Xenopus oocytes and embryos: *


Preparation *

On The Day *


Embryo Injection: *

Embryo labelling *

BrdU *

Embryo Histology *

Whole Mount in situ hybridisation *

Wax Embedding *

Rehydration of sections *

Cryostat Sectioning & Non-isotopic in situ hybridisation to sectioned material *

Cryostat Sectioning & Non-isotopic Fluorescence staining of sectioned material, e.g. for F-Actin, (Xenopus Protocol) *

Membrane staining with soybean agglutinin. *

Hoechst Staining of nuclei *



Xenopus oocytes and embryos:


(D.R. Smith)


At least the day prior to the micro-injection prepare the following:

1. Glass capillary needles.

Pull glass micro-injection needles with a micro-electrode puller. Under a binocular microscope break the end of the needle with a tapered tungsten filament (produced by vaporizing the end of a tungsten wire with an oxy-acetylene torch) fused into the end of glass tube as holder. The end of the needle is broken to a diameter of 10-20 um. The tungsten filament is simply used to apply force to the needle. Compare the broken needle end with a standard needle under a magnification of 30x or more and rebreak if necessary. Select needles with single near 45O angled single points. (It is useful though not essential to mark the point side of needle. The point is then oriented to be on the lower needle surface when injecting.

2. Petri dishes.

Prepare small discs of nylon netting of the correct size to fit 9cm petri dishes. Pipette about 1 ml of chloroform into the bottom of the petri dish. Leave for between 10 and 30 seconds. Remove chloroform, and place one of the discs into the bottom of the dish. Press net firmly down against petri, moving around the edge of the netting. When the chloroform appears dry the netting disc should be firmly and evenly stuck to the bottom of the disc. To aid evaporation of chloroform while fixing netting in place a hair dryer can be used or the petri simply held in the airflow into the fume hood. Sterilise dishes with diethyl pyrocarbonate (0.1% in H2O made up freshly) for about 30 mins then wash three to four times with sterile distilled water. Store closed until required. UV sterilisation under the lamp of a culture hood is often also performed just before use.

3. Stock solutions.

DNA or RNA solutions to be injected should be clean (if possible have been purified on a caesium chloride gradient without EtBr) and the concentration should be accurately known. The DNA should be dissolved in injection buffer (10 mM Tris HCl pH 7.5, 88 mM NaCl). From this make up a 10x injection stock. The actual concentration of this 10x stock will vary depending upon what is to be injected. For the standard polymerase I vector (pX1108c) the concentration is 250 ng/ul of insert. For the M13 ribosomal promoter insert the concentration is 280 ng/ul. From the 10x stock, dilute down to a 1x micro-injection stock, still in injection buffer. This can then be used directly in the injection mix (if doing a competition or internal standard type experiment) as follows:

3 ul 1x injection stock of plasmid (i.e. pX1108c)

3 ul 1x injection stock of control (i.e. pX1 ES1)

6 ul Inj. Buffer or alpha-amanitin (500 ug/ml) if required to inhibit polII & III

Mix the solutions well and spin briefly. Store at -20oC until required and then on ice.

4. Modified Barths Media







88 mM





1.0 mM

100 mM




2.4 mM

240 mM


) Solution I


7.5 mM

750 mM





0.82 mM

82 mM


) Solution II



0.33 mM

33 mM


) Solution III


0.41 mM

41 mM



Make up stock solutions and autoclave. For 5 litres of 1x Barths media, take 4 litres distilled H2O, add 88 ml 5 M NaCl and 50 ml of the three stock solutions. Adjust to pH 7.6 with conc HCl. Make up to 5 litres dispense into 500 ml bottles and autoclave. Store at 4oC until just prior to use, then transfer to 18oC incubator.

5. Antibiotic

Aliquot 2 ml from the stock bottle (100x pen. strep. (Gibco)) into 4 ml tubes. Store at -20oC. Thaw and use as required. Add one 5 ml aliquot to each 500 ml bottle of Barths media immediately prior to use.

6. Operating equipment

Sterilise all the operating equipment that will be required, such as scissors, tweezers and arterial clamp. Wash the equipment thoroughly, rinse with distilled water then bake in an oven at 180oC for 2 hours or autoclave.

7. Pasteur pipettes

(These have now been superceded by 1ml pipette tips cut to the appropriate size just before use and flamed). Free oocytes are manipulated using a pasteur pipette whose tip has been broken off to yield a wide lumen and flamed. Sterilise by baking at 180oC for 2 hours.


