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Protocols2 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.

  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 4C 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.

  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 of OCT is placed over the culture, then the tube placed in liquid nitrogen. It is convenient to keep the tissue near the top of the cryotube. Liquid nitrogen-cooled isopentane may also be used for freezing. Tissue may be kept frozen for several weeks.
  2. Precoat cleaned glass microscope slides with poly-L-lysine at 100 mg/ml.
  3. Mount the tissue on a stub using OCT as adhesive.
  4. Cut 7 mm sections using a cryostat. Air dry the sections then place in a tray or box, wrap in foil and store at -80C. Sections should be used as soon as possible and certainly within 2 months.
  5. Allow sections to warm to room temperature then fix with dry acetone for 10 min. Rehydrate sections using PBS then incubate in protein blocking solution (Dako; 10 min).
  6. Standard immunostaining protocols may be used with peroxidase or fluorescent conjugates (14). Note that alkaline phosphatase conjugates should be used with caution as this enzyme is expressed by some trophoblast.

Resin embedding of explant cutures.

Examination of semithin resin sections can show the pattern of cell growth across and within the gel.
Electron microscopy enables cell-cell and cell-matrix interactions to be visualised.

Equipment and reagents

  1. Rinse the explant culture in PBS, then fix in freshly prepared 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer pH 7.3 for 2-3 hours at room temperature.
  2. Rinse in 0.1 M sodium cacodylate buffer pH 7.3 with 3 mM CaCl2 three times over 24 h and store at 4C until ready for processing. Then, using a dissecting microscope and razor blade, trim the gel around the explant into a rectangular shape about 4 x 8 mm, ensuring that one short edge is cut at right angles across the explant growth. This will be the face that is sectioned and the maximum area of outgrowth possible is desirable. Transfer to a glass vial for processing.
  3. Post-fix in 1% osmium tetroxide in 0.05 M sodium cacodylate buffer pH 7.3 (equal parts 2% aqueous OsO4 solution and 0.1 M buffer) at 4C for 1 h in a closed glass vial. Prepare in a fume cupboard and wear protective gloves. Do not inhale vapour. After fixation, decant the fix and rinse in 0.1 M sodium cacodylate buffer pH 7.3.
  4. Dehydrate in an ascending alcohol series: 15 min each in 50, 70, 95, 100, 100% ethanol.
  5. Incubate in propylene oxide 2 x 15 min, in a fume cupboard, wearing protective gloves.
  6. Infiltrate in equal parts epoxy resin and propylene oxide for 1 h (Epon, Araldite, Taab Embedding Resin or other epoxy resin mixtures can be used). This must be done in a fume cupboard, wearing gloves. Do not inhale vapour. Follow the manufacturer's instructions for the preparation of resin.
  7. Leave overnight in closed vials at 4C on a rotator in a mixture of three parts resin and 1 part propylene oxide.
  8. Next day, give three changes of resin at 48C and then embed in rectangular silicone rubber moulds approximately 6 x 11 x 5 mm deep. Place the gel so that the cut surface of the explant is up against the 6 mm edge of the mould, which will be the cutting face. Fill the moulds to the top with resin. Polymerise at 60C for 72 hours.
  9. With a razor blade or using an ultramicrotome trimming facility, trim into the block face until the outgrowth is reached. This can be seen as a fine black line, and the cut villus surface as a round profile. Then cut 0.5 m thick sections and mount on a glass slide. Leave on a hotplate (70-80C) to dry and stain with 1% toluidine blue in 1% borax. Wash and examine.
  10. Suitable areas can be photographed and then ultrathin sections cut with a glass or diamond knife and mounted onto copper grids, contrasted with uranyl acetate/ lead citrate and examined in the electron microscope.