Organotypic Culture - Science topic

Hello Ilia V. Potapenko,

PTFE inserts are used in organotypic cultures because of its biocompatibility and non-adhesive nature. Being a chemically inert material, it does not react with cells or tissues, ensuring a biocompatible environment. Its non-stick nature prevents the explant from adhering to the insert, allowing for easy manipulation and preventing damage to the tissue. It allows for the maintenance of the 3D structure of the tissue, which is crucial for studying organotypic cultures. Also, the porous nature of PTFE membrane allows for the efficient diffusion of nutrients, oxygen, and waste products across the membrane, supporting the viability of the cultured tissue. 

Detailed information on the properties of PTFE may be obtained from the link below.

Best,

Hi,

I think that the data sheet for that solution says that is not suitable to 3D cultures as it has been sterilised using UV light, that eventually can interfere with the cross linking that is necessary for 3D cultures

You are most welcome Kristýna Valášková

Precisionary makes vibrating microtomes: precisionary.com

Ted Pella used to make a model, but I'm not sure if they still do.

Some other options can be found through electron microscopy sciences. https://www.emsdiasum.com/microscopy/products/equipment/vibrating_microtome.aspx

I know this is a few years late, but I happen to be looking for one myself now and a google search brought up your question, so perhaps this information will help others.

i use a 1ml sterile tip with a tip modified (just cut with scissors :D) s o you can just suck the slice out from the petri dish and release it on the membrane.

Thanks Matteo Donegà I would be very glad to hear about any update you have.

You embed the tissue block in low-melting agarose on ice. Cell viability would be sufficient for most of the experiments.

I agree with the comments regarding the filter change, electrode position, lower submerged setting. However, there is still one basic condition: careful preparation of acute slices. It looks like they're not alive.

The color of a collagen solution will sometimes revert over time back to acidic if the buffering capacity is low. I use 10x EMEM for the mix. HBSS itself in the incubator becomes acidic. Additionally, the color should be more red when you add the cells; if the final concentration of the collagen is around 1.5 mg/ml or less, the cells will sink to the bottom before it polymerizes if it is too acidic. If you can get the pH to 7.5 but less than 8, you can quickly mix and plate without such problems. But I agree change from HBSS to a better bufferend formula like MEM or EMEM or DMEM, whatever is available at a 10x concentration. Hope this helps.

If you want to stain live cells, formalin fixation is not possible. After 2 days in formalin everything will be dead.

I could imagine that it depends on the cellular position of the antigen you want to stain: Staining cell surface proteins will be much more likely than proteins sitting in the mitochondrial membrane for example as you need to get access to the epitope.

Good luck, it sounds like an interesting but difficult idea!

But I want to use AAV...

hi Debora,

Most of commercially avalable AAV-based vectores give 80-90% knockdown. I am sure, that in vitro everything will be fine)). The question is, would you move forward to the mice models or not? If so, take into account, that AAV are eliminated from cells within 2 weeks. If your futere plans are about drug to neurons, then non-integrative system is not a good idea. If your aim is just to look for the best way of knockdown - no difference in virus based shRNA - delivery systems, just the choice of personal preference)))

Hi Enrico,

I'm also interested in the suitability of the Alamar Blue assay for organ culture and came across your request. Have you tried it in the mean time?

Cheers, Cornelia

Many thanks to you Prof. Raghunath and Prof. Leavesley. Your suggestions are very helpful. 

O.K,. have managed to adjust (find) the media which supports the optimal growth of each cell line individually?

Me! I used it to assess proliferation at 1d, 3d 7d of fibroblasts in fibrin/HA gels, ill look for the protocol and send ot back to you 

Hi Apoorva,

Generating skin organotypic cultures is tricky and I am afraid the major issue with your system is that the HaCaT cell line is not ideal for generating these skin equivalents since the cells already have a partially to fully differentiated phenotype under normal culture conditions due to the high calcium content of both standard media and fetal bovine serum. However, I attached a protocol for you from the guys who actually derived the HaCaT cell line (Stark 2004). They also use DMEM/FBS for cell culture and then, after seeding the cells onto the collagen gel, used TGF-b to induce terminal differentiation. If your HaCaTs appear already fully differentiated during normal culture, you might want to try thawing a new vial or even consider purchasing new cells from ATCC.