8. Collection of oocytes

Oocytes are taken from Xenopus laevis or Xenopus borealis females. Best results are obtained from females which have been aclimatised to the lab for several weeks and a week or more after an injection of 50 to 100u of hCG. For removal of the oocytes, the frog is narcotised in 0.5 litre of MS 222 (Sandoz or Sigma) at a concentration of 1 to 1.6 g/litre in tap water at near room temperature. After 5 to 12 minutes exposure (2 min after stimulated movement ceases), depending on size, the frog remains anaesthesized for 1/2 to 1 hour. The frog is placed on its back, on a wet cloth and rinsed with Barths plus antibiotic berfore and at regular intervals during operation. An incision about 1 cm long, slightly offset from the ventral line, is made in the skin. The peritoneum and the muscle tissue are cut open. A small portion of the ovary is pulled out with forceps and removed with a pair of scissors. The clump of oocytes is immediately transferred to a petri dish containing modified Barth medium with antibiotic. The rest of the ovary is pushed back into the body cavity. The peritoneum and the muscle tissue are sewn up and then the skin closed off using cat gut. The oocytes are kept at +18oC or at 4oC and can be used for at least 2 days.

On The Day

9. Preparation of oocytes

Oocytes of stage 5 and 6 are separated from the ovary with a platinum wire loop. These can be distinguished morphologically from the other developmental stages by a change in the animal pole from a dark brown to a light brown or beige, and the formation of a rather distinct lighter coloured boundary between the hemispheres. This colour change occurs when the oocyte reaches a diameter of 1000 to 1200 um. Stage 5 represents about 10% of the stage 2 to stage 6 population. Stage 6 oocytes are identified by their relatively unpigmented equatorial band, which measures about 0.2 mm wide. The animal hemisphere returns to a dark brown colour similar to that of the earlier stages. The size of the oocytes is 1200 - 1300 um and polarisation of yolk platelets in the vegetal hemisphere and smaller platelets in the animal hemisphere is readily apparent.

Free oocytes are manipulated using a pipette tip with a wide lumen. About 40 oocytes are manipulated on to the petri dishes containing the bonded netting. The oocytes are orientated on the netting in such a way that the brown animal pole is upwards. The petri dish and its contents are placed in an MSE 6L or Sorvall RC3, 6x 1 litre rotor bucket and centrifuged for about 10 mins at 18oC at 1000-1600 rpm. The results of centrifugation are variable and it is often best to carry out a small series of tests to determine the best centrifuge conditions. Centrifugation is considered optimal when the dark pigment ring is just displaced, indicating the position of the nucleus. During the centrifugation the oocytes are forced into the meshes of the nylon netting and remain fixed. When the centrifuge has come to a stand still the oocytes will present themselves with the brown pigment ring facing the injector. Centrifuged ocytes are best used immediately. Centrifugation of the oocyte does not impair the capacity to synthesize RNA.

10. Setting up the needle

When using the positive displacement system, the needle is filled with paraffin oil from its non-tapered end by means of a syringe. Care must be taken to ensure that no air bubbles are introduced into the needle. The needle is fixed to the micro-manipulator via the needle holder and a short length of PVC tubing, also filled with paraffin oil, again being careful not to introduce air bubbles.

1.6 to 3 ul of 1x injection solution is delivered from a standard tip onto a piece of Parafilm. From now on all operations are carried out under the binocular microscope. The injection needle is introduced into the drop with the aid of the micro-manipulator. The solution is aspirated into the needle by turning the micrometer screw slowly, to avoid shearing the DNA. The hydraulic system must be free of air bubbles so that when suction is applied, the miniscus within the needle rises immediately and stops rising as soon as the the micrometer stops turning. When most of the liquid has been sucked up, apply positive pressure until a small volume of solution is ejected. Then remove the needle from the remaining drop of liquid and immediately immerse the needle in some buffer to prevent drying until required.

11. Injection

Place the petri dish containing the centrifuged oocytes under the binocular microscope. The needle attached to the micromanipulator is aimed at the oocytes at an angle of about 60-70o. The oocytes are aligned relative to the tip of the needle by manoeuvering the petri dish. Before injecting the first oocyte it is advisable to ensure that the system is working by pressing the foot control until a drop of liquid can be seen. Repeat at least once through the course of the injection (i.e. after about 20 oocytes). The tip of the needle is introduced into the dead center of the ring of displaced pigment and just below the surface of the oocyte. 20 nl of the injection fluid is injected into each oocyte nucleus using the micrometer movement. An alternative method is to continuously run the drive motor running and to count the seconds that the needle remains in the oocyte. This method has the great advantage that it is extremely reproducable and that there is less chance that the needle will become plugged by yolk. This method is also less sensitive to any air within the hydraulic system.

