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For all these fixatives the most common protocol is to fix, then rinse in TBS, exchange TBS for 100% methanol and store samples in the freezer, where they will be stable for many months (years?). Methanol is itself a fixative, and also increases the porosity of the samples, giving antibodies better access to internal structures. It can however block some epitopes, so if your samples don't stain try skipping the methanol. Our normal fix. Good for most antibodies, and for lacZ enzymatic activity. Fix in neutral buffered formalin (=4% formaldehyde final, stabilized with methanol) for up to two hours at room temperature. Rinse with TBS and store in 100% methanol. 0.1M MOPS pH 7.4 (with NaOH) + 2 mM EGTA + 1 mM MgSO4 + 4% formaledhyde (1/10 vol of concentrated stock, 37 or 40%) make up 10 x salts with first three reagents, autoclave and store at room temp. It will yellow with exposure to light but this is O.K. Add formaldehyde fresh, from a bottle that has been opened less than one month. Paraformaldehyde can be substituted if formaldehyde. Store paraformaldehyde at 4 degC for two weeks. Fix at room temp for up to 2 hours. 4 vols Methanol plus one volume DMSO. Fix in freezer at -20 degC overnight Fix at room temp for 30' to 2 hours, wash with a number of changes of 70% ethanol. Some people say you must wash out all traces of the yellow stain if you plan to do parafin sections on Bouin's fixed embryos. I'm no gun histologist so I won't comment. Fix at room temp for up to three hours, transfer to 100 % ethanol. Powerful well penetrating fix, preserves nucleic acids. Stable at room temperature. Good for performing dissections for RNA isolation. Add embryos to fix just before dissection. Collect bits, remove excess fix, rinse with 0.3M tris pH 7.5, homogenize in RNA isolation buffer.