Getting the organoids out of the hydrogel

[Ellen Nuttall Musson]

Hi TooT!

Thanks for creating this great resource.

We are trying this for the first time in the lab and had some difficulty in separating the organoids from the hydrogel completely, despite doing 3 extra spins at increasing speeds.

Now that they are in the plate, you can kind of see that they still have some hydrogel associated with them.

Is this a problem that you have come accross before? Do you have any additional advice to overcome it? Do you anticipate that it would affect e.g. cell engraftment?

Best wishes

Ellen

[Abdullah Khan]

Hi Ellen, 

You're very welcome! Thanks for being involved. 

This is always a bit of a pain and unfortunately it varies depending on the lot of Matrigel/collagen etc. We are working on a modified protocol that we think will fix this, but it's not fully validated yet. As soon as it is we will share! 

For now what we say is if there is a little bit of hydrogel in the wells around the organoid that is not a problem - usually we spin the organoids in the 96-well plate at a low speed (like 50-100G for a couple of minutes) so there isn't lots of free floating material. Usually the cells grow into the excess ECM and it isn't a huge issue after about a week or so you get a nice round structure. This is what I would suggest for now, and is what we do routinely and it doesn't seem to impact patient engraftment (we get good engraftment and results afterwards). 

Does this make sense? You will find it is very ECM batch dependent. Sometimes you get beautiful separation... but sometimes... not so much. We should have an improvement ready in a few months but for now a lot of the 'OG' experiments we do are as above and we (touch wood) have no issues with end point. 

Hope that helps, 

Abs

[CAMILLA TOMBARI]

Hi TooT!

Thank you for creating this group and for sharing your experience.

We are starting to work with organoids and we are experiencing the same problems as other people in extracting individual organoids from hydrogel, even if we have followed the suggestions you shared in this chat. Do you have any new updates?  Have you ever tried at this stage to work in cold conditions to improve hydrogel dissolution? Do you think organoids may suffer cold centrifugation and washing?

Thank you!

Camilla

[Abdullah Khan]

Hi Camilla, 

We have some updates we are putting together, probably be a few weeks though if you don't mind hanging on it's a busy couple of months but I'm working on some formal protocols. 

In the interim, in theory yes though we haven't tried cold conditions to improve hydrogel dissociation. It depends on how cold and how long for, personally I'd be a bit hesitant to cold shock cells. The thing you can try if you're really struggling is doing a brief collangease wash post extraction to remove the excess (short being something like 5 mins at 37 deg). If too long you run the risk of damaging structure. 

If you post some photos of how much ECM you have left floating around I can better judge how big the issue is. A lot of the time excess ECM gets absorbed into the structure as it grows/cells proliferate

BW + thanks for TooTing 

Abs

[Ka Ka Yu]

Hi Toot! 

I am just starting to make these organoids and ran into a similar problem. Would love to get your advice on this and wondering if there are any updates on optimization protocols? 

The hydrogel seems to form a clump with the other organoids. I have tried frequent centrifugation steps, 400g 4mins, 500g 5mins, 700g 5mins, 800g 2mins, the hydrogel seems to still be attached to the organoids and I cannot see a distinct interface between hydrogel and organoids, so I couldn't manage to remove any of the hydrogel. Shown in photo attached. 

I have tried to treat half of the organoids with collagenase type II for 5mins 37C in waterbath. I will see how this affects the organoid structure in validation experiments. 

Additional note, I saw that VitroGel® Organoid Recovery Solution does help to remove hydrogel. Has anyone tried this? 

Thanks all! 

Ka Ka 

[Abdullah Khan]

Hi Ka Ka, 

The best method we've found to be honest is to switch to the approach described here: https://www.biorxiv.org/content/10.1101/2025.02.16.638505v1

When this happens this is often because of a batch (usually matrigel) which is protein rich so it makes it harder to separate them. 

My advice is, if possible, try out the granular method in the manuscript. It is actually much more straightforward than the bulk extraction! If you are just getting started, it might be a good time for the switch. Like I said to Salim, I am a bit snowed under for a few weeks but will try and get a formal protocol up here in the next couple of months. We put a lot of detail in the methods section though hoping it would help people reproduce. 

BW
Abs