Any uninjected oocytes are removed from the nylon with a jet of media. The oocytes are then incubated at 18oC (23oC must not be exceeded) for the desired time (about 14-18 hours).

12. RNA extraction

Injected and incubated oocytes are removed from the nylon mesh by a jet of media as described above, and oocytes showing an unequal distribution of pigment are rejected. The remaining oocytes are transferred to a disposable 4 ml or eppendorf tube. Remove all of the barths media and resuspend in 400ul room temperature TE or RSB:

1x RSB:

10 mM Tris HCl pH 7.5

200 ul 1 M

1.5 mM MgCl2

30 ul 1 M

10 mM NaCl

40 ul 5 M



total volume

20 ml

Add 50 ul Proteinase K (20 mg/ml), gently whirlimix and leave 1 min. Add 50 ul 20% SDS and whirlimix vigorously until all the oocytes have lysed. Incubate for about 1 hour at room temperature.

After the above stage the oocytes can either be frozen or they can be taken through the next steps.

Add 50 ul 3 M NaAc (or NaCl) and whirlimix. Extract 3 times with phenol-chloroform and once with chloroform.

Add 2.5 vol. ice cold EtOH, mix well and leave at -20oC 1hr.

Spin 12,000 rpm, 10 mins and 4oC in Sorvall RC-5B, pour off supernatant and wash with 80% EtOH 12,000 rpm, 5 mins.

Dry samples (but be careful not to over dry) and for S1 mapping resuspend in formamide at a concentration of 4 ul/oocyte. Resuspension can prove to be difficult. Incubation at 65oC for a short time can help.


Have ready the following prior to the days micro-injecting:

1. Glass capillary needles (broken)

2. Petri dishes (with netting) and a few without netting

3. Stock solutions of DNA

4. Modified Barths media

5. Antibiotic solution

6. Operating equipment (sterilized by baking)

7. Optional wide lumen pipettes (sterilized by baking) or 1 ml cut pipette tips lightly flamed.

On the Day:

8. Operate frog to recover ovary fragment

9. Remove oocytes with wire loop

10. Manipulate about 40 oocytes onto a tray (for each solution to be injected

11. Start centrifugation of one tray of oocytes (10 mins, 1,200 to 1,600 rpm, 18oC)

12. Fill needle with paraffin oil from blunt end using syringe

13. Insert into needle holder

14. Put 3 ul of injection solution onto piece of parafilm

15. Under binocular microscope slowly aspirate solution

16. Place tip of needle in buffer until ready to inject

17. Under binocular microscope inject each oocyte

18. Remove any uninjected oocytes with a jet of medium

19. Incubate at 18oC 14-18 hours

20. Resuspend in 500 ul RSB

21. Add 50 ul Proteinase K (20 mg/ml), mix and leave 1 min.

22. Add 50 ul 20% SDS, whirlimix until oocytes totally lysed

23. Incubate at room temp 1 hour

24. Add 50 ul 3 M NaAc (or NaCl).

25. Extract 3 times with 1 volume phenol/chloroform

26. Extract once with 1 volume chloroform

27. Add 2 volumes ice cold EtOH, mix well, -20oC 1hr

28. Spin 12,000 rpm, 10 mins and 4oC. Pour off S/N and discard

29. Wash with 80% EtOH 12,000 rpm, 5 mins and 4oC. Pour off S/N and discard

30. Dry pellet lightly

31. Redissolve in formamide at a concentration of 4ul/oocyte

32. Store at -20oC or -70oC.


Embryo Injection:

Female Xenopus are induced to ovulate with human Chorionic Gonadotropin (CG) from ICN dissolved at 1000 I.U./ml in sterile distilled water or 1/10 Ringer's:-

1x Ringer’s:-




0.15 gm




0.15 gm



final volume

1 l

Normally females are injected with 100 I.U., 0.1ml, 2 days before eggs are required, then again with 400 - 700 I.U., 0.4-0.7 ml, 8 to 12 hr before.

Females may be kept in normal aquarium water or if the eggs they lay may be required for fertilisation, in High Salt-MBS.