In general, I would recommend using primary human keratinocytes (also availbale from ATCC) in combination with serum and calcium-free growth medium (e.g. EpiLife https://www.thermofisher.com/order/catalog/product/MEPICF500). Cell differentiation at the air-liquid interface is then induced by adding calcium to the medium. I attached a detailed Nature Protocol paper for you. Good luck!

Tara,

the very basic editing of any stack image can be done by selecting, on ImageJ:

Image-Adjust-Color Balance

Image-Color-Channel Tool

Always have both open to play around with your images. Channels tool is crucial, under "Color" you can modify each channel (including the color), under "Composite" you can look at co-localization. Be aware that double or triple channel images can be exported (using Image-Type-RGB Color) only in Composite mode.

If you have stack images, you can either show single planes or Maximum intensity projections (Image-Stacks-Z Project).

Also, the Despekle and the Smooth features can help you cleaning up your images.

Thanks for the answer. i will try your suggestion.

Hi Veronica,

I tried few times in vitro  mouse skeletal muscle culture, overall i test Soleo and EDL muscles in order to guarantee edium diffusion in whole tissue avoiding necrosis deeplu in the organotypic culture. I used in parallel standard plastic dishes or teflon grid to leave the muscle in supsesion not attached to the plastic in DMEM 10%FBS. I did not notice much differences, the muscles were alive, but the intercellular space was noteworthy increased do to medium soaking.

I hope this will help you,

  good luck

Cesare

Dear Luz

It is difficult to answer your question without more details of how you set up the experiment, but if the section is sufficiently thick (I assume it must be 300 micron or thicker) then perhaps you can inject a tiny volume (say 1 or 2 microliters) directly into the slice. If you take a small needle (25 gauge) and shove it onto a 10 microliter pipette (fits well on eppendorf pipettes) then you can pipette as low as 1 microliter of cell suspension (very high density) accurately. If it works you should get localized pocket of cells where you inject. In that case they would interact with and perhaps invade the surrounding tissue more rapidly.

If you stick with the gel matrix delivery, find a way to position it on top of or adjacent to the slice using some kind of improvised well/spacer as described here: http://www.jove.com/video/50881/coculture-system-with-an-organotypic-brain-slice-3d-spheroid

Good luck!

Best Regards,

Mark

Que:Also, is it necessary to adjust the pH of ACSF before or after bubbling with carbogen? I would assume beforehand?

I don't have an indication of specific culture media, but perhaps to apply different size of slices could be helpfull. 

Thank you!

you can use glycerol . the culture grow for day and centrifuge after Platte get add glycerol sterilized and keep in -20 c

Thank you all for your answers.

- Daniel, we don't use organotypic cultures for electrophysiology only, we also use them to evaluate cell viability after Oxygen and Glucose Deprivation and we also perform immunofluorescence, protein extraction and RNA extraction on them. We do not destroy the insert, so we thought that we might be able to reuse it. We were given a protocol, but in our experience, recycling the inserts is not convenient because liquid gets through the pores and floods the top of the insert.

- Ulrich, there is another method to fabricate your own inserts in Koyama R et al., 2007, J Pharmacol Sci.

For anyone interested, the protocol for recycling the membranes is the following:

1. Remove the cultures from the insert gently with a spatula or brush

2. Rinse inserts in tap water

3. Place inserts in Trypsin-EDTA (0.05% trypsin, 0.02% EDTA in 0.9% NaCl or HBSS) (we use leftovers from cell passages)

4. Incubate 1h 37º in incubator.

5. Rinse 3 times in demineralized water

6. Place in 70% alcohol for at least 24h

7. Dry alcohol under the hood and store under sterile conditions