Centrifugation can't be guaranteed to remove cells from the medium, so if it is important to do that, you can pass the medium through a wide 0.45 um filter (available from tissue culture suppliers).

If the medium loses a lot of its CO2 during this and/or during collection from the cells and storage, the high pH can be damaging to cells. So it is best to re-gas it, ideally by placing in a too-large bottle, adding excess gas from a cylinder of 5%/10% CO2 (whatever you use), and swirling- not so much that the serum (if any) froths. Or otherwise by placing in the incubator in a container with a loose top, say overnight. But some activity (cytokines etc) might be lost in the second method.

Collecting conditioned medium and demonstrating its effect on another cell population is very difficult to reproduce. If you do suspect chemokines and cytokine involvement, its best to prove that with SELDI or another proteomics method. Then you can add this chemokine/ cytokine in serum free media to your target cells and recheck the proteome. Hope this helps!

I use fibroblast conditioned media to examine the paracrine signaling on myocytes. In order to condition the media, I grow the fibroblasts to confluence, wash them well with serum-free media and incubate them with serum-free for 48 hours. I collect the media and use it at various dilutions. Analysis of chemokines/cytokines is rather difficult if you condition using serum-containing media. There is not really any reason you need to stimulate your cells unless you have a specific question, i.e. does hypoxia result in differential activation of EPCs.

Dear Hou,

You can culture the EPCs until confluent.At confluency change into media with atleast 1% serum.EPCs are sensitive to serum starvation and a large number of cells may detach and float.

Another alternative is to change into complete media (EBM+EGM) and collect the media after 24 hours/48 hours.In this case you have to use complete media as your control in estimation of the chemokines or while performing functional assays.

Hope this helps!

If you need to have serum in your medium, which becomes your culture medium - then your control conditions are essential! What you need to do is to have wells with no cells, which are treated in exactly the same way as the cells used to condition the medium e.g. 1 dish with cells, one dish without and that is the only difference. You then need to assess the impact of both conditioned media on the target cells of interest. Not an ideal situation (better to not have serum present), but at least this provides you with a decent comparator.

Centrifugation can't be guaranteed to remove cells from the medium, so if it is important to do that, you can pass the medium through a wide 0.45 um filter (available from tissue culture suppliers).

If the medium loses a lot of its CO2 during this and/or during collection from the cells and storage, the high pH can be damaging to cells. So it is best to re-gas it, ideally by placing in a too-large bottle, adding excess gas from a cylinder of 5%/10% CO2 (whatever you use), and swirling- not so much that the serum (if any) froths. Or otherwise by placing in the incubator in a container with a loose top, say overnight. But some activity (cytokines etc) might be lost in the second method.

Hi Yuning

we have grown EPC in full EGM 2MV + 5% FBS, then swithced in EBM2 %1% FBS (same medium as before without Bullet kit cytokines) and incubated the cells either in normoxia or hypoxia (1.5% O2, 5% CO2) for 3 days. The supernatant was centrifuged and filtered through a 0.45 um filter.

You might check the following paper for more details:

Di Santo S, Yang Z, Wyler von Ballmoos, Völzmann J, Diehm N, Baumgartner I, Kalka C. Novel cell-free strategy for therapeutic angiogenesis: In vitro generated conditioned medium can replace progenitor cell transplantation.

PLoS ONE, 2009;4(5):e5643.

You may consider first using an insert membrane system (semipermeable) in which the cells are placed in 2 chambers that allow the media but not the cells to diffuse. I would perform this PRIOR to using conditioned media. We have successfully used this in several studies. I think they are sold as 'transwell' membrane systems.

Dear Hou,

As most of them suggested it is better to avoid serum when collecting the conditioned medium. And EPCs in general secrete lot of angiogenic cytokines, stimulating them in the hypoxic conditions may further enhance their angiogenic secretion profile.

Based on the molecular weight of the cytokines of your interest, you can separate and concentrate particular fraction of the conditioned medium using centricon concentraters.

Lahoucine izem

I worked with conditioned media from different cell type for a long time. My experience is that you should not have serum or other variables in your collection media, they complicate the analysis of you data, If you need to stimulate your cell with a compound do that first then wash it off extensively before adding collection media. In my hand this was the best way. Good luck

In collecting CM from human fibroblasts, I find substitution of serum with 0.1% BSA useful in collecting CM. Confluent fibroblasts are quite happy in medium with 0.1% BSA from several days (routinely use 48 hrs to collect CM, but cells can go even longer) although of course growth rate slows, but they are confluent after all. BSA protein seems to provide some beneficial factors (maybe free fatty acids as an energy source or other bound mediators). Addition of some protein source to CM also reduces loss of any secreted proteins that might bind to plastic surfaces or other mechanisms. BSA does not seem to interfere with any bioassays or ELISA assays that I employ. Just make sure you use cell culture grade and that it is relatively endotoxin-free or low endotoxin. I do not use FFA-free for this purpose.

We grow stromal and glial cells in coculture system in 2% FBS in order to get conditioned media. Then we draw off 2/3rd of the media and replace it by fresh media. Stimulation by a single factor may bias the culture system and very difficult to get rid off.

As James Fabisiak has suggested, it makes a lot of sense to avoid for conditioned media any serum and add some BSA instead. In the blood albumin acts as a carrier for many things like lipids or bioactive peptides presumably including also transport of noxious metabolites. In addition, in vitro it is beneficial to prevent peptides from sticking to culture vessels. We have used successfully BSA as supplement in defined media, though a very good quality should be chosen to avoid unspecific effects.

My conditioned medium protocols for various cell types is as follows. Note: because there are many variables, it's important to keep as many constant as possible. Keep the serum batch constant, and make sure it supports both conditioning and target cell types.

First: how low a serum concenntration can your conditioning cells cope with?

Passage cells in a range of serum concentrations to find the minimum they will grow in.

Conditioning:

Either

method 1 uses non-growing cells: Grow cells as normal and when they are confluent, instead of passaging them, change to low serum medium. Leave that medium on for a set (arbitrary or experimentally determined) period of time. Usually 3 or 4 days, or the amount of time between passsages, or a week. Collect the medium.

Method 2 uses actively growing cells: Passage cells at usual density into the low serum medium. At the density you would usually passage them, collect the medium.

Spin down hard and long. Filter sterilize. Can usually store in frozen aliquots, but test.

Dear Lahoucine Izem and Lori Walker ,

In this case that you produce your conditioned media in serum-free media, what is your control? Is your control a serum-free media cultured during the same 24h of condition media? Then in this case, do you treat your target cells direct with 100% conditioned media or do you dilute it in full media? I am asking it because I would like to know how exclude the effects of serum starvation from the effects of C26-cells secreted factors?