Staining protocols for placental explant cultures

Staining protocols for placental explant cultures

Whole mount staining.

The protocol given uses a single primary antibody with nuclear counterstaining, but it can be extended to double antibody staining. It requires an inverted microscope with fluorescence illumination and long working distance objectives.

Whole mount staining in multiwell plates.

Method

1. Wash the cultures gently and thoroughly in PBS twice.
2. Fix in 1 ml MeOH. Replace the first solution immediately with a second aliquot. Incubate 30 min.
3. Wash three times with PBS.
4. At this stage cultures can be trimmed under the dissecting microscope with a small pair of scissors. Remove parts of the explant that are not participating in anchorage to the gel. Take care not to damage the gel surface. If floating villi are removed there is less likelihood of damaging the culture during processing.
5. Incubate overnight (at least) at 4°C in blocking solution (4% BSA/PBS). Include 0.02% sodium azide if the plates are to be kept longer.
6. Incubate in 1st antibody for 30 min at room temperature. Ideally the culture should be submerged. If necessary the volume can be minimised (eg 100 ml) by repeatedly pipetting the solution over the explant, preventing it from drying out. Antibody should be diluted in 4% BSA/PBS. A good positive control antibody is anti-cytokeratin 7 (OV-TL 12/30; Dako; use at 1/50). This identifies the outgrowing cells unequivocally as trophoblast. Anticytokeratin 8/18 can be used for marking cells (e.g CAM5.2, Beckton-Dickenson) but beware that some placental fibroblasts express this marker so it is not specific for trophoblast.
7. Wash in PBS, then overnight on a platform shaker in PBS/BSA.
8. Incubate for 30 min - 2 h in fluorescent 2nd antibody diluted in PBS/BSA. Plates should be wrapped in foil.
9. Wash overnight in several changes PBS using a platform shaker. Cultures can be inspected under the inverted microscope at any stage to monitor the removal of background fluorescence in the gel. Keep the plates in the dark throughout.
10. Counterstain nuclei using eg propidium iodide or DAPI at 5 mg/ml in PBS, 30 min. PI or DAPI are made up in water as a stock solution of 5 mg/ml.
11. Wash further in PBS.

Whole mount staining in tubes

The protocol can be modified for more economical use of antibody by detaching the cultures and staining in Eppendorf tubes.

Method

1. Fix as described in the previous protocol and rehydrate in PBS.
2. Trim the gel using a scalpel retaining the central area with explant.
3. Gently tease the remaining gel off the well surface using a small spatula.
4. Use a pastette to transfer it to a 0.5 ml Eppendorf tube.
5. Add blocking solution (PBS/4% BSA) and proceed as in Protocol 3. Incubations can be carried out on a roller mixer.
6. After staining, transfer the gel fragment to a microscope slide and remove excess PBS by blotting with a tissue. Orient it as during culture.
7. Encase the gel fragment in aqueous hardening mountant (Histotec, Serotec, UK). Do not use a cover slip. Incubate at room temperature overnight in the dark.
8. The culture can now be inspected directly using water or oil immersion objectives and either upright or inverted optics.

Cryosectioning explant cultures.

For immunostaining with a larger panel of antibodies on the same culture, cryosections are required.

1. Trim the edges of the gel with a scalpel if the culture is confined to one part of the collagen drop. Gently tease the gel carrying the culture from the floor of the well and transferre to a cryotube containing a drop of OCT compound. A further drop

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