I currently using FACS (Aurora Cytek) to quantify necrotic Gr1+ cell death from the spleens of immune competent mice. I target Gr1+ cells in the spleens of tumor-bearing mice using a photodynamic agent which has been shown to be very effective in vitro.
After delivering in vivo cytotoxic phototherapy to the spleens of mice, I wait for 4 hours to dissociate the spleens. Four hours after targeted phototherapy, the spleens are passed through a 70 um filter with DMEM media, then red blood cell lysis carried is out for 10 minutes. After counting cells, I am able to determine a reduction in viable splenocytes through cell viability assays such as CCK8 (a 40% viable cell reduction in the phototherapy treated mice vs control).
I must demonstrate the selective targeting of Gr1+ splenocytes using a more sensitive technique such as flow cytometric analysis. Currently, I am using anti-Gr1 conjugated-PE, anti-CD11b-FITC and fixable live/dead stain to detect changes in non-viable Gr1+ cells. Of the Gr1+ cells, only about 7-14% (n = 3) of them are non-viable.
Is it possible that most of the dead Gr1+ cells are washed away during the multiple washing/centrifugation steps, resulting in a lower non-viable Gr1+ population? Should I use propidium iodide instead?
My staining protocol is conducted as follows:
1) Spleen dissociation with DMEM
2) Centrifugation (1,600 rpm for 7 min)
3) Aspirate, then resuspend in 1X RBC lysis buffer (10 min), then neutralize
4) Centrifuge (7 min), aspirate
5) Resuspend in 10 mL DMEM
6) Count viable cells in Hanks and trypan blue, then aliquot 1-1.5 million viable cells (dead cells are, of course, present) into FACS buffer
7) Incubate in anti-CD11b-FITC and anti-Gr1-PE (FACS)
8) Centrifugation
9) Wash with FACS buffer
10) Resuspend in 1X PBS, then incubate in far red live dead stain for 30 min
11) Centrifugation
12) Wash with 1X PBS
13) Fix cells in formalin
14) Store samples in FACS buffer for flow cytometric analysis.
Any suggestions are welcome! Thank you.
If I were you, I would probably add simple trypan blue after each step. It would be good to know the absoslute number of your cells (which is most likely decreasing) and the percentage of viable cells at each step of the preparation. Your idea with adding PI is also good.
Best method to quantify the Necrotic cell population has to be Annexin-V FITC/PI dual staining by flow cytometry. This is a very well established protocol and exact. You can refer to some of my papers for reference.
Awesome, Dr. Satyam Kumar Agrawal . I thought so as well. I happened to use the you the LIVE/DEAD™ Fixable Far Red Dead Cell Stain we generally use in my lab, but I've come to realize that certain staining protocols may not be optimal for quantifying AND retaining dead cells. Thanks for the advice.
Remember that during the preparation of cells for FACScan many are getting lost and many will be gated out. The good old trypan blue shows you ALL cells. See Zhivotosky, B., & Orrenius, S. (2001). Assessment of apoptosis and necrosis by DNA fragmentation and morphological criteria. Current protocols in cell biology, 12(1), 18-3.
Dear James,
Try double plot assay Annexin V and PI. It will hrlp you to distinguish the necrotic, live and apoptotic cells as well.
I have some questions for you:
1. Which type of formalin you are using?
2. Have you checked your centrifuge speeds for this testing? A centrifuge speed validation could help.
3. Have you checked pH of washing buffer?
4. There are many other dyes for measuring viability with flow cytometry such as 7-AAD (we use this dye for our CD34 counts), PHK-26 and such (Molecular Probes, now Invitrogen has many different ones), if you think that everything else is fine after your check try one of these other dyes. Hope this information is beneficial for you.
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