Just before laying a male is sacrificed and the testes taken into 1 x MMR, 10% foetal calf serum, 100ugm penicillin, 50ugm streptomycin.

1x MMR,


0.1M NaCl

20ml 5M NaCl

2mM KCl

1ml 1M KCl

1mM MgSO4

1ml 1M MgSO4

2mM CaCl2

2ml 1M CaCl2

5mM Hepes

10ml 0.5M Hepes

0.1mM EDTA

0.2ml 0.5M EDTA

Final pH 7.8.


Eggs are massage from female into 10 - 15 ml of 1 x MMR for immediate use or into High Salt-MBS for storage, (up to 12hr is claimed for very good batches, but this does not usually work for us). The testes is opened slightly at one end with forceps and wiped over the eggs in a minimal solution volume. The eggs are then flooded with 0.1 or 0.2 x MMR.

10x MBS (less CaCl2)











pH to 7.6 with NaOH


Make up to 1l and filter sterilise.



10 x MBS (No CaCl2)


0.1M CaCl2




Gentamycin (50mg/ml)


(or 100u/ml penicillin, 100ug/ml streptomycin SO4 (Gibco))


High Salt MBS

10 x MBS (No CaCl2)


0.1M CaCl2


5M NaCl




Gentamycin (50mg/ml)


The first division occurs in about 1.5hr, the second after another hour. At the earliest 20 to 30 min. after fertilisation the eggs are dejellied in cysteine soln for two minutes, use 15 - 20 ml per batch of 50 - 100 embryos change after 2 to 3 min. for fresh if necessary. Keep time of dejelling as short as possible, around 5 min. though 6 to 10 min. has been used quite successfully as long as subsequent wash is thorough.

Cysteine solution, make fresh :- 100ml

cysteine HCl

2 gm

NaCl (5M)


10M NaOH to pH 7.8-7.9.

1.5 ml

Wash very thoroughly with 0.1 or 0.2 x MMR (e.g. 6 x 100ml) finally transfering into the injection buffer, 0.2 x MMR, 5% Ficoll 400.

Inject between 35 - 100 pgm of DNA or up to 2 ngm RNA in a volume as small as possible. Generally a maximum of 10 to 20 nl is used but smaller is better, much smaller volumes ~50pl have been used by some. Inject into the animal pole of one or two cell embryos.

Leave at 14 to 18oC in 0.2 x MMR, 5% ficoll 400 for a few divisions or up to 32 cell stage, then transfer to 0.1 x MMR and incubate O/N at 14 to 18oC adding 1ug/ml Gentamycin or penicillin streptomycin (100x solution Gibco) if necessary. Lower temperature incubation leaves embryos at gastrula next morning, but we have found that viability is often poorer than at higher temperature.



Embryo labelling


Embryos are placed in 0.1mM BrdU, 0.1 x MMR for 2 to 4 hr before fixation or further treament.


Embryo Histology

Whole Mount in situ hybridisation

Islam & Moss (1996) TIGs 12, 459

Incubations were performed in a 24 well tissue culture plate and embryos contained in ~1ml plastic cups with nylon net bases. These were prepared from 1.5ml microcentifuge tubes by removing the tapered section of each tube and welding this to a fine nylon net. R/T incubations were performed on a Nutator, while those at elevated temperatures were performed in a shaking water bath.

1) Fixation of embryos;

The embryos are fixed at R/T in MEMFA (0.1M MOPS, pH 7.4, 1mM EGTA, 2mM MgSO4 and 3.7% formaldehyde for two or more hours, (possibly O/N). After fixation the embryos are transfered to methanol and can be kept many months at -20oC.

MEMFA; (0.1M MOPS, pH 7.4, 1mM EGTA, 2mM MgSO4 and 3.7% formaldehyde

2) Rehydration;

The embryos are rehydrated at R/T by sequential washings in 75% methanol-25% PBST, 50% methanol-50% PBST, 25% methanol-75% PBST and finally PBST, each 5 min.

PBS; 0.145M NaCl, 10mM Na-phosphate pH 7.4

PBST; PBS containing 0.1% Tween 20)

3) Enzyme treatment.

a) Hatched embryos; Rehydrated embryos are incubated at R/T in 10 ug.ml-1 proteinase K, PBS for 20 to 30 min. (20 min stage 23 to 30, 30 min for later stages).

b) Prehatching embryos; Rehydrated embryos are incubated at R/T in 10 ug.ml-1 proteinase K (Sigma), collagenase A (Boehringer) 2 mg.ml-1, hyaluronidase 20u.ml-1 (Sigma), PBS for 10 min. only.

