Hi Melissa Corcini Berndt . What days of culture were these images taken on?. The last two images seems to be having proper cell density than the others. Try splitting in 1:3 ratio. Add cold dissociation reagent to the wells and leave it for 5 minutes or scrape and collect the matrigel dome completely and keep the tube on ice for some time before centrifuging.

As for contamination:

Has your matrigel dome inside the media changed color like straw yellow or looks turbid?

Do you observe any movement in your wells when undisturbed under microscope?

Please share if you have a picture taken in higher magnification.

Hello Janani Shanmugam ! Thank you for your answer.

The pictures were taken three days after splitting. Prior to that, the organoids had been in culture for ten days after thawing from cryovials.

The Matrigel looked relatively clear. It always picks up a bit of the medium’s color, but that’s all. For the very full wells, I admit it’s harder to judge, perhaps they were slightly more turbid.

I’ll try your tips for splitting. I hadn’t kept everything on ice during the process, so far. Do you think I should limit the up‑and‑down pipetting to 10–15 strokes to reduce shear stress on the organoids?

There was no movement whatsoever, which makes me wonder if the culture is contaminated. In standard plate cultures without Matrigel, you can clearly see organoid movement. Is movement also visible when they’re embedded in Matrigel?

Unfortunately, I don’t have higher‑magnification images.

To clarify, the movement I mentioned is about the particles other than the organoids to rule out contamination not the organoid itself.

Yes during the passaging reagents should be cold. So try collecting the organoids using cold dissociation reagent. Pipetting should be complete and swift. After collecting the organoids, you can keep the suspension in ice for 5 to 10 minutes, so that the matrigel dissolves completely. If you find a white jelly like layer after centrifuging it is probably matrigel. You can centrifuge again with cold 0.1% BSA in PBS which will give you a refined pellet.

But the media you add should be at room temperature.

1. Try to increase the split ratio to 1:3

2. Use cold reagents and even cold pipette tips if you can during passaging.

3. Minimize the pipetting strokes.

4. Tip: If there are tiny blebs like bubbles around your organoid, they are stressed.

5. Since you have mentioned, there is no movement or turbidity of media or matrigel, contamination might not be an issue as per my knowledge.

Hope this will be helpful. Let us know if it helps.

Best of luck Melissa Corcini Berndt

I apprechiate your tips Janani Shanmugam! I will try that and get back to you to report if I achieve more consitent results. Thank you!

Janani Shanmugam

Hello! I just wanted to share some feedback.

I tried a new isolation and followed most of your suggestions and it really helped!

The changes that made the biggest difference were:

• Keep everything cold/on ice during splitting and harvesting. When I process a full plate, I keep the finished tubes on ice in the meantime. This improved viability and, more importantly, gave me much more defined pellets, every step was easier to handle. The same was true for RNA harvests; I’m getting very good, consistent results now.
• Split/harvest before they overgrow. Organoids seem much more prone to contamination once they get too big or dark, with cells starting to die. StemCell recommends 7–10 days in culture for colon organoids, but in my experience 9-10 days is almost always too long.
• Keep the split ratio around 1:2. I avoid 1:3, as growth was poorer when fewer cells were in the gel.
• Pipetting 45 µL domes instead of 50 µL (24-well plate), I’ve had better results with that.

Thank you for the input. I hope this helps someone else who’s struggling, too!

Melissa Corcini Berndt

Happy to help and glad that you are getting good results.

Thanks for getting back. Best of luck.