Hi Naveen - your main problem is that you are getting an enormous amount of bleedthrough from your green channel into your red channel. You can see this in the areas of bright green, the signals overlap pretty much completely. Look in the area that is dark for Calcien AM - you'll see nuclear staining with PI. This is what dead cells look like. If you solve your channel bleedthrough you'll have fixed a bulk of your problem. I also agree with Goodwin that EtHD1 is much better than PI - PI at high doses or in long treatments tends to also stain cytoplasmic RNA. You can definitely get your conditions right so that you can use it but generally speaking EtHD is better. I've published a protocol on live/dead staining in cells and a protocol on confocal microscopy of live tissue slices (in which I show analysis of live vs dead cells by depth using z-series) that may be helpful: "Ex vivo imaging of excised tissue using vital dyes and confocal microscopy" - PMID: 22752953 and "Assessment of Cell Viability" - PMID: 23546778. Both list good spectral replacements for the dyes you're using, and a couple viability dye sets are demonstrated in the live-tissue protocol. Feel free to email me questions if you have any.

Dear colleague,

As you said Z-Stacks of slides is an easy thing while 3D is a tough one. As I understood you are using tissue culture plastic ware instead of glass slides. As there are sometimes a little bit thicker it you should check for the LWD of you objective if you are using inverted imaging. Regarding you problem, I am hope a microscopy expert is able to help you.

As an alternative I wanted to suggest that you could also try to make a single cell suspension and run the cells by flow cytometry if only numbers matter and not spatial distribution of dead cells within your tissue. As cells will be sticky I would suggest usage of fixable viability (ebioscience) or zombie aqua dyes to discriminate for dead cells and fix the cells afterwards.

Dear Sander Bekeschus,

Thanks for your reply!

Actually, my microtissue is like a small ball (1mm thickness). So I just place them on a glass slide.

The problem with a single cell suspension is that I would have to use trypsin and collagenase which might kill some of the cells.

Naveen

Use Ethidium homodimer1 instead of PI. Ethidium homodimer cannot enter the cells unless the cell membranes are compromised. If you trypsinize cells avoid trypsin step.

Good luck.

Dear Goodwin Jignesh,

Does not Ethidium homodimer 1 behave in the same way as PI?

Naveen

Hi Naveen - your main problem is that you are getting an enormous amount of bleedthrough from your green channel into your red channel. You can see this in the areas of bright green, the signals overlap pretty much completely. Look in the area that is dark for Calcien AM - you'll see nuclear staining with PI. This is what dead cells look like. If you solve your channel bleedthrough you'll have fixed a bulk of your problem. I also agree with Goodwin that EtHD1 is much better than PI - PI at high doses or in long treatments tends to also stain cytoplasmic RNA. You can definitely get your conditions right so that you can use it but generally speaking EtHD is better. I've published a protocol on live/dead staining in cells and a protocol on confocal microscopy of live tissue slices (in which I show analysis of live vs dead cells by depth using z-series) that may be helpful: "Ex vivo imaging of excised tissue using vital dyes and confocal microscopy" - PMID: 22752953 and "Assessment of Cell Viability" - PMID: 23546778. Both list good spectral replacements for the dyes you're using, and a couple viability dye sets are demonstrated in the live-tissue protocol. Feel free to email me questions if you have any.

Dear Naveen,

Yes. EtHD1 gives red fluorescence. Like Simon said, bleed through is a problem in your image. Since you're using confocal microscope, its easy to set excitation and emission spectra you need. You had mentioned only 488 which is good for Calcein-AM. You need to set one another around (excitation/emission maxima ~528/617) for PI or EtHD1. Have control cells stained with single stains such as Calcein-AM alone and EtHD1 alone or PI alone and make sure you set the wavelengths right, so that it cannot bleed through other channel (if you check Calcein-AM make sure it is not bleeding to 546/red). This can also be fine tuned by reducing the exposure time, by increasing the averaging values and by adjusting the pinhole settings. Good luck..!!

Hi,

I've manged to image my microtissue again, this time with a fluorescence microscopy instead of a confocal and from the image the live cells can be differentiated from the dead cells quite easily.

But since red light penetrates tissue more easily, I can see the dead cells in the center more easily than the live cells. Is there any other way I can overcome this? Like may be using a different dye? (Digesting the cells is not something I am looking for).

Actually, my final step is to check if there are any dead cells on the surface of the microtissue (In terms of percentage). And its posing to be quite a problem!!!

I think it may be best to use a confocal microscope rather than a brightfield, if you have the option. With a confocal you can collect a z-stack and get an idea of cell viability in 3D.

If switching dyes might help you have a number of options, here are just a few suggestions: one easy one would be to use a blue nuclear dye (Hoechst 33342) with PI or EtHD1 and just count viable as blue without red, inviable as blue + red. This may end up being less sensitive but works. You could also try Sytox Green for dead cells (it is FAR superior to PI in my experience - very bright and very nuclear specific) and a blue or red cell tracker dye for live cells. I think Sytox Green, Cell Tracker Red, and Hoechst 33342 would be a very good dye set for this - you'd get all nuclei blue, dead cells would have green nuclei, and live cells would have red cytoplasm.

Hi Naveen, I am experiencing pretty much the same problem with you. Did you manage to overcome yours? 

Hi Gozde, 

Yes, it work quite well for me. Instead of using the confocal microscope I used the carl zeiss ApoTome.2. with optical sectioning and filters to image my micro-tissues and they looked perfect! Just like what I wanted. 

I would say that the best way to image your micro-tissues would be to first have a live control and dead control. And then to calibrate using the microscope so that your dead micro-tissues look red and live ones look green.

Hi Naveen, thanks. I  do not have any other option apart from the confocal so I have to give it a try with the confocal. Hope I will also find a way like you did :)

Instead of using two different fluorescent tags, use only PI for counting dead cells (Quantify) and for representative purpose use DAPI+PI.