Hi everybody,

Thank you very much for all the opinions, advice and support. With some of the suggestions on here, I have managed to overcome the problems with CaCO-2 adhesion in my 96-well assay sufficiently.

This included:

Washing cells with serum free media

Growing the cells in 10% FBS media

Using manual aspiration methods rather than the pump.

Just for anybody elses information:

The CaCO-2's were p.40 in my experiments

The 96 well plates were Nunclon T.C treated

I had to use 96 well formats, because that was the point of developing my assay for high throughput analysis.

The media was ATCC formulation (MEM-Earle's Salts supplemented with 2mM L-Glutamine, 1mM Na Pyruvate and I reduced the serum to 10% FBS).

From my observations I think 'doming' may have been an issue (as suggested by Norbert). As CaCO-2 cells like to group together in tight clusters when grown in monolayer’s, I suspect the reason I was only having problems with 96-well growth surfaces was due to the small surface area, so a high percentage of the wells' cell population was not anchored to the growth surface instead being more tightly connected with each other. It might still be the case that they are not 'happy' being washed in PBS (apologies but I do not have time to fully investigate this, it worked with Serum free media washes so that is good enough for my purpose), but the cell population could also have been disrupted through sheer forces of pipetting liquid into the wells at a velocity high enough to cause some 'floating' of cells away from the growth surface, and with pump aspiration, the suction seemed to rip high numbers of cells away and out of the assay. By being gentler in these steps, I have been able to overcome the problem and produce data in the assay repeatedly.

If anybody else experiences any problems like this and has the opportunity to investigate adhesion problems in greater detail than I currently can, it would be interesting to find out what exactly is going on with CaCO-2's in 96 wells.

But for now, thank you very much to all for your help. This was my first time using researchgate (or any forum to be honest), and I can really see the benefits of being able to easily discuss practical problems and bounce ideas within a research community. I will definitely be using the webstie and participating in discussions frequently into the future.

Many thanks,

Mark

I haven't worked with Caco-2 in a while, but I remember similar problems. You might try a few different plate brands, as they can vary a lot with regard to cell adhesion. More to the point, I have overcome these problems for assays with numerous cell lines just by using a flooding/flicking wash. Cheap, fast and effective, you just use a wash bottle (cut off the tapered tip so you have a large opening in the dispensing tube) and gently flood the plate with buffer, then flick it by hand into a disposal container. Repeat as required. Obviously not easy to make this a sterile method, but for assays this is usually not a consideration. You also have to spend a little time wiping the plate afterwards so that optical surfaces are clean for the plate reader. There are expensive programmable plate washers that allow adjusting flow rates, so you could reduce the dispensing rate of buffer if that feature is available, but the flooding/flicking method is a tried and true method that has been used by many people for decades.

I haven't worked adheion assay on 96 well plates but quite often I use either the 24 well standard cell culture plate or the 8 chamber slide cell culture plates and both of them worked fine. The trick is start with small amount of cells like 10,000 cells/well (500 ml media/well) i.e in a 24 well cell culture plate so that they will have enough space to grow and they will not overlap one over the other. In my case I grow them for 21 days, changing the media everyother day. During washing be gentle, always add the PBS to the side and aspirate it gently and manually.

Another point: ATCC recommends 20% FBS for Caco-2 growth. This difference could potentially be a factor affecting cell adhesion. See http://www.atcc.org/ATCCAdvancedCatalogSearch/ProductDetails/tabid/452/Default.aspx?ATCCNum=htb-37&Template=cellBiology

Yes ATCC recommends 20% FBS for Caco-2 growth but if I am not mistaken it is only if use thier media. We use 10 % FBS either with MEM or DMEM and worked fine.

You could also try washes with sterile PBS + 0.1mM CaCl2 + 1mM MgCl2 to prevent junctional damages.

Caco2 adhere really well on most plastics but they can sometimes start behaving strangely at late passages. Are yours relatively young? What can happen with old caco2 cells is that they may detach as a sheet as they get very confluent when you wash them with PBS.

You could try a different brand of plate just in case. However, sometimes if the cells are too confluent they may start detaching as they start relying on eachother for adhesion. Gentle PBS wash with sterile PBS with the addition of 0.1mM CaCl2, as Sebastian above suggested, works well. However, it depends on the subsequent assay, so check before you add it. I tend to use large pipettes (10-25ml)with wide opening and using the "slow" setting on the pipetteman so adding the PBS dropwise and then gently aspirate. High serum concentration may actually interfere with adhesion so I would stick to 10% or bellow as a general rule.

