I'm trying to establish a baseline of how fed and starved oenocytes look like in L3 larvae on my wild type strains. But I'm having a hard time finding them.
This is how I'm doing the experiment: I synchronize the larvae by collecting eggs for two hours at 25C, followed by 3 day incubation in a humidified chamber. I then dissect the larvae in PBS on ice (so that they don't move as much). I've tried various dissecting techniques - an abdominal longitudinal cut that exposed the internal organs but keeps everything in it's place; cut out the head and tail to let the solutions go in with minimum disturbance of internal structures; an abdominal longitudinal cut followed by removal of all internal organs and flattening of the skin. I then fix and stain with Oil Red O as previously described.
My starved larvae have much less lipid droplets than the fed ones but I cannot see oenocytes. Is there a dissection trick I'm missing? Or maybe a staining trick?
From the published images the clusters seem to be visible at low magnification, is this true, or are these cells only visible at very high magnification?
I have tried your last dissection methodology: i.e. Flaying the larvae by pinning the carcass on a silicone plate, removing all organs but the muscle/epidermis and fixing it on the plate. The staining of the cells with oil red was indeed a little week but I could see the cells at 20X using a Phase contrast microscope (same on used for in situ samples). The cells are pretty big and can be seen easily although they are buried behind the muscle layer. I would recommend staining some starved larvae as positive controls as oil red staining is much stronger in such samples. You may also try using the OK72-Gal4 to mark the cells with GFP to standardize the protocol to your convenience.
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