Did you think about differentiating them into macrophages then perform the assay?

The size of the pores in the membranes you used are too small. Use membranes with larger pores or look into the devices we designed recently for monocyte migration. Also keep in mind that monocytes do not start migrating right away, there is usually a time dealay that can be as long as a few hours.

Have you tried to optimize your assay with fresh monocytes? How about using 5-10% FBS as chemoattractant?

What you need to do is set up a Zigmond chamber (google it) and there are many out there. You would use the amount migration of monocytes....per thaw....you will need a microscope with video capablities.

Freshly thawed monocytes will not respond very well. It is better to use them freshly isolated, and then, they do respond better. You have to consider to give them time (a few hours) to recover after the freezing process. Check the viability of your cells as well.

Also, as other people noticed, maybe 3um is too small, although 8um will give you more background. The 3um works ok with, again, fresh monos: the numbers will be much lower, but will much less background, since you have them "actively" stretching to migrate through the pore (quite an effort for cells 10-12um diameter; that's why 8um pore is not really a good option either). Maybe , for your conditions, 5um should do better.

Use RPMI 0.5-1%BSA or some other alternative richer medium than HBSS (that's what I use, along with most of the people I know)

Also, take into account that if you let them rest for too long, they'll become adherent. You'll need to find and optimal resting time for your purposes. Perhaps using teflon wells, on which monos will not adhere, is a good alternative.

DO NOT try increasing the time, since your chemotactic gradient will be gone soon and you'll be seeing mostly chemokinesis.

THP-1 expressed high CCR2 receptor could response to MCP-1,which only a few of human primary blood monocyte may expressed CCR2

Good morning, I do similar work but with primary Drosophila haemocytes, particularly plasmatocytes the putative Drosophila homolog to mammalian macrophages. Although not the same, the optimisation techniques would be the same in principle. Primary cultures are a difficult one to optimise as you want to try to recreate the in vivo behaviour but in ex vivo conditions. Where I started with mine was simply the culture conditions, keeping it basic to just media and supplement. I found that primary cultures prefer higher concentrations of FCS/FBS, mine work best at 25% (v/v) FBS supplementation. Although it would seem the culture conditions appear well optimised already, you need to take into account the other factors within your media and the concentrations you have them at, as pH, etc. also play as factors in cell behaviour.

Now moving on to ECM coating, there are two types of ECM application - wet and dry, which by the sounds of it you are using a dry application. I have found that both work effectively for observing cell motile behaviour and migration, but there are subtle differences. Moreover, the way you treat the glass coverslip (or surface to which you apply cells to) also plays a huge factor in how the ECM spreads across that surface and thus how your cells spread across that ECM. For any wet application I would suggest autoclaved glass coverslips that have undergone a high molar HCL treatment and thorough washing, in advance of ECM application. Next, are you planning on testing other ECMs other than fibronectin? You might find a generic ECM like collagen IV, or a collagen mix ECM like gelatine might work. Conjoined to this you need to optimise the ECM concentration, which I think for most in vitro mammalian cell lines is anywhere between 1:10-1:40 d.f. from stock solution.

Once this is all optimised, you may or may not observe migration. Migration on an ex vivo ECM can only occur if - the cell is capable of auto-polarisation through integrin-ECM interaction alone, - the cells are in some form of active state by chemo signal/ electrical signal/ mechanical stress signal whether that be by your additions to culture condition, or to some innate biological reason from when you sourced the cells. You may find that even application of growth factor (like fMLP) chemo-signals, for chemotaxis, may not work as the cell is not in an active state. Plus you will need to be careful considering the well size and the number of cells you seed as you may get instant saturation of cells with chemo signal due to a small well surface area, but that is just an idea.

Anyways, as I mentioned before my work is in Drosophila primary haemocytes not mammalian so naturally the cells are different. But I hope the core optimisation steps may help you out. All the best, and sorry for the essay!

In short, this is my experience:

CELLS.- The best cells for setting up a chemotactic assay are polymorphonuclear cells (PMN) and peripheral blood monocytes or monocyte-derived macrophages (macrophages -human and murine ones- from other tissues such as lung or milk do not work well).

ASSAY.- The best assay (although modern devices work also perfectly well) is that performed with the so called Boyden chambers or the modified Boyden ones.

Take in account first if you wish measure the number of cells migrated or the leading front (distance travelled by the leading front of cells using the microscope micrometer). I recommend this technique to assess locomotion, because is more accurate and reproducible although is more tedious.

FILTERS.- Polyvinylpyrrolidone-free polycarbonate filters are a good choise for monocytes and macrophages. Using PMN and the leading front method you should use nitrocellulose filters. By the way, do not autoclave filters or PMN will migrate less.

PORE SIZE.- The pore size should be 3 um for PMN, 5 um for monocytes and 8 um for macrophages.

CHEMOTACTIC FACTORS.- Zymosan-activated serum or LPS-activated serum are the best ones. N-formyl-methionyl- leucyl-phenylalanine have been used with very good reproducible results. Immune complexes (albumin-antialbumin, for instance) is other option. Supernatants from Concanavalin A-activated lymphocytes is a good chemotactic factor as well as some purified lymphokines. However, nterferon-gamma is not.

In summary, chemotaxis is very useful technique but a boring one. So, patience and good hands.

If you want to see primary monocytes moving, here is a movie showing monocytes (red) and neutrophils (gray) chemotaxis towards LTB4. Monocytes take few hours before they start moving, and then move three times slower than neutrophils. It all depends on what you want to do know about monocyte chemotaxis from your assay...

http://www.pnas.org/content/suppl/2012/11/22/1210269109.DCSupplemental/sm02.mp4

We used the 2D chemotaxis µ-slide from Ibidi (Martinsried, Germany) in 3D mode for real-time microscope-based human monocyte chemotaxis assays. You can freshly isolate human peripheral mononuclear cells using centrifugation with Ficoll (1.077 g/ml), wash once or so (minimal handling), add fluorescent (Alexa Fluor 488-conjugated) anti-CD14 antibodies, wash, mix the cells with 2x collagen type I (e.g. use final collagen conc. of 1.6 mg/ml), and add the suspension to a last-minute prefilled chamber with 10-20 nM fMLP in one of the two 40 µl reservoirs. The cells migrate toward the chemoattractant source with a mean speed of about 4-5 µm/min for over 8 hours. The assay is quite tricky to establish, but monocytes tend to be quