ß-Galactosidase Activity Staining using X-gal
The protocol from Y. Hiromi contains several staining solutions that allow adjustment to
the staining intensity in particular genotypes. The use of cacodylate as buffer in the
fixative and DMF as solvent for X-gal can be omitted and replaced with less toxic
solutions; PBS as buffer during fixation and DMSO as solvent for X-gal.
However, the more recent protocol by Ch. Klämbt appears much easier to use (no/or
chemical devitellinization of embryos;) and is certainly the one of choice when screening
large numbers of different genotypes. It also allows counter staining with antibodies
following the ß-gal activity reaction.
Protocol from Yash Hiromi
1) Dechorionate embryos ;with 50% bleach. Wash well with water and collect on a
Nitex mesh. Blot mesh on tissue paper to remove excess water.
2) In a deep depression slide, fix embryos in 0.5 ml of heptane which is saturated with the fixative for 15 minutes at room temperature. Cover the wells with a glass slide to prevent heptane from evaporating. Embryos should turn light yellow. Whitish embryos are not well fixed and will not stain well.
3) Take embryos with a Pasteur pipette and transfer onto a slide glass. Use a piece of filter paper to remove excess heptane. After all the heptane has evaporated, take embryos by gently touching with a double stick tape and stick the tape on a slide glass, embryo side up (Siliconized slides are easier to work with.) Cover embryos with a drop of PBS.
4) Devitellinize using a dissection needle (a tungsten needle is not necessary). You can use a sewing needle whose tip is sharpened with a sand paper. Well fixed embryos should be stiff and should come out from the membrane by gently scratching the vitelline membrane. Transfer the embryos into an Eppendorf tube.
5) Remove the PBS and add ~300 ml of staining solution without X-gal (i.e. Fe/NaP or Fe/CP). Leave at room temperature for ca. 5 minutes.
6) Remove liquid and replace with staining solution. Incubate from 2 hours to overnight at room temperature or 37° C.
7) Remove the staining solution. Rinse embryos by vortexing once in 70% ethanol and once in 100% ethanol.
8) Store the embryos in 90% glycerol/PBS at 4° C. They can be kept for months at this stage. Mount using two #1 coverslips as spacer.
Comments:
a) Staining with Fe/NaP pH 7.2 and higher temperature (37° C) results in stronger
staining, presumably because the conditions are closer to the optimum for the
enzymatic activity. For ftz/lacZ fusion gene transformants, staining using Fe/CP
overnight at room temperature is sufficient for visualizing stripes in the extended
germband stage embryos and for CNS staining. To stain stripes at the blastoderm
stage, use Fe/NaP pH 7.2 and stain overnight at 37° C. In this case, do not include
older embryos in the same tube, because they will stain too strongly and may
transfer the blue reaction product to the other embryos in contact. For best
resolution, try to use conditions that allows you to stain overnight, rather than
stopping the reaction after a short incubation period. Staining at 37° C will increase
the size of reaction product crystals and will make identification of stained cells
more difficult.
b) White precipitate (X-gal crystals; ?) will form in the staining solution during incubation. This does not affect staining reaction, but will stick to the embryos. Vortexing in ethanol helps to remove it from the embryos.
c) Embryos can also be dehydrated and mounted in Epon for permanent preparations. GMM can not be used because blue dye appears to dissolve (although slowly) in this medium.
Solutions:
Fixative (prepare fresh solution every day):
0.1 M Na cacodylate buffer pH 7.3 1 ml
50% glutaraldehyde
(EM grade Fluca #49631) 1 ml
heptane 2 ml
Shake well and let the phases separate. Use the heptane (upper) phase for fixation. Fixative (lower) phase can be reused by adding heptane to 2 ml.
Fe/NaP-solution (pH 7.2):
0.2 M Na2HPO4 1.8 ml
0.2 M NaH2PO4 0.7 ml
5 M NaCl 1.5 ml
1 M MgCl2 50 ml
50 mM K3(Fe(CN)6) 3.05 ml
50 mM K4(Fe(CN)6) 3.05 ml
H2O to 50 ml
Store in dark at room temperature.
Fe/CP-solution (pH 8):
50 mM K3(Fe(CN)6) 5 ml
50 mM K4(Fe(CN)6) 5 ml
CP pH 8 40 ml
Store in dark at room temperature.
