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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;


Table of Contents

1. Preparation of DIG-labeled DNA probes for in situ hybridization
1.1. DIG-labeling by linear PCR
1.2. Chopping the probes by DNaseI digestion
2. Fixation of embryos from a small number of worms
3. Large scale fixation of embryos
3.1. Harvesting of embryos
3.2. Fixation
4. Hybridization and signal detection
4.1. Proteinase K treatment
4.2. Pre-Hybridization
4.3. Hybridization
4.4. Washing
4.5. Probe detection
4.6. Mounting
5. Reagents

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

1.1. DIG-labeling by linear PCR

  1. Make the following reaction mix (total 10 μl):

    distilled water 4.4 μl
    anchored oligo dT primers* 3.0 μl
    10 × Taq buffer 1 μl
    Taq polymerase 0.1 μl
    10 × DIG-dUTP/dNTP mix** 1.5 μl

    cDNA insert (> 3.5 ng/ μl)

    amplified from yk clone using T7/T3 primers

    0.6 μl

    * anchored oligo dT primers:

    5.3 μM (dT)17dG/5.3 μM (dT)17dC/21.3 μM (dT)17dA (This is to avoid the effect of poly-A stretch. Other vector primers may be used.)

    ** 10 × DIG-dUTP/dNTP mix:

    0.35 mM DIG-dUTP/0.65 mM dTTP/1mM each d(A, G, C)TP

     
  2. Subject to thermal cycling:

    • hot start at 95 °C for 45 sec

    • 95 °C × 15 sec

    • 45 °C × 1 min

    • 72 °C × 1 min

    • 50 cycles

  3. Add 10 μl of 10 mM EDTA

  4. Pass through G-50 spin colum chromatography (ca. 250 μl bed volume)

  5. Add 5 μl of TSE

1.2. Chopping the probes by DNaseI digestion

  1. Make the following reaction mix (total 25 μl) on ice:

    The G-50 elutate 20 μl
    10 mg/ml Salmon sperm DNA 1 μl
    distilled water 0.5 μl
    10 × DNase buffer* 2.5 μl
    DNaseI (16 μg/ml)** 1 μl

    * 10 × DNase buffer: 0.5 M TrisHCl pH 7.5, 0.1 M MgCl2

    ** Dilute stock solution (1 mg/ml) with 0.1 M 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 37 °C for 30 min

  3. Transfer on ice

  4. Add 5 μl of 0.1 M EDTA

  5. Heat at 75 °C for 5 min

  6. Check the size by alkaline agarose gel electrophoresis and DIG detection, if necessary

  7. Store frozen

2. Fixation of embryos from a small number of worms

  1. Take siliconized 1.5 ml eppendorf tubes

  2. Place about 100 μl 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 μl of suspension of E. coli OP50 in S-basal

    2. Cover the lid with the body of the tube

    3. Let stand at 20 °C overnight

  4. Spin down the worms.

  5. Add equal volume of 2 × alkaline-bleach solution and mix well

  6. Leave at r.t. for 10 min to dissolve the adult bodies

  7. Add 1 ml of M9 buffer at 4 °C

  8. Centrifuge at 2500 rpm for 30 sec at 4 °C in a swing rotor

  9. Remove the sup carefully, leaving about 100 μl of the sup to avoid removing the embryos

  10. Repeat steps 7-9 three more times

  11. Add an 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 4 °C

  14. Reduce the volume to about 50 μl

  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 × 40 mm)

  19. Place it on the top of a dry ice block

  20. Freeze for 7 min at -70 °C

  21. Peel off the cover slip quickly

  22. Soak the slide in methanol cooled at -20 °C for 5 min

  23. Rehydrate by soaking the slide in the series of the following solutions pre-cooled at 4 °C:

    methanol for 5 min
    methanol:formaldehyde-Hepes-PBS* = 35:15 for 2 min
    methanol:formaldehyde-Hepes-PBS* = 25:25 for 2 min
    methanol:formaldehyde-Hepes-PBS* = 15:35 for 2 min
    formaldehyde-Hepes-PBS* for 20 min

    *formaldehyde-Hepes-PBS  
    Hepes 200ml
    10xPBS 25ml
    formaldehyde 25ml

  24. Dehydrate by soaking the slide in the series of the following solutions at r.t.:

    ethanol:PBS=15:35 for 5min
    ethanol:PBS=25:25 for 5min
    ethanol:PBS=35:15 for 5 min
    ethanol for 5 min × 2 times
  25. Store in ethanol at −20 °C.

