I am using fluorescence microscopy to detect apoptosis and necrosis. I use cancer cells and treat it with anticancer compounds. I am using Annexin V (apoptosis) and Propidium iodide (1 mg/ml ) for necrosis. My protocol is as follow:
1. Seed cells on cover slip (16 hours)
2. Treat cells (24 hours)
3. Wash cells twice with assay buffer (provided with Annexin V kit)
4. Add Annexin V and PI solution and keep in dark for 15 min
5. Wash cells twice with assay buffer
6. Fix cells with 4% para-formaldehyde solution
7. Wash cells twice with PBS
8. Transfer cover-slips to microscope slides and using DAPI
9. Observe under fluorescence microscope.
However, I am running into difficulties.
1. Although, I can see blue stained nuclei. I am seeing green colored cytoplasm and dark red nuclei as well as cytoplasm. I expect my compound o cause apoptosis as well as necrosis. But,even in the control samples, I see the same.
I am not sure what wrong.
I would be grateful if you can suggest me how to improve on this.
Hi Pratik,
I don't exactly work with cells now, but based on what I read I can suggest you few things.
1. May be the two channels that you are using are overlapping. Try using Annexin V and PI separately and see how they look, since they channels wont overlap.
Also, you can try increasing the dose of your compound.
Hope it helps,
Good luck.
Wrong choice of experiment. 1. Washings before fixation remove most of apoptotic and necrotic cells. 2. cell membranes are compromised so Annexin stains intracellular annexins. 3. Hard to have accurate quantification due to loss of cells and small area (cover slip).
I would suggest PI-FACS which is quantitative and you can detect apoptosis (complete shifting of peaks to left) and necrosis (a minor shift of G1 peak to left). If you gate these two populations you can quantify both. Note: 24 hr time point will also have apoptotic but secondary necrotic cells (which is different from necrosis). Good luck.
@ Goodwin: Thanks!
1. I am actually going to do flow cytomtery for accurate quantification. However, I decided on using fluorescence microscopy as my second step. My first step was to use trypan blue to study the percentages of necrotic cells.
2. I did modify my protocol. After removing treatment media, I simply added the Annexin V and PI dyes. I then kept in dark for 15 mins. I then washed twice and fixed cells with 4%PFA. I then washed twice and mounted coverslips on microscope slides. However, even this time all dyes were inside cells (even in control sample).
It would be better to go for the Flow
cytometric analysis rather than microscopic analysis.
Hi Pratik,
If it is not a problem of your microscope then definitely you are loosing the membrane integrity as your control cells are also PI positive. How does your cells look before staining? Double check that you are using the proper dilutions of the washing/staining buffers and stains? Try the protocol with some other cells (just untretaed). May be your present cells are sensitive to the reagents. Also washing with PBS after annexin V staining is not a good idea as it will remove Calcium ions and then the annexin V will fall off and might give nonspecific signal. Use the washing buffer provided with the kit.
To troubleshoot if it's caused by staining or by cell culture, you can stain with PI in the cell culture media and check the cell by live cell imaging. You'll see if control cells taking up the PI dye in live cell imaging. If control cells don't take up PI in nuclei, then wash the cells and then probably you can stain AnnexinV quickly. If the cells don't stain PI but mostly stains AnnexinV, then the annexinV staining is causing rapid cell permeability. If both PI and annexinV staining are good in this sequential staining, then you can further optimize this cell staining procedure.
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