CP-solution (pH 8):
0.1 M citric acid 27.5 ml
Na2HPO4 27.61 g
H2O to 1 l
Staining solution:
Warm up Fe/NaP or Fe/CP to 37° C. Add 1/30 volume
of X-gal (5-bromo-4-chloro-3-indolyl-b-D-galactopyranoside)
solution (8% in DMSO, store at -20° C) . Use 200 to 300 ml per tube
of embryos.
Modification for weak ß-Galactosidase Expression
With weak ß-galactosidase expression, long incubation of glutaraldehyde fixed embryos
in staining solution results in an undesirably yellow embryo. It is possible to avoid this
with the following modification
1) Fix dechorionated embryos for 20 minutes in a monolayer in heptane previously equilibrated with 18% formaldehyde in 0.1 M cacodylate buffer, pH 7.4.
2) As in other X-gal protocols, dry embryos and mount on double sided tape, inside a silicone sealant well on a slide.
3) Wash with staining buffer, then incubate for up to 3 days at 30° C in staining solution with Xgal. Refresh solutions half way through.
4) Wash out buffer and replace with fixative (for example 4% formaldehyde in PBS).
Poke a hole in the vitelline, let fixative penetrate for at least 10 minutes, and then
devitellinate embryos. Store in 80% glycerol/20% PBS until photographed.
Protocol from Christian Klämbt.
All steps are performed in custom made multi well plates unless otherwise noted.
I. Preparation of embryos still in their vitelline membrane:
1) Collect embryos and wash with PBT.
2) Dechorionate in 50% bleach for 4 minutes and wash for 2 to 5 minutes with PBT.
3) Blot the embryos dry and place them in n-heptane saturated with 2.5% glutaraldehyde/PBS. The PBS/glutaraldehyde solution can be kept and used for the saturation of fresh n-heptane for a long time (up to 8 months at 4° C).
4) Fix for 7 minutes while gentle agitating at RT.
5) Blot away excess heptane and wash embryos with PBT until they sink to the bottom and do not stick to each other any more.
6) Wash and permeabilize for another 2 hours with several changes of PBT (Even slight contamination with heptane on the vitelline membrane will cause diffuse blue background staining.)
II. Staining:
1) Heat the staining solution to 65° C, or over a flame until the solution gets cloudy (This will greatly reduce the formation of crystals during the staining process). Add 1/50 volume of 10% X-gal solution in DMSO to the warm staining solution.
2) Blot away excess PBT and place embryos in the staining solution.
3) Stain for 2 to 4 hours at 37° C in a humid chamber and stop the reaction by washing with PBT.
III. Preparation of devitellinized embryos for X-gal staining:
Methanol and also ethanol will destroy lac-Z activity. However, if the devitellinization
step is done fast, the methanol as well as the ethanol do not significantly destroy the
enzyme's activity.
1) Wash, dechorionate and fix the embryos as in (I.).
2) Blot away excess heptane fix and wash the embroys in n-heptane.
3) Transfer the embryos in 1 to 2 ml heptane into a 5 ml glass tube, add 1 to 2 ml
methanol.and shake vigorously.
4.) After 10 to 20 seconds the interphase becomes visible. Aspirate the heptane phase
and half of the methanol phase off (at this point not all the embryos will have made
it to the bottom of the glass tube. You will lose 30 to 50% of the embryos).
5) Add 2 ml methanol for a wash. Aspirate all methanol and wash twice with 80%
ethanol. Do not let the embryos sit in methanol or ethanol for longer times than
neccessary.
6) Wash 3 times for 5 to 10 minutes with 4 ml PBT on a rocking platform.and wash for an additional 2 hours in PBT.
7) Transfer the embryos back into a multi well plate and proceed with the staining as in
(II.). Do not let the embryos dry at this point. If you want to look at the X-gal stained
embryos, wash briefly in PBT and mount them in PBS/50% glycerol.
If counter staining of the embryos with specific antibodies is desired, wash away
staining solution with several changes of PBT for 2 hours. Subsequently wash embryos
with PBT containing BSA and 5-10% goatserum for 1 h and proceed with the usual
staining protocol for antibodies.
