Corrected by Ikuko Muramatsu, Masumi Obara, Wakako Shimizu, Aya Hamakawa and Yuji Kohara (April 27, 2005)
Correspondence: Ikuko Muramatsu <isugiura@lab.nig.ac.jp>, Yuji Kohara <ykohara@lab.nig.ac.jp>

Contents:

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 μl);

   distilled water	4.4 μl
   anchored oligo dT primers *	3.0 μl
   10 x Taq buffer	1 μl
   Taq polymerase	0.1 μl
   10 x 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 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 μl of 10mM EDTA.
4. Pass through G-50 spin colum chromatography (ca. 250 μl bed volume).
5. Add 5 μl of TSE.

II. Chopping the probes by DNaseI digestion

6. Make the following reaction mix (total 25 μl) on ice;

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

   * 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.)
7. Incubate at 37^oC for 30 min.
8. Transfer on ice.
9. Add 5 μl of 0.1M EDTA.
10. Heat at 75^oC for 5 min.
11. Check the size by alkaline agarose gel electrophoresis and DIG detection, if necessary.
12. Store frozen.

B. Fixation of embryos from a small number of worms

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 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 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 μl (packed volume) of the embryos and adjust the volume to 200 μl with M9.
2. Add 200 μl 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 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^oC to settle the embryos to the bottom.
9. Remove the buffer, and immediately immerse in methanol at -20^oC for 5 min.
10. Rehydrate the embryos by immersing the slides in the follwoing series at 4^oC. The solutions must be pre-cooled at 4^oC.

    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 22oC

11. Dehydrate the embryos by immersing the slides in the follwoing 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 x 2 times

12. 12. Store in ethanol at -20^oC.

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 : 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, 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 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.

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 μl of heat denatrued (at 99^oC 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 48^oC for 1 hr.

III. 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^oC overnight in the moist chamber.

IV. Washing

1. Wash the slides in the following series of washing solutins at 48^oC 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 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 reagetnts) twice for 5 min each at r.t..
6. Colour development
   1. Mix 180 μl of NBT and 140 μl of BCIP in 40 ml of staining buffer.
   2. Immerse the slides in the mixture for 1 hr at 22^oC 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 μg/ml DAPI in Tris buffer at 4^oC for 30 min.

VI. Mounting

VI.A. 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.

VI.B. 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 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 40^oC for several hours.

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

M9

KH2PO4	3g
Na2HPO4	6g
1M MgSO4	1ml

Add DW to total 1 liter and autoClave

S-basal

NaCl	69g
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

NaClO	3ml
5M KOH	2.5ml
DW	19.5ml

PBS

NaCl	137mM
KCl	2.7mM
Na2HPO4	4.3mM
KH2PO4	1.5mM

Adjust pH to 7.2 and autoClave

PBT

PBS + 0.1%Tween 20

EH buffer (Embryo Handling buffer)

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

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
	Hepes	100mM
	MgSO4	2mM
	EGTA	0.04%

Add NaOH to pH6.9 and autoClave

Hybridization solution

deionized formamide	50%
SSC (pH7, autoClaved)	5x
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 (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 μm syringe filter.
3. store at -20^oC.

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)

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

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 80^oC.
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 65^oC 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^oC.

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