Okay gang,

We got a sticky wicket research problem here, and I thought some of you
might have
ideas to help us out.

My colleague uses DiI to retrogradely label motoneurons in manduca sexta
larva, which
enables her to identify the cells of interest in each segment.  She then
dissects out
the DiI-labeled motoneurons, which as you can imagine is a fairly laborious
process,
and now wants to do acid phosphatase histochemistry on these cells.  She
has been
trying to pellet these cells in agar and then cryosection the pellet for
the enzyme
assay.  But she has so few cells in each pellet (7-8 cells per pellet),
that she
can't find them, even though she keeps the pellet quite small.  So she
needs a more
efficient way to pellet them or needs to take a different approach.

So now she plans to make a small well on a polylysine-coated slide,
transfer the
dissected cells to the well and fix the cells with paraformaldehyde in the
well for
an hour at room temperature.  The thought is the fix will crosslink the
proteins in
the cell membrane to the polylysine, and the cells will adhere during the
subsequent
acid phosphatase assay.  I know that alkaline phosphatase activity survives
paraformaldehyde fixation, so my assumption is that acid phosphatase will also.

She can't assay the cells in place in the tissue because the assay she's
using has an
acetone step, which will leach out the DiI.  Even if she modified the
protocol to
eliminate the acetone, chromophoric substrates will quench DiI, and DiI has
such
broad excitation and emission spectra, it will interfere with any fluorophoric
substrate she might use (except maybe the ELF substrate, which is another
whole can
of worms).

So if you have any thoughts on her current approach or any other ideas she
might try,
we're all ears.

Karen in Oregon