The embryos are then briefly rinsed with PBST.

4) Refixation

The embryos are refixed in 4% paraformaldehyde in PBS for 20 min. at R/T.

5) Inactivation of endogenous alkaline phosphatase

Embryos are briefly rinsed in hybridising solution and then incubated in fresh hybridising solution (HB) at 75oC for 30 min.

Hybridising solution (HB); 5 x SSC pH6.0, 50% formamide, 50ug.ml-1 heparin, 0.1% Tween 20, 0.5mg.ml-1 torula RNA.

6) Prehybridisation

Without change of solution, simply lower the temperature to hybridising temperature (55 to 70oC) and incubate 4 to 5 hours.

7) Hybridisation

Change HB for fresh preheated solution 30 min before beginning hybridisation. Add 500ng.ml-1 of digoxygenin labelled riboprobe and hybridise O/N.

8) Washings

Incubate at temperature of hybridisation Thyb in;


10 min.

75% HB-25% 2 x SSC-0.1% Tween 20

10 min.

50% HB-50% 2 x SSC-0.1% Tween 20

10 min.

25% HB-75% 2 x SSC-0.1% Tween 20

10 min.

2 x SSC-0.1% Tween 20

10 min. x 3, at Thyb plus 2 to 5 oC

0.2 x SSC-0.1% Tween 20

30 min. Thyb plus 2 to 5 oC

0.2 x SSC-0.1% Tween 20

10 min. Optional.

2 x SSC-0.1% Tween 20

10 min. Thyb plus 2 to 5 oC.

The embryos are brought to R/T and washed in PBST for 3 times, 5 min each.

9) Detection with alkaline phosphatase

a) Block for 1 hr at R/T with 5% sheep serum, 2mg.ml-1 BSA (Cohn fract. V), 1% DMSO in PBST.

b) The antidigoxygenin phosphatase conjugated Ig is preabsorbed with heat inactivated embryo powder for 1 hr at R/T. 10mg of embryo powder is resuspended in 500ul of PBS and heated at 75oC for 30 min and centrifuged for 1 min in a microcentrifuge to recover the pellet, which was then resuspended in 500ul of blocking solution. 2.5ul of antibody was then added and the solution incubated for 1 hr at R/T. The solution is then centrifuged to remove all embryo powder. The clear preabsorbed antibody solution was then diluted to 5ml with blocking solution (final Ig dilution 2000x). 1.2 ml of preabsorbed antibody solution was delivered into each well of embryos and incubated for 2hr at R/T.

Embryo Powder

Late embryos/early larvae were anesthetised in 3-amino benzoic acid ethyl ester (40ug. ml-1, in PBS, pH7.0). The embryos are then frozen in liquid N2 and ground to a fine powder under N2 in a mortar. The powder was then transfered to a centrifuge tube, cold actone added and left on ice for 30 min with occasional vigorous shaking. After recovering the powder by centrifugation, 12,000g, 10 min, it was again resuspended in cold acetone and left on ice for 10 min with occasional vigorous shaking. The centrifugation was repeated and the pellet dried under vacuum and stored at -20oC in a vacuum dessicator.

10) Washing

The embryos were washed 12 x in 2 ml per well of PBST, 15 min each. The washings may be continued O/N at 4 oC if necessary.

11) Staining

Just prior to staining the embryos were washed in 2 changes of NTMT, 15 min each at R/T. They are then transfered to a clean 24 well plate and incubated at R/T in developing solution. Development is judged visually and the developing solution changed for fresh as soon as it becomes slightly coloured. Development is terminated by washing once or more with PBS and then fixed in MEMFA (0.1M MOPS, pH7.4, 2mM MgSO4, 1mM EGTA and 3.7% formaldehyde) for 5 min or more, (they may be stored in this solution at 4 oC).


100mM tris-Cl pH9.0, 100mM NaCl, 5mM MgCl2 and 0.1% Tween-20.

Developing Solution;

50ul NBT (4-nitro blue tetrazolium chloride) and 37.5ul of BCIP (5-bomo-4-chloro-3-indolyl-phosphate, 4-toluidine salt) (Boehringer) are added to 10ml of NTMT.

12) Clearing

The embryos may be cleared (in glass vials) by washing;

1 x PBS, 1x methanol, 1 x 50%, methanol-50% benzyl benzoate and finally in 1 part benzyl alcohol, 2 parts benzyl benzoate. This solution will leach some colour and so should not be used until just before photographic recordings are made.

