I am trying to set up the in vitro assay to check glucose uptake in mouse adipocytes. I use collagenase digestion of mouse fat pad tissue, centrifugation and a few washing procedure then. At the end I get oily cell suspension and try to culture it in usual 24-well plates. I have a few questions:
- how to count mature adipocytes after isolation to prepare cell suspension with needed concentration? I tried to use trypan blue and hemocytometer but this way doesn't work cause cells are very big, always floating and can be disrupted easily.
- how to make cell culture free from oil released from disrupted adipocytes? I tried a few washings but oil is still in wells.. is this oil toxic for culture?
- how to maintain this culture - all cells are floating and no attached cells in the wells.. I tried to stain them by Oil Red O, it is rather difficult to do because only a small amount of cells can be fixed on the well because of their floating..
- is this floating culture normal to check parameters like glucose uptake or cytokine production? I mean these parameters could be affected by absence of cell attachment or presence of oil in culture?
- did anyone try to use adipose tissue explants instead adipocyte cell culture? how to culculate exact percentage of cells in each well in this case? Should I use just weight of tissue fragments per 1 ml?
thank you in advance for any help!