In situ hybridization on whole mount embryos of C.elegans

Tomoko Motohashi, Keiko Hirono, and Yuji Kohara
Genome Biology Lab
Center for Genetic Resource Information
National Institute of Genetics
Mishima 411-8540, Japan

Corrected by Ikuko Muramatsu, Masumi Obara, Wakako Shimizu, Aya Hamakawa and Yuji Kohara (April 27, 2005)
Correspondence: Ikuko Muramatsu <>, Yuji Kohara <>


  1. Preparation of DIG-labeled DNA probes for in situ hybridization
  2. Fixation of embryos from a small number of worms
  3. Large scale fixation of embryos
  4. Hybridization and signal detection
  5. Reagents

A. Preparation of DIG-labeled DNA probes for in situ hybridization

I. DIG-labeling by linear PCR

  1. 1. Make the following reaction mix (total 10 );
    distilled water4.4
    anchored oligo dT primers *3.0
    10 x Taq buffer1
    Taq polymerase0.1
    10 x DIG-dUTP/dNTP mix **1.5
    cDNA insert (> 3.5 ng/) amplified from yk clone using T7/T3 primers0.6
    * anchored oligo dT primers
    5.3 (dT)17dG / 5.3 (dT)17dC / 21.3 (dT)17dA
    (This is to avoid the effect of poly-A stretch. Other vector primers may be used.)
    ** 10 x DIG-dUTP/dNTP mix
    0.35mM DIG-dUTP / 0.65mM dTTP / 1mM each d(A, G, C)TP
  2. Subject to thermal cycling;
     hot start at 95oC for 45 sec
     95oC x 15 sec
     45oC x 1 min
     72oC x 1 min
    50 cycles
  3. Add 10 of 10mM EDTA.
  4. Pass through G-50 spin colum chromatography (ca. 250 bed volume).
  5. Add 5 of TSE.

II. Chopping the probes by DNaseI digestion

  1. Make the following reaction mix (total 25 ) on ice;
    The G-50 elutate20
    10mg/ml Salmon sperm DNA1 μ
    distilled water0.5
    10 x DNase buffer *2.5
    DNaseI (16 /ml)**1
    * 10 x DNase buffer: 0.5M TrisHCl pH 7.5, 0.1M MgCl2
    ** Dilute stock solution (1mg/ml) with 0.1M NaCl
    (Note : Best size of probes is about 100 bases. Longer probes may cause high background. The concentrations of the enzyme should be optimized by pilot experiments.)
  2. Incubate at 37oC for 30 min.
  3. Transfer on ice.
  4. Add 5 of 0.1M EDTA.
  5. Heat at 75oC for 5 min.
  6. Check the size by alkaline agarose gel electrophoresis and DIG detection, if necessary.
  7. Store frozen.

B. Fixation of embryos from a small number of worms

  1. Take siliconized 1.5 ml eppendorf tubes.
  2. Place about 100 of distilled water on the (inside) top of the lids.
  3. Pick and transfer 40-50 gravid worms into the water. If you need very late stage embryos;
    1. Add 50 of suspension of E.coli OP50 in S-basal.
    2. Cover the lid with the body of the tube.
    3. Let stand at 20oC overnight.
  4. Spin down the worms.
  5. Add equal volume of 2 x alkaline-bleach solution and mix well.
  6. Leave at r.t. for 10 min to dissolve the adult bodies.
  7. Add 1ml of M9 buffer at 4oC.
  8. Centrifuge at 2500 rpm for 30 sec at 4oC in a swing rotor.
  9. Remove the sup carefully, leaving about 100 of the sup not to remove the embryos.
  10. Repeat 7. - 9. three more times.
  11. Add equal volume of 3 mg/ml chitinase.
  12. Mix and incubate at r.t. for 3 min.
  13. Spin at 2500 rpm for 30 sec at 4oC.
  14. Reduce the volume to about 50 .
  15. Transfer the embryos to a poly-L-lysine coated 3-well slide using a siliconized pipette tip.
  16. Add a half volume of 4% gelatin, 2% BSA, and mix gently by pipeting.
  17. Let stand for several minutes to allow the embryos settled down to the bottom.
  18. Cover with a cover slip (24 x 40 mm).
  19. Place it on the top of dry ice block.
  20. Freeze for 7 min at -70oC.
  21. Peel off the cover slip quickly.
  22. Soak the slide in methanol cooled at -20oC for 5 min.
  23. Rehydrate by soaking the slide in the series of the following solutions pre-cooled at 4oC;
    methanolfor 5 min
    methanol : formaldehyde-Hepes-PBS* = 35 : 15for 2 min
    methanol : formaldehyde-Hepes-PBS* = 25 : 25for 2 min
    methanol : formaldehyde-Hepes-PBS* = 15 : 35for 2 min
    formaldehyde-Hepes-PBS*for 20 min
    * formaldehyde-Hepes-PBS
  24. Dehydrate by soaking the slide in the series of the following solutions at r.t.;
    ethanol : PBS = 15 : 35for 5 min
    ethanol : PBS = 25 : 25for 5 min
    ethanol : PBS = 35 : 15for 5 min
    ethanolfor 5 min x 2 times
  25. Store in ethanol at -20oC.

