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New fixation WBG 14-3

C. elegans whole mount immunohistochemistry with Bouin's Fixative

I have developed a fixation protocol which provides an alternative set of conditions to test antisera that work poorly on paraformaldehyde fixed worms (e.g. the Finney protocol). A Bouin's fixative of 75 ml saturated picric acid [real nasty stuff- explosive in crystal form!], 25 ml of formalin, and 5 ml of glacial acetic acid is made (many fly labs use this fixation solution which may be stored at 4°C for many months). Nematodes grown on NGM plates (usually from chunks for 2 days at 22.5 °C) are washed off the plate using 2 to 3 ml of H2O and spun down in a glass 12 ml conical tube in a clinical centrifuge at RT (approximately 23°C in my lab). The supernatant, save 50 µl, is removed by aspiration. A mix of 400 µl of Bouin's, 400 µl methanol and 10 µl of ßME is placed in the glass tube to initiate fixation and subsequently the worms are transferred to an eppendorf tube using a glass pipette (600:200:10 Bouin's: MeOH: ßME also works). The tube is rocked on a Labquake shaker at RT for 30 minutes, then quick frozen in liquid N2. The tube is quickly thawed under running hot water until the solution melts (but before it warms up past RT). The tube is rocked an additional 30 minutes. A solution of BTB [ 1X Borate buffer, 0.5% Triton X-100, 2% ßME] is made from 50X borate buffer ( 1.0 M H3BO3, 0.5 M NaOH). The worms are spun from fixative in a centrifuge [ 2-3 seconds in an eppendorf 5415C ]. The fixative is aspirated, and 1.4 ml of wash solution is added, rocked a few seconds to mix. Spinning and aspiration completes the first wash. Two more washes are done, and the worms are then resuspended again in 1.0 ml BTB ( in the lower volume worms move more during rocking). The worms still have a yellow tinge due to the picric acid at this point. The worms are now rocked for approximately 1 hr at RT. The speed at which the yellow picric acid destains is a good sign of how well the worms are going to stain. If the worms remain yellow after an hr of washing, they usually end up impermeable to antibodies. Wash again with BTB and incubate an additional 2 or 3 hrs. Wash with BT (BTB lacking BME) one time, and AbA (1X PBS, 1.0% BSA, 0.5% Triton X-100, 10 mM NaAzide) solution two times. Incubate 30 minutes or more then begin standard Ab incubations. I never switch tubes during this procedure. I store these worms at 4°C and they often stain relatively effectively a month later (and sometimes they don't).

In my hands this procedure gives good staining in greater than 90% of the worms. Our antibodies to RAB-3 and SNB-1(synaptobrevin/VAMP) work much better on Bouin-fixed worms than on well permeabilized [Finney protocol]-fixed worms. UNC-64 (syntaxin) Ab works about the same for both protocols, and SNT-1 (synaptotagmin) Ab works much better in the Finney fixation than the Bouin's fixation, although staining is still reasonable in Bouin's. Thus, Bouin's fixation provides an alternative condition that may reveal different epitopes on proteins. I have not altered fixation length very much, but I have found that increased fixation does reduce permeability. A 1 hr fixation seems to be a good compromise between fixation length and retaining permeability, but I would suggest reducing fixation to 30 minutes if you find your worms are not permeable when you try the procedure. I am relatively certain that small changes in the temperature during fixation can cause dramatic changes in the extent of fixation.

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