We would like help with methods people have used to break open culture
cells (that contain a lot of chloroplasts) to isolate plasma membranes
using 2-phase partitioning. Our moss cultures consist of small filaments with
2-5 cells. We have tried homogenizing them with a mortar and pestle, an
omnimixer, a tissue tearer, and freezing the tissue and then grinding;
none of these methods allow us to break open the cells.
We have recently tried a bead beater and can disrupt the cells, but our
microsomal preps are loaded with chloroplasts or isolated thylakoids (in
other words they are sticky and green). We have tried using fatty acid
free BSA in our homogenization buffer to prevent sticking of chloroplasts
etc to our membranes, but don't see any improvement. When we load these
resuspended microsomes on our 2-phase gradients for plasma membrane
isolation, the phases don't separate well (overloaded?) and we don't get
good recovery of clean pm (even if we add additional gradients). We have
been able to use 2-phase partitioning to isolate pm from vegetative tissue;
however, this tissue can be ground with a mortar and pestle and contains
fewer chloroplasts.
Can anyone help us with methods successfully used to isolate plasma
membranes from green culture cells? We are most interested in knowing
how the cells are broken open and any methods used to remove
chloroplasts/pieces of chloroplasts. I would be happy to provide any
>additional information. Thanks.
Karen Schumaker
University of Arizona
Department of Plant Sciences
schumake at ag.arizona.edu
Phone: (520) 621-9635
Fax: 520-621-2012