Thank you very much everyone, some interesting considerations for me to try out! I've tried different plastics before, but have to keep with a 96 well format for the assay. I'm gentle with my sterile PBS washes, but think it is the aspiration step causing issues. My cells do grow to a high level of confluencey (they're p.37 by the way), and I did suspect they may use eachother for anchorage due to the 'bunching' nature of their cell-cell interaction. Therefore I will have another go at reducing the serum content in the growth media, and adding CaCl2 to the PBS. I'll let you know how it work, but thank you for the sussgestions so far :)

Hello again, high passage number can be an issue depending on the cell line.

It is generally recommended that you don't use cells that have been in culture for extended periods of time as their characteristics may change. So it maybe good to try a new vial of cells if available for comparison.

Good luck

One of the guys in our lab has exactly the same problem with CaCO-2 cells grown on Mattek dishes. When he has multiple manipulation steps for IF preparation they detach in sheets. This might well be down to conditions, but it's also a cell line specific phenomenon. For example NIH3T3 behave the same and detach in sheets while HeLa do not. Have you tried these manipulation steps at a lower confluency? It may stop a detaching cell taking its neighbours with it.

I don't use PBS for washing cells any more, I use the media with out any additives, for your example them MEM without serum, L-glu and Pyruvate. I have done test on several confluent cells in 24 well cell culture dish, If I leave a PBS on them for 10 to 15 min they start to detach from the wells. Using MEM only, for washing, might help.

All the best

HH

Firstly I strong agree with Christophe, in addition, the younger they are the more fragile they become when cultured. Do you grow them for long (e.g longer than 5 days without changing media or sub-culturing)? if so consider a high passage like >20, at that passage number they are very matured and will attach without a problem, in fact the older they become the more they become resistant to detachment. That is why they are considered a "gold standard" model for permeation/absorption assays in which you grow them in a membrane for about 21 days without splitting them. Another important thing to consider is your cell density for a specific assay. Area of 96 well plate is very small, its highly possible that they over multiply within a short period (their doubling time is less than 24 hours).

Good luck!

Salts could Help during filtration but PBS may not form strong ionic pair to pass together. Can PBS form a stable ionic salt?

Confluent Caco-2 cells growing on hard plastic or glass surfaces differentiate to form nearly impermeable polarized monolayers with tight junctions between cells. Because of selective ion flux they transport fluid into the basolateral space and consequently form domes, which are in fact detached from the plastic (search "Caco-2 domes" and you will find references and images). How much dome formation you have depends mainly on how long the cells have been confluent in culture. It may just be a matter of timing for your experiments... do you even need confluent cells? (which, because they differentiate and are polarized, are quite different biochemically from sub-confluent cells). Also, it's difficult to compare your experience with that of other labs using Caco-2 because there is so much heterogeneity from lab to lab, especially among labs that have used the cells for many years. This problem is discussed in PMID 19131524. You may be able to solve your problem by growing cells on plates that have permeable membrane surfaces. They adhere better and don't form domes, but this is an expensive alternative. The gentle washing method that I pointed out above does work, but it is not a high-throughput method, if that is a consideration.

I fully agree with Norbert Kartner. Growing Caco-2 cells on plastic is all right and so is using them in assays for sub-confluent undifferentiated cells. However, as the cells start the differentiation process leading to formation of cells in monolayer linked by tight junctions, the plastic substrate becomes non physiological as the cells transport fluid in the basolateral space and form domes that facilitate cell detachment upon washing. Differentiation on permeable substrates should be the rule for epithelial cells such as the Caco-2 cell line, although it is an expensive option and high throughput assays are more complicate to develop. But it is wrong to assume that the permeability of the substrate does not influence these cells biology and differentiation.

Hi

I would recommend to check the pH of PBS,. Also, you can dilute the stock of PBS and try. Good luck

Caco-2 cells could be easily detached from plastic support with PBS since they need, like all epithelial cells, Ca++ and Mg++ ions to adhere (Trypsin solution contains EDTA in order to chelate divalent ions and facilitate cell dissociation). Washes must be performed by using complete medium and cells could be used under confluence to avoid cell monolayer disruption.

I have the same problem for several years but in breast cell lines. I have used MCF7, MDA-MB-231 and MCF10A (non tumoral) with the same results; in 96 weel-plates washes steps I lost cells. I tried manually with a micropipette, with an automatic 8 channel pipette, with a plate washer at the lower velocity of draw in and rinse and overturning the plate, but more or less the results are the same. Also I tried with PBS, as well as cold PBS or complete MEM, but always with the same results. I changing plates brand too, without results.

Finally I modify wash steps on my protocol to reduced them at maximum, because I give up with this misterious (this is science!)

you can also try to use specific pre-coating to promote a better cell adhesion (Poly-Lysine for example).

I've tried with Nunclonc surface, negative and positive charge surfaces and now I order a new phisical treatment to adhere cells. But with poly-lysine I've not tried.

Thank for the suggestion.