3. Large scale fixation of embryos

3.1. 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 μm 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 4 ml water in a 15 ml Falcon tube (clear type)

  7. Add 5 ml of 2 × alkali-bleach solution, mix well and let stand for 10 min

  8. Force the worms out through a 23-gauge needle onto nylon mesh

  9. Collect embryos by spinning the filtrate at 800 × g using a swing rotor

  10. Wash the embryos 4 times with M9 and transferred into a siliconized eppendorf tube

3.2. Fixation

  1. Take 100 μl (packed volume) of the embryos and adjust the volume to 200 μl with M9

  2. Add 200 μl of yatalase (15 mg/ml in 0.3 M 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 μl/well of Basal EH buffer onto each well of poly-L-lysine coated 8-well slides

  7. Dispense 5 μl/well of the embryo suspension into the buffer at each well

  8. Let stand for 10 min at 4 °C to settle the embryos to the bottom

  9. Remove the buffer, and immediately immerse in methanol at -20 °C for 5 min

  10. Rehydrate the embryos by immersing the slides in the follwoing series at 4 °C. The solutions must be pre-cooled at 4 °C.

    methanol 5 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 22 °C
  11. Dehydrate the embryos by immersing the slides in the following series at r.t.

    ethanol:PBS = 3:7 5 min
    ethanol:PBS = 1:1 5 min
    ethanol:PBS = 7:3 5 min
    ethanol 5 min × 2 times
  12. Store in ethanol at -20 °C.

4. Hybridization and signal detection

4.1. 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 = 1:1 5 min
    ethanol:PBS = 3:7 5 min
  2. Wash the slides once by immersing in PBT for 5 min. *For late stage embryos, additional HCl treatment is effective, which can cut glycosid bonds of the proteoglycan that appear on late stage embryos.

    1. a) Immerse the slides in 0.2 N HCl for 20 min at r.t.

    2. b) Wash the slides 2 times in PBT for 5 min

  3. Immerse the slides in proteinase K (10 μg/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

4.2. Pre-Hybridization

  1. Immerse the slides in the following series of mixtures;

    50% formamide, 5 × SSC, heparin, 0.1% Tween:PBT = 1:1 10 min
    50% formamide, 5 × SSC, 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 μl of heat denatured (at 99 °C 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, 5 × SSC. (No need to use coverslips)

  6. Incubate at 48 °C for 1 hr

4.3. Hybridization

  1. Add 50 μl of heat-denatured DNA probes for each slide. (The final concentrations of probes is about 0.06 μg/ml)

  2. Cover the slide with a parafilm coverslip to reduce evaporation

  3. Incubate the slides at 48 °C overnight in the moist chamber

4.4. Washing

  1. Wash the slides in the following series of washing solutins at 48 °C with slight agitation

    50% formamide, 5 × SSC, heparin, 0.1% Tween:PBT = 1:1 (First washing is performed in separate containers for every 2 min slides)  
    50% formamide, 5 × SSC, heparin, 0.1% Tween:PBT = 1:1 10 min × 2 times
    0.8 × PBS, 0.1% CHAPS 20 min × 4 times
  2. Wash the slides twice in PBT for 5 min at r.t. to remove CHAPS.

4.5. 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 μl 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 reagents) twice for 5 min each at r.t.

  6. Color development

    1. a) Mix 180 μl of NBT and 140 μl of BCIP in 40 ml of staining buffer.

    2. b) Immerse the slides in the mixture for 1 hr at 22 °C in the dark, monitoring the extent of the staining.