Wax Embedding

After fixation as for wholemount or after wholemount hybridisation the embryos are embedded as follows:-

Either dehydrate stepwise, 10 min. each through;

50% EtOH 50% saline (0.85% NaCl),

70% EtOH in water,

85% EtOH in water,

95% EtOH in water,

100% EtOH,

or ;

100% EtOH directly,

Then 30 min. in each of;

1) 100% dry EtOH (Store 99% EtOH over Type 3A molecular Sieves (Fisher),

2) Dry EtOH : Toluene :: 1 : 1,

3) Toluene

4) Paraffin wax (Paraplast : Tissueprep 2 :: 1 : 1, (both Fisher) melted at 60 to 62 oC O/N). This step is best done directly in the moulds.

5 to 7) Repeat the treatment with paraffin three more times.

8) Finally remove the moulds from the oven and rapidly orient the embryos under dissecting microscope while the wax begins to set. A heat platinum wire may help.

Sectioning & mounting Wax Embedded embryos

The blocks are left at R/T to set then carefully placed at 4oC O/N. They are then sliced at 10um. 5 to six slices are cut from ribbon and dropped onto the surface of 45oC preboiled water in a constant temperature bath. The sections are then lifted out on a Tespa treated slide, see below. The slides are placed on a slide warmer at 37oC for 30 min. or just left standing to drain and finally placed in a slide box containg dessicant, sealed and heated at 37oC O/N. They are then stored in the same box with dessicant at 4oC until required.

Slide Preparation with Tespa

Long Protocol, (RNase free):

1. Soak slides in 1M HCl (or 10% HCl, 70% EtOH) for 15min.

2. Soak in hot tap water containing a dash of detergent ( e.g. Beckman rotor cleaning detergent) for 30 min.

3. Rinse in running tap water 45 min.

4. Rinse in Milli-Q water, leave for few minutes.

5. Bake overnight 250oC.

6. Cool slides and place for 30 to 60 sec in 2% 3-aminopropyl-triethoxysilane (TESPA, Sigma A3648) in acetone (9ml in 450ml acetone) at R/T.

7. Dry at 42oC ( no higher than 55oC) for 30 - 60 or more minutes.

  1. Store at R/T dust free.


Short Protocol:

1. Soak slides in 10% HCl, 70% EtOH for 15min followed by EtOH 95%, 15 min..

2. Baked to dry at 150oC for 5 min. and allow to cool.

3. Dip slides in 2% 3-aminopropyl-triethoxysilane (TESPA, Sigma A3648) in acetone (9ml in 450ml acetone) for 10 sec.

4. Wash twice with acetone, once with Milli-Q water.

5. Dry at 42oC ( no higher than 55oC) for 30 - 60 or more minutes.

  1. Store at R/T dust free.


Protocol of Strähle, Adam & Ingham (TIG 10, 75-76, 1994):

1. Wash slides overnight in 1% HCl, 70% ethanol.

2. Rinse in H2O and dry at 70oC.

3. Submerge in 2% (v/v) TESPA (3' aminopropyl-triethoxy silane, Sigma A3648) in acetone for about 10 s.

  1. Rinse in acetone for 10 s, then rinse in H2O for 10 s and bake at 160oC

Rehydration of sections

To remove paraffin wax and rehydrate sections place slides for 5 min. in each of the following;

1) Xylene,

2) Repeat with fresh Xylene

3) 95% EtOH

4) 85% EtOH

5) 50% EtOH

6) H2O

7) PBS or other aqueous buffer as required, e.g. for hybridisation or staining.

Cryostat Sectioning & Non-isotopic in situ hybridisation to sectioned material

Originally for Zebrafish embryos. Strähle et al. TIG 10, 75-76, (1994).

Fixation And Sectioning

1. Fix zebrafish embryos in BT fix (4% w/v paraformaldehyde, 4% w/v sucrose, 0.12 mM CaCl2, 0.1 M NaP04 pH7.4: store at 40C for up to 7 d) at 40C overnight.