Coating is a good idea, but before switching to poly-lysine, or any other surface, I would make sure to verify that the new substrate is not changing the cell behavior -- many adherent cell types have distinct responses to substrate features (including -- surface texture, chemistry, charge state, coating with purified extracellular matrix molecules, biomaterials, biomimetics, etc). In our experiments we found that attachment of bone marrow mesenchymal stem cell monolayers can be undermined by proteases (i.e., MMP-2, a gelatinase that cleaves non-fibrillar collagen type I) released into the conditioned media (Guzman-Morales et al 2009, Bone). In an osteogenesis assay, our thick monolayers start to detach sporadically, around 7 to 10 days into the assay. Detachment is attenuated -- and osteogenic differentiation promoted -- by adding dexamethasone to the culture media which suppresses MMP-2 release. Stromal cell attachment that involves integrins and collagen type I is probably different from epithelial cell attachment.

Very interesting your answer. I am going to read more about it on epithelial cells.

Make sure that your PBS contains Ca and Mg. These divalent cations are important to cell attachment. The original Dulbecco's PBS contained these, but there has been a tendency over time to treat PBS as divalent free. Mainly that is because the divalents can be nuisance when dissolving the salts. The original recipes made a 10x as divalent free and a 10x divalent solution and then one mixed 1 part 10x divalent free with 8 part water and then 1 part 10x divalents. Divalents with phosphate are not very soluble.

Dear Mark, One of my students also experienced this problem. She worked really carefully and even used manual pipetting instead of the pump. She also pipetted the PBS onto the sides of the well and not directly onto the cells.

I see that you have received quite a lot of advice but according to a number of papers the method that you used should work. My student used 96 well costar flat bottom plates before. Somebody suggested using nunc plates instead, so she is going to try this. Also, she is going to dilute the PBS(which is a 1x dilution) a little before using it to wash the cells. In the paper of Wang, et al., 2008. (Title: CdSe Nanoparticles in Caco-2 Cell Cultures) they gave the following protocol:

Cell culture plates were filled with Dulbecco's phosphate buffered saline (D-PBS) and then inverted for 10 minutes, enabling unattached cells to be removed by precipitation. Cell layers were then washed gently with D-PBS five times at room temperature." We will try this as well. Please let me know about your successes in this regard.

Regards

Anne Grobler

Im going to try this as well.

.

Kind regards

Lizette

Cells are easily washed way from 96-well plates. I have overcome this problem by using a bigger well size whenever my assay allows (i.e. the reagents are not critically expensive).

Hi Mark,

We had a similar issue with a different cell line and were able to dramatically improve adhesion by coating our plates with 0.25% (w/v) gelatin (bovine skin in MQ H2O; autoclave to dissolve). Gelatin, though, is just heat denatured collagen so Caroline's comment above is of particular concern. Plating the cells on such an ECM may engage integrins etc and can dramatically affect whatever biology you're looking at, hence such methods should be approached as an experiment unto themselves.

Good luck!

Hi everybody,

Thank you very much for all the opinions, advice and support. With some of the suggestions on here, I have managed to overcome the problems with CaCO-2 adhesion in my 96-well assay sufficiently.

This included:

Washing cells with serum free media

Growing the cells in 10% FBS media

Using manual aspiration methods rather than the pump.

Just for anybody elses information:

The CaCO-2's were p.40 in my experiments

The 96 well plates were Nunclon T.C treated

I had to use 96 well formats, because that was the point of developing my assay for high throughput analysis.

The media was ATCC formulation (MEM-Earle's Salts supplemented with 2mM L-Glutamine, 1mM Na Pyruvate and I reduced the serum to 10% FBS).

From my observations I think 'doming' may have been an issue (as suggested by Norbert). As CaCO-2 cells like to group together in tight clusters when grown in monolayer’s, I suspect the reason I was only having problems with 96-well growth surfaces was due to the small surface area, so a high percentage of the wells' cell population was not anchored to the growth surface instead being more tightly connected with each other. It might still be the case that they are not 'happy' being washed in PBS (apologies but I do not have time to fully investigate this, it worked with Serum free media washes so that is good enough for my purpose), but the cell population could also have been disrupted through sheer forces of pipetting liquid into the wells at a velocity high enough to cause some 'floating' of cells away from the growth surface, and with pump aspiration, the suction seemed to rip high numbers of cells away and out of the assay. By being gentler in these steps, I have been able to overcome the problem and produce data in the assay repeatedly.

If anybody else experiences any problems like this and has the opportunity to investigate adhesion problems in greater detail than I currently can, it would be interesting to find out what exactly is going on with CaCO-2's in 96 wells.

But for now, thank you very much to all for your help. This was my first time using researchgate (or any forum to be honest), and I can really see the benefits of being able to easily discuss practical problems and bounce ideas within a research community. I will definitely be using the webstie and participating in discussions frequently into the future.

Many thanks,

Mark