I would like some help with formulating a new protocol for a rough fractioning of the organel content of some cell culture samples (fibroblasts).
What I currently am doing:
Basically freeze/thaw cycles and a brief waterbath sonification of the whole cell.
What I would like to do:
Harvest cells and lyse them with a hypo osmotic lysis buffer (possibly with some nonionic detergent (and protease inhibitors)). It's essential that the mitochondria should be largely unaffected by this.
Alternatively I have read that 30 min of aggitation at 4 deg celcius in a isotonic lysis buffer can is sufficient to disrupt the plasma membrane, any experience with this?
Then spin them for 5-10 mins at 4 deg celcius. The supernatant should then ideally consist of cytosolic proteins.
Then resuspending the pellet in a mitochondria lysis buffer and using a disruption technique that is rough enough to open both the inner and outer membrane. Here I'm properly back to freeze/thawing, waterbath sonification or a blunt-ended needle and a syringe (the latter option can be done without detergent!?).
A further centrifugation would leave me with a supernatant full of mitochondrial proteins, though the disruption method will properly affect the degree to which there is any of membrane bound proteins left.
Please share your experience with: plasmamembrane disprution (method and buffer) & mitochondrial lysisbuffer.
Sorry for the long winded post, but felt context to be important :-)
Thanks in advance for any advice, sincerely Martin
Martin LundMember Since 06 Sep 2012
Offline Last Active Oct 24 2012 02:10 AM
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My research interests
Mitochondrial Proteomics & diseases.