C. Large scale fixation of embryos

I. Harvesting of embryos

  1. Get a plenty of worms from a mixed stage population.
  2. Wash the worms 2 times with M9 buffer.
  3. Collect L1-L3 by sieving through 50 Nylon mesh.
  4. Allow the collected worms to grow to young adults in liquid culture.
  5. Take 1 ml packed worms from the culture, which will give 8-15 slides for in situ.
  6. Resuspend the worms in 4ml water in a 15ml Falcon tube (clear type).
  7. Add 5ml of 2 x alkali-bleach solution, mix well and let stand for 10 min.
  8. Force the worms ou through a 23-gauge needle onto nylon mesh.
  9. Collect embryos by spinning the filtrate at 800 x g using a swing rotor.
  10. Wash the embryos 4 times with M9 and transferred into a siliconized eppendorf tube.

II. Fixation

  1. Take 100 (packed volume) of the embryos and adjust the volume to 200 with M9.
  2. Add 200 of yatalase (15mg/ml in 0.3M mannitol) and vigourously shake for 75 sec.
  3. Wash the embryos 3 times with EH buffer (Embryo Handling buffer).
  4. Wash the embryos with Basal EH buffer.
  5. Resuspend the embryos in 1 ml of Basal EH buffer. (Note : For success, it is desired that 20-30% of embryos are devitellinized at this step.)
  6. Place 30 /well of Basal EH buffer onto each well of poly-L-lysine coated 8-well slides.
  7. Dispense 5 /well of the embryo suspension into the buffer at each well.
  8. Let stand for 10 min at 4oC to settle the embryos to the bottom.
  9. Remove the buffer, and immediately immerse in methanol at -20oC for 5 min.
  10. Rehydrate the embryos by immersing the slides in the follwoing series at 4oC. The solutions must be pre-cooled at 4oC.
    methanol5 min
    methanol : 3.7% formaldehyde in hepes-PBS = 7 : 3 2 min
    methanol : 3.7% formaldehyde in hepes-PBS = 1 : 1 2 min
    methanol : 3.7% formaldehyde in hepes-PBS = 3 : 7 2 min
    3.7% formaldehyde in hepes-PBS 75 min at 22oC
  11. Dehydrate the embryos by immersing the slides in the follwoing series at r.t.
    ethanol : PBS = 3 : 75 min
    ethanol : PBS = 1 : 15 min
    ethanol : PBS = 7 : 35 min
    ethanol5 min x 2 times
  12. 12. Store in ethanol at -20oC.

D. Hybridization and signal detection

I. Proteinase K treatment

  1. Rehydrate the embryos by immersing the follwoing ethanol series;
    0.03% H2O2 in ethanol : PBS = 7 : 3 2 min
    ethanol : PBS = 1 : 15 min
    ethanol : PBS = 3 : 75 min
  2. Wash the slides once by immersing in PBT for 5 min.
    *For late stage embryos, additional HCl treatment is effective, whcih can cut glycosid bonds of the proteoglycan that appear on late stage embryos.
    1. Immerse the slides in 0.2N HCl for 20 min at r.t.
    2. Wash the slides 2 times in PBT for 5 min.
  3. Immerse the slides in proteinase K (10 /ml in PBT) and incubate at r.t. for 11 min.
  4. Stop the digestion by immersing the slides in 2 mg/ml glycine in PBT for 2 min.
  5. Wash the slides twice by immersing them in PBT for 2 min each.
  6. Refix by immersing the slides in 3.7% formaldehyde in hepes-PBS at r.t. for 50 min.
  7. Wash the slides twice in PBT for 5 min each.
  8. Immerse the slides in 2 mg/ml glycine in PBT at r.t. for 5 min.
  9. Wash the slides in PBT for 5 min.

II. Pre-Hybridization

  1. Immerse the slides in the following series of mixtures;
    50% formamide, 5xSSC, heparin, 0.1% Tween : PBT = 1 : 1 10 min
    50% formamide, 5xSSC, heparin, 0.1% Tween 10 min
  2. Wipe off the slides
  3. Draw a rectangle surrounding the sample wells using a IMMUNO pen to make a ridge.
  4. Add 250 of heat denatrued (at 99oC for 10 min and quickly chilled for 5 min) hybridization solution for each 8-well slide.
  5. Place the slide in a moist chamber containing a paper towel wetted with 50% formamide, 5XSSC. (No need to use coverslips.)
  6. Incubate at 48oC for 1 hr.

III. Hybridization

  1. Add 50 of heat-denatured DNA probes for each slide. (The final concentrations of probes is about 0.06 /ml.)
  2. Cover the slide with a parafilm coverslip to reduce evaporation.
  3. Incubate the slides at 48oC overnight in the moist chamber.