Solutions:
Staining solution (200 ml):
10 mM PO4-buffer pH 7.2 2.0 ml 1M
150 mM NaCl 6.0 ml 5M
1 mM MgCl2 0.2 ml 1M
3 mM K4[FeII(CN)6] 0.3 g
3 mM K3[FeIII(CN)6] 0.2 g
0.3 % Triton X-100 0.6 ml
PBT: PBS with 0.5 % Triton X-100 (but without BSA)
Visualization of ß-Galactosidase for EM Histology ;(Protocol from Roger Jacobs)
1) Process for X-gal staining as described above for light microscopy. After staining,
transfer to 0.1 M cacodylate buffer, pH 7.4, and select the embryos you wish to
embed.
2) Fix in primary EM fix (2% glutaraldehyde, 2% paraformaladehyde in 0.1 M cacodylate buffer, pH 7.4) for 30 minutes.
5) Wash in cacodylate buffer with at least 3 changes for 5 minutes.
6) Fix in 1% osmium tetroxide in cacodylate buffer for 30 minutes
7) Wash in cacodylate buffer, followed by distilled water.
8) Stain in 5 % uranyl acetate for 30 minutes.
9) Dehydrate and infiltrate as in standard EM protocol. Older whole-mount embryos need longer infiltration times. Infiltrate in pure plastic for 24 to 48 hours (Note dissections infiltrate through a graded ethanol-plastic series!). Use for example Epon-Araldite plastic. You may also use methacrylate based plastics, but this will make semi-thin sectioning (0.5 µm) more difficult. It is much easier to embed whole-mount embryos in the middle rather than the bottom of the block. Cook plastic in half filled molds for about 4 hours, then add embryos in new plastic to the top half of the mold. You can orient the embryos in the mold with a fine probe.
Sectioning and Staining:
Trim and orient your tissue by conventional technique. Sections of 1.0 µm or thicker can
be picked up individually off a dry glass knife and placed on a small drop of water on a
subbed (1% gelatin and 0.1% chromium potassium sulphate x 12 H2O) slide. For serial
sectioning, use a glass knife with a small water boat behind it [made with mylar tape, or
LKB's prefab plastic boats (#2208-100)]. 0.5µm is the easiest to handle. Ribbons of
sections can be spread with a chloroform wick, and transferred to a dry subbed slide
using the wire loop technique. Merely touch the loop containing the ribbon and water to
the slide, and remove the water by touching a sliver of filter paper to the edge of the
loop.
Best tissue contrast is obtained with 0.5% Toluidine Blue O in 1% Borax buffer.
This stain is preferred when you are trimming and orienting, but will not allow you to
visualize X-gal product.
To preserve contrast with X-gal, counterstain with saturated aqueous Basic
Fuchsin. This stain penetrates epon slowly, but there are two tricks which will increase
permeability. First, you might soak your unstained slide in xylene for 20 to 30 minutes,
wash in acetone, and dry before staining. Secondly, add a drop or more of acetone
containing 1% Basic Fuchsin to the slide when staining with saturated aqueous Basic
Fuchsin on a hotplate (65° C).
Slides should be permanently mounted with permount and coverslip, an viewed
under oil immersion optics. You can now enhance X-gal staining contrast by using
appropriate filters (for example a yellow filter or in extreme cases, a red filter will
darken the image of the crystals relative to the rest of the tissue).
In EM, X-gal crystals make large ugly needles that actually penetrate cell walls,
and fill vacuoles inside cells. It is therefore useful mostly in low magnification work.
Immunohistochemical Detection of ß-Galactosidase
The detection of ß-galactosidase with an antibody and immunohistochemical staining
gives the best possible single cell resolution. However, it is more laborious than activity
staining.
I usually follow the standard protocol for immunohistochemistry in embryos as described in the relevant section of this methods book. The following reagents and dilutions are normally employed:
primary antibody: anti-ß-gal monoclonal from Promega
use at 1/200 to 1/500 dilution in PBT(PBS, BSA,
Triton; for recipe see immunohistochemistry protocol) cont.
5% goat serum
(The quality of the anti-ß-gal monoclonal is obviously important. However, we have sometimes had problems obtaining consistent results due to lower quality of the primary antibody. Nevertheless, Promega still provides the best monoclonal commercially available.)
secondary antibody HRP-conjugated goat-anti-mouse polyclonal from Biorad
use at 1/200 dilution in PBT/goat serum (see above)
HRP reaction 0.5 mg/ml DAB in PBT
add 1µl of 3% Hydrogenperoxide and 8µl 8% NiCl(2)
7H(2)O for each ml of DAB solution
Embryo pretreatment, incubations, washes, staining reaction and mounting of the
embryos is performed as described by the standard immunohistochemistry protocol.