35S-UTP (50mCi at 10 mCi/ml), 1.0ml SP6 (or T7 or T3) polymerase at 15-20 units/ml. Bring to final volume of 20ml with H2O. Incubate at 37oC for 60 mins.
2. Destroy the DNA template. Add 20 units RNasin, 1.0ml tRNA (25mg/ml) 1.0 units DNase1 (RNase free). Incubate at 37oC for 15 minutes.
3. Add 200ml of 10mM DTT.*At this point remove 0.5ml and place in 10ml of Northern sample buffer (see section on checking full length transcripts). * Also remove a further 0.5ml and add to 400ml of 0.5M NaH2PO4 to assess 35S-UTP incorporation(see below).
4. Extract sample twice with equal volumes of phenol and chloroform.
5. Add 1/10 volume of 3M NaAcetate to the aqueous phase and precipitate the RNA by addition of 2.5 volumes of 95% ethanol at -20oC (keep on dry ice 10 minutes). Remove the supernatant and dry the pellet before dissolving in 50ml of 10mM DTT.
Alkaline hydrolysis of RNA probe
For optimal penetration of tissue sections, the 35S-labelled RNA probe requires digesting to around 100-150bp fragments. This is achieved by limited alkaline hydrolysis at 60oC for X minutes.
where X = (Lo-Lf)/KLoLf
where Lo= original transcript length in kb. Lf= Final transcript length in kb (0.1-0.15) K = 0.11
1. To the 50ml of RNA probe add 50ml of 100mM carbonate buffer pH10.2 (100mM NaHCO3 and 100mM Na2CO3: titrate against each other to pH10.2 Store stock at -20oC).
2. Incubate at 60oC for X minutes.
3. Neutralize with 100ml of neutralising buffer: 0.2mM NaAcetate, 1.0% (v/v) glacial acetic acid, 10mM DTT
* Remove a 0.5ml aliquot and place in Northern sample buffer (see section on checking digested transcripts).
4. Add 1/10 volume of NaAcetate and precipitate with 2.5 volumes of 95% ethanol as described above.
5. Dissolve pellet at 1ng/ml/kb probe complexity in 10mM DTT, 50% deioninised formamide. This is 10X stock. The recovery in terms of ng RNA probe can be determined as follows:
35S incorporation efficiency
1. Take 6 Whatman GF/C x 2.3cm diameter glass microfibre filters and add 50ml of 0.5M NaH2PO4 containing 0.5ml freshly made probe.
2. Wash three of these filters in 0.5 litres of 0.5M NaH2PO4 for a few minutes, to remove unincorporated probe. Dry all 6 filters and count in a scintillation counter. The % incorporation of 35S-UTP can be calculated from the total counts added (unwashed filters)and the incorporated counts (washed filters). From this the recovery can be assessed - roughly 4ng RNA per 1% 35S-UTP incorporated.
Check the lengths of the full-length and alkaline-hydrolyzed transcripts by Northern blot.
References
Du Pont Biotech Update on in situ hybridisation, (1989), Vol 4, No.5.
Lawrence and Singer, Quantitative analysis of in situ hybridization methods for the detection of actin gene expression, (1985), Nucleic Acids Res, 13, 1777-1799.
Maniatis et al., (1989) Molecular cloning: A laboratory Manual. 2nd edition. Cold Spring Harbour Laboratory Press, Cold Spring Harbour, NY.
2. Non-radioactive in situ hybridization with digoxygenin (DIG)-labelled probes.
1. Preparation of tissue sections
Tissue is prepared, sectioned and the sections collected as described above (ISH using radiolabelled probes). However, after air drying at room temperature for 2 hours, the sections are not treated further before hybridization.
2. Preparation of digoxygenin-labelled probes
Template DNA is prepared as described above (ISH with radiolabelled probes). In vitro transcription reactions are set up at room temperature in the following order:- 2.5ml linear template DNA, 4.0ml 5 x transcription buffer (as above), 6.0ml of 100mM DTT; 2.0ml 10 x DIG RNA labelling mix (10mM each of ATP,CTP,GTP, 6.5mM UTP, 3.5mM DIG-UTP (Boehringer), 1ml RNasin (Promega), 20 units of the appropriate RNA polymerase, H2O to 20ml. The mixture is incubated at 37oC for 2 hours, extracted twice with equal volumes of phenol and chloroform/isoamyl alcohol (IAA) (24:1) and once with chloroform/IAA, precipitated with ethanol, washed with 70% ethanol, resuspended in 100ml of 10mM DTT and stored at -70oC in small aliquots (5m more-or-less indefinitely. The optimal dilution of each probe is determined by titration on control sections. Usual the optimal dilution is between 1:500 and 1:2000.
3. Hybridization with digoxygenin-labelled probes.
Probes are
diluted immediately before use in hybridization buffer (see
below), denatured at 75
Hybridization buffer: 1x "salts", 50% formamide, O.1mg/ml yeast tRNA, 10% (w/v) dextran sulphate, 1x Denhardt's. Make up a large volume (100 ml will do around 1000 slides) with highest quality reagents and water (DEPC-treated) and store in aliquots at -20o.
10x "salts": 2M NaCl, 50mM EDTA, 100mM Tris-HCl pH7.5, 50mM NaH2PO4.2H2O, 50mM Na2HPO4
4. Post-hybridization washes.
All incubations are carried out in coplin jars. After overnight hybridization, slides are incubated in wash buffer (1x SSC, 50% formamide, 0.1% Tween-20) at 65oC for at least 15 minutes or until the cover slips fall off, then washed twice further in wash buffer at 65oC for 30 minutes each. The slides are then incubated 2 x 30 minutes in MABT (100mM maleic acid pH7.5, 150mM NaCl, 0.1% (v/v) Tween-20).
5. Blocking and antibody staining.
The slides are dried off around the sections using tissue paper, the sections circled with a Pap Pen (Agar Scientific), transferred to a humidified chamber and incubated in blocking solution (MABT containing 2% blocking reagent [Boehringer] and 10% heat-inactivated sheep serum) for 1 hour at room temperature without a coverslip. The blocking solution is then replaced with anti-digoxygenin AP-conjugated antibody (Fab fragments; Boehringer) diluted 1:1500 in blocking solution, and the incubation continued overnight at 4oC (or two hours at room temperature for strong signals).
6. Post-antibody washes and colour reaction.
The slides
are transferred to coplin jars and washed 3 x 5 minutes in MABT;
then 2 x 10 minutes in prestaining buffer (100mM Tris-HCl pH9,
100mM NaCl, 5mMgCl2). The prestaining buffer is
replaced with staining buffer (100mM Tris-HCl pH9, 100mM NaCl,
5mM MgCl2, 5% (w/v) polyvinyl alcohol (av. Mw 70-100k,
Sigma cat P-1763), 0.2mM 5-bromo-4-chloro-3-indolyl-phosphate
(BCIP, Boehringer), 0.2mM nitroblue tetrazolium salt (NBT,
Boehringer) and incubated in the dark at 37