  7. Wash the slides three times with PBS, 20 mM EDTA to stop the reaction.

  8. If necessary, incubate the slides in 1 μg/ml DAPI in Tris buffer at 4 °C for 30 min.

4.6. Mounting

4.6.1. Permanent mount 1

  1. Add about 90 μl 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

4.6.2. Permanent mount 2

  1. Dehydrate with the following ethanol series:

    ethanol:PBS = 3:7 5 min
    ethanol:PBS = 1:1 5 min
    ethanol:PBS = 7:3 5 min
    ethanol 5 min × 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 40 °C for several hours

(Note : Hybridization signals by this method tend to be weaker than those by other methods and to diffuse, but preservation of morphology is better than other methods)

4.6.3. Glycerol mount

  1. Add drops of 90% glycerol, 10 mM 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

5. Reagents

M9

KH2PO4 3 g
Na2HPO4 6 g
1M MgSO4 1 ml
Add DW to total 1 liter and autoclave  

S-basal

NaCl 69 g
1M K-PO4 (pH6) 100 ml
cholesterol (5 mg/ml in EtOH) 2 ml
Add DW to total 2 liter and autoclave

2 × alkali-bleach solution

NaClO 3.0 ml
5M KOH 2.5 ml
DW 19.5 ml

PBS

NaCl 13 7 mM
KCl 2.7 mM
Na2HPO4 4.3 mM
KH2PO4 1.5 mM
Adjust pH to 7.2 and autoclave

PBT

  • PBS + 0.1% Tween 20

EH buffer (Embryo Handling buffer)

mannitol 0.3 M
Hepes pH 7.2 50 mM
NaCl 10 mM
MgCl2 10 mM
EGTA 0.04%
NH4NO3 2 mM
gelatin 0.1%
DTT 2 mM

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 × PBS = 8:1:1

*hepes buffer

Hepes 100 mM
MgSO4 2 mM
EGTA 0.04%
Add NaOH to pH6.9 and autoclave

Hybridization solution

deionized formamide 50%
SSC (pH7, autoclaved)
sonicated salmon testis DNA 100 μg/ml
yeast tRNA 100 μg/ml
heparin 100 μg/ml
Tween 20 0.1%

yatalase (15 mg/ml) and chitinase (1 mg protein/ml = 5 mg crude/ml)

  1. Dissolve powder of yatalase (TAKARA No.T017) or chitinase (SIGMA No. C-6137) in 0.3 M mannitol, 50 mM Hepes pH 7.2, 10 mM NaCl, 10 mM MgCl2, 2 mM DTT

  2. Filtrate through a 0.45 μm syringe filter.

  3. store at -20 °C.

Digoxigenin-11-dUTP (Roche 1570013)

PBT

  • 0.1% Tween-20 in PBS (0.01% DEPC treated)

PBtr

  • 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)

NaCl 100 mM
MgCl2 5 mM
TrisHCl pH 9.5 100 mM
Levamisol 1 mM
Tween-20 0.1%

poly-L-lysine coated slides

  1. Immerse glass slides in solution of kitchen detergent for 20 min

  2. Wash in tap water for 1 hr

  3. Wash in ion-exchanged water

  4. Autoclave and dry at 80 °C

  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 at 65 °C for 1 hr

Parafilm coverslips

  1. Dribble beads of rubber cement along the edge of square pieces of parafilm

  2. Dry briefly at 35-40 °C



*Edited by Ikuko Muramatsu, Masumi Obara, Wakako Shimizu, Aya Hamakawa, and Yuji Kohara. Last revised April 27, 2005. Published May 19, 2015. This chapter should be cited as: Motohashi T., Hirono K., and Kohara Y. In situ hybridization on whole mount embryos of C. elegans (May 19, 2015), WormBook, ed. The C. elegans Research Community, WormBook, doi/10.1895/wormbook.1.176.1, http://www.wormbook.org.

Copyright: © 2015 Tomoko Motohashi, Keiko Hirono and Yuji Kohara. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

§To whom correspondence should be addressed: Email: Ikuko Muramatsu, isugiura@lab.nig.ac.jp, Yuji Kohara, ykohara@lab.nig.ac.jp

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