Dear all,
I have been trying to optimize a transwell migration assay. However I have some difficulties regarding the counting part. I am doing crstal violet staining and cleaning the upper part of the membrane with a kimtech paper. (Because the surface of 96 well transwell membrane is too small for regular cotton swabs) Then I cut the transwell membrane, mount them and later I clean the upper part again and take some pictures. But I don't know which cells I should count.
Here I attached a picture that I took in 63X and these are my questions:
-Number 1, 2 and 3 are the pores?
-Number 4 is a cell which is about to migrate through the pore?
-Number 5 is the right cell to count for the assay?
I would be really glad if you can give me some feedbacks.
Your criteria for designating a cell as 'migrated' may need to be more stringent, but as long as you count all conditions with the same threshhold you should see a phenotype.
1. Many people only count cells that spread out after crawling through the membrane. The cells in the image appear to be stuck in the pores. Adjusting time of assay (at least 6 hours) or serum/chemoattractant in the bottom chamber can help solve that problem. The speed of migration will be cell type/context dependent.
2. At 63x , the residual cells on the top (unmigrated), are usually out of focus and will not be included in the count.
3. For washing the cells from the top, use a PBS-soaked cotton swab and a twisting motion to remove nonmigrated cells.
4. Alternatively, you can trypsinize the membranes and the migrated cells will detach and fall/adhere to the bottom plate.
Good luck!
Dear James Preston,
Thank you so much for your feedback!! Actually I am using 10%FBS as chemoattractant and stop the experiment after 24 hours of seeding time. I am wondering whether I should use a different attractant or a different time point. Another question is how you perform the trypsinization. You just add trypsin to the upper side of the transwell, trypsin penetrates through the pores and only the cells at the bottom fall to the receiver plate, am I correct? Could you please give more information about that too?
A usual positive control is soluble CXCl12.
Another option is to collagen coat the inserts. THe collagen must be dilute, or the assay becomes an invasion assay rather than a migration assay.
The trypsin should be applied to the bottom chamber.
If using a single membrane (e.g. Neuroprobe) then disassemble the boyden apparatus and add a 96 well plate with trypsin/PBS, as if you were going to plate another migration assay. If using single inserts then simply transfer them to a new well containing trypsin.
Thank you so much again for your answers. Unfortunately I am using a whole 96 well inserts. I guess I can just transfer them to a new receiver plate with trypsin inside.
From what I understand, the experiment must be in duplicates - one for trypsinization and one for staining.
You can extract the crystal violet using SDS.
1. After imaging, simply cut your membrane and place in a 24-well plate.
2. Add 150µl of 1-5% SDS to extract crystal violet (~5 min on shaker)
3. Plate 50µl (Triplicates) into a 96-well plate and read the O.D at 570nm.
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