2. Wash embryos twice in BT fix without paraformaldehyde. Remove chorions with sharp watchmaker's forceps.

3. Embed embryos in 1.5% agar (Gibco BRL), 5% sucrose.

4. After trimming, transfer agar blocks to 30% sucrose, 0.1% azide at 40C until blocks sink (usually overnight).

5. Cut 15 um cryostat sections. Transfer sections to TESPA-coated slides. [Wash slides overnight in 1% HCl, 70% ethanol. Rinse in H20 and dry at 700C. Submerge in 2% (v/v) TESPA (3' aminopropyl-triethoxy silane, Sigma) in acetone for about 10 s, rince in acetone for 10 s, then wash in H20 for 10 s and bake at 160oC]. Air-dy sections for 2 h at room temperature, then store at -200C in an air-tight box over silica gel (we have stored sections for up to one year).

Probe Preparation

1. Synthesize digoxigenin- and fluorescein-labelled antisense RNA probes according to the protocol recommended by the supplier of the modified digoxigenin and fluorescein nucleotides (Boehringer Mannheim). Best results are obtained using transcripts > 1 kb; shorter transcripts work, but some detection sensitivity is lost. For use with fish sections, probes need not be hydrolysed, although when working with chick sections, hydrolysed probes may give better results.

2. Store probes in 50% formamide at -200C.


1. Dilute antisense RNA probe in hybridization buffer [0.3 M NaCl, 10 mM NaPO4, 10mM EDTA, 10 mM TrisHCL pH 7.5, 50% formamide (Fluka), 10% dextran sulphate (Pharmacia), 1 mg ml-1 rRNA (Sigma R7125), 1 x Denhardt's (modified from Ref.6)]. Dilution is usually 1:100, but the optimal dilution can vary for different probes. In double-labelling experiments, the digoxigenin-labelled and the fluorescein-labelled probes are mixed at the appropriate dilutions. Denature the probe mix at 700C for 5 min immediately before applying to the sections.

2. Add 30-50 ul diluted probe to each slide and coverslip. Initially the probe might not cover all sections completely. This is not a problem, as it will spread out during hybridization.

3. Place slides on filter paper soaked with 50% formamide, 0.3 M NaCl, 10 mM NaPO4, 10 mM EDTA, 10 mM TrisHCl pH. 7.5 in a sealed box. Hybridize at 550C for 8 h or overnight.

4. Transfer slides to a slide rack and submerge in prewarmed (650C) 50% formamide, 2 x SSC for 15 min, to detach coverslips. Repeat wash at 650C for 30 min.

5. Wash twice in 25% formamide, 1 x SSC, 0.5 x PBS at 650C for 30 min.

6. Transfer to PBS at room temperature for 5 min, and then block for 30 min in PBT [PBS, 0.2% (w/v) BSA, 0.1% Tween 20].

Preabsorption of Anti-Digoxigenin and Anti-Fluorescein Antibodies

1. Make zebrafish powder by sacrificing adult zebrasfish in 3-amino benzoic acid ethyl ester (40 ug ml-1, pH7). Freeze fish in liquid nitrogen and grind to a fine powder under nitrogen in a mortar. Transfer the powder into a centrifuge tube, add cold acetone and incubate on ice for 30 min, with occasional vigorous shaking. Spin in a cooled Sorvall centrifuge (HB4 rotor),10 000 r.p.m. for 10 min. Resuspend pellet in ice-cold acetone, and incubate on ice for 10 min, shaking occasionally. Repeat the centrifugation step and resuspend pellet in acetone. Dry on a filter paper and store at 40C in a sealed vial. Chick embryo powder is prepared as in Ref. 7.

2. Dilute anti-digoxigenin (DIG) and anti-fluorescein alkaline phosphatase conjugated antibodies (Boehringer Mannheim) 1:400 and 1:100 in PBT, respectively.

3. Incubate antibodies with 6 mg ml-1 zebrafish powder at 40C overnight. Centrifuge to remove tissue debris.

4. Dilute preabsorbed anti-DIG and anti-fluorescein antibody in PBT to give 1:2000 and 1:500 final dilutions, respectively.

Antibody Staining

1. Add 30 ul antibody to each slide and coverslip. Incubate at room temperature for 30 min or in a box on filter paper soaked with PBS at 40C overnight.

2. Was for times for 10-20 min in 1 x PBS, 0.1% Tween 20.

3. Transfer slides into staining jars with NBT-BCIP or Vector Red staining solution. [NBT-BCIP staining solution is prepared freshly: 100 mM NaCl, 50 mM MgCl2 100 mM TrisHCl pH 9.5, 0.1% Tween 20, 5 mM levamisole, 0.34 mg ml-1 nitroblue tetrazolium salt (NBT, Gibco BRL) and 0.175 mg ml-1 5-bromo-4-chloro-3-indolyl-phosphate (BCIP, Gibco BRL), Vector Red staining solution is prepared according to the manufacturer's instructions (Vector Labs).]