IV. Washing

  1. Wash the slides in the following series of washing solutins at 48oC with slight agitation.
    50% formamide, 5xSSC, heparin, 0.1% Tween : PBT = 1 : 1
    (First washing is performed in separate containers for every slides.)
    2 min
    50% formamide, 5xSSC, heparin, 0.1% Tween : PBT = 1 : 1 10 min x 2 times
    0.8xPBS, 0.1% CHAPS 20 min x 4 times
  2. . Wash the slides twice in PBT for 5 min at r.t. to remove CHAPS.

V. Probe detection

  1. Incubate the slides in PBtr (PBS, 0.1% Triton-X100, 0.1% BSA, 0.01% NaN3) for 1.5 hr at r.t..
  2. Cover the embryos with 250 of anti-DIG conjugate (dilute 1 : 2500)/8-well slide.
  3. Incubate for 2 hrs at r.t. in a moist chamber. (NO need to use coverslips.)
  4. Wash the slides with PBtr 4 times with slight agitation.
  5. Wash the slides with the staining buffer (see reagetnts) twice for 5 min each at r.t..
  6. Colour development
    1. Mix 180 of NBT and 140 of BCIP in 40 ml of staining buffer.
    2. Immerse the slides in the mixture for 1 hr at 22oC in the dark, monitoring the extent of the staining.
  7. Wash the slides three times with PBS, 20mM EDTA to stop the reaction.
  8. If necessary, incubate the slides in 1 /ml DAPI in Tris buffer at 4oC for 30 min.

VI. Mounting

VI.A. Permanent mount 1.

  1. Add about 90 of "MOUNT-QUICK AQUEOUS" onto the embryos on the slide.
  2. Cover with a coverslip.
  3. Let stand one day to dry up.
  4. Seal up the edge of the coverslip using nail varnish.

VI.B. Permanent mount 2.

  1. Dehydrate with the following ethanol series;
    ethanol : PBS = 3 : 75 min
    ethanol : PBS = 1 : 15 min
    ethanol : PBS = 7 : 35 min
    ethanol5 min x 2 times
  2. Wash once with ethanol : Histo-Clear (National Diagnostics) = 1 : 1.
  3. Wash once with Histo-Clear.
  4. Add drops of Mount-Quick onto the embryos and cover with a coverslip.
  5. Leave the slide at 40oC for several hours.
(Note : Hybridization signals by this method tend to be weaker than those by other method and to diffuse, but preservation of morphology is better than other methods.)

VI.C. Glycerol mount

  1. Add drops of 90% glycerol, 10mM Tris, 1% n-propylgallate onto the embryos.
  2. Cover with a coverslip which are dotted with vaselin : solid paraffin = 9 : 1 at the 4 corners as spacer.

E. Reagents

1M MgSO41ml
Add DW to total 1 liter and autoClave
1M K-PO4 (pH6)100ml
cholesterol (5 mg/ml in EtOH)2ml
Add DW to total 2 liter and autoClave
2 x alkali-bleach solution
5M KOH2.5ml
Adjust pH to 7.2 and autoClave
PBS + 0.1%Tween 20
EH buffer (Embryo Handling buffer)
Hepes pH 7.250mM
Basal EH buffer
(= EH buffer without EGTA, NH4NO3, gelatin and DTT).
Glycine in PBT
Glycine 2 mg/ml in PBS
AutoClave, then add 0.1% Tween 20
3.7% Formaldehyde in hepes-PBS
hepes buffer* : formalin : 10 x PBS = 8 : 1 : 1
*hepes buffer
Add NaOH to pH6.9 and autoClave
Hybridization solution
deionized formamide50%
SSC (pH7, autoClaved)5x
sonicated salmon testis DNA100 /ml
yeast tRNA100 /ml
heparin100 /ml
Tween 200.1%
yatalase (15 mg/ml) and chitinase (1mg protein/ml = 5 mg crude/ml)
  1. Dissolve powder of yatalase (TAKARA No.T017) or chitinase (SIGMA No. C-6137) in 0.3M mannitol, 50mM Hepes pH 7.2, 10mM NaCl, 10mM MgCl2, 2mM DTT
  2. Filtrate through a 0.45 syringe filter.
  3. store at -20oC.
(Roche 1570013)
0.1% Tween-20 in PBS (0.01% DEPC treated)
0.1% BSA (Fraction V), 0.1% Triton X-100 in PBS
proteinase K stock solution
20 mg Proteinase K (Roche 30U/mg)/ml water
Staining buffer (Alkaline phosphatase reaction buffer)
100mMTrisHCl pH 9.5
poly-L-lysine coated slides
  1. Immerse glass slides in solution of kitchen detergent for 20 min.
  2. Wash in tap water for 1hr.
  3. Wash in ion-exchanged water.
  4. Autoclave and dry at 80oC.
  5. Drop poly-L-lysine solution (SIGMA P-8920) onto individual wells of the slides.
  6. Leave for 25 min.
  7. Aspirate off the excess solution (only when 3-well slides are used).
  8. Dry up at 65oC for 1 hr.
Parafilm coverslips
  1. Dribble beads of rubber cement along the edge of square pieces of parafilm.
  2. Dry briefly at 35-40oC.