4. Allow colour to develop in the dark at room temperature for several hours or overnight. Slides can be removed, checked for staining and returned to the staining jar.

5. Wash in 1 x PBS, 0.1% Tween 20 for 10 min.

6. Apply second antibody (repeating steps i-iv) or dehydrate (1 min each step in 30, 50, 75, 95 and, finally, 100% ethanol).

7. Clear in histoclear and mount. (The red precipitate obtained with the Vector Red alkaline phosphatase substrates fluoresces strongly when a rhodamine filter set is used).

Cryostat Sectioning & Non-isotopic Fluorescence staining of sectioned material, e.g. for F-Actin, (Xenopus Protocol)

Originates from Drechsel et al. Curr. Biol.7, 12-23.

TBSN:- 50mM Tris-HCl, 155mM NaCl, 0.1% NP-40.

PEMFA:- 4% paraformaldehyde, 0.5% glutaraldehyde, 80mM PIPES, 5mM EGTA, 1mM MgCl2, 0.2% Triton X100, pH7.5.

5X stock;


24g ,




1ml 1M



final volume




(Dissolve in PEMFA & water by boiling.)


50% glutaraldehyde


10% Triton


5x stock PEMFA




Final volume


i) At selected development stage flood embryos with 4% paraformaldehyde, 0.5% glutaraldehyde, 80mM PIPES, 5mM EGTA, 1mM MgCl2, 0.2% Triton X100, pH7.5 (PEMFA) and fix for 2hr at R/T.

ii) Wash twice in TBSN (50mM Tris-HCl, 155mM NaCl, 0.1% NP-40, pH 7.5) and stored overnight in TBS or PBS.

iii) Transfer to PBS, 1.6M sucrose and leave for >12 hr.

iv) 6 to 10 embryos are placed in a drop of Tissue-Tek embedding solution (Miles) and frozen on a metal block on dry ice.

v) Cut 16mm sections and pick up on gelatin coated coverslips (slides?) and allowed to air dry for 30-60 min.

vi) Sections are then postfixed in 2% formaldehyde, PBS, washed in PBS and treated with 3mg.ml-1 NaBH4 in PBS(?) for 15 min to reduce fluorescence.

vii) Sections are blocked with 5% appropriate normal serum in TBSN for 30min and then incubated for 1 - 2hr with first antibody in 5% serum in TBSN blocking solution.

viii) After four washes with TBSN, sections are incubated with second antibody as for the first one.

ix) rhodamine-phalloidin staining for F-actin is done by addition to second antibody solution.

x) After four washes in TBSN, sections are mounted in 50% glycerol. 100mM Tris-HCl pH 9, 1mg.ml-1 p-phenylenediamine.

Gelatin coating of coverslips and slides.

i) Add 2.5 g (0.5%) of gelatin to 500ml boiling water.

ii) When dissolved remove from heating and add 0.25g chromium potassium sulphate (chrome alum).

iii) Filter solution.

iv) Soak slides for 2 min in the solution and then dry at 60oC or O/N at R/T.

Membrane staining with soybean agglutinin.

Originates from Drechsel et al. Curr. Biol.7, 12-23.

Lectins are present on the surface membrane of the embryo animal pole. By binding these with biotinylated soybean agglutinin before the first division, the recruitment of new and hence unbound membrane can be followed by subsequent staining with fluorescent or enzyme conjugated anti-biotin antibodies.

i) After fertilisation and early standard cysteine degellying, vitelline membrane is removed manually, embryos placed in agarose wells made using 1mm ball bearings

ii) Place embryos in 50mg.ml-1 biotinylated soybean agglutinin, 1 x MMR for 10 min..

iii) Wash in 1 x MMR and inject as normal.

iv) At selected development stage flood embryos with 4% paraformaldehyde, 0.5% glutaradehyde, 80mM PIPES, 5mM EGTA, 1mM MgCl2, 0.2% Triton X100 (PEMFA) and fixed for 2hr at R/T.

iv) Staining is done on cryosections (or have tried whole embryos).


Hoechst Staining of nuclei

100 x Hoechst stock solution (20ug/ml in H2O, kept at 4oC) is diluted in H2O and the rehydrated slides treated with this for 5 to 10 min. Finally slides are washed in H2O and mounted in aqueous solution. The cover slips are sealed with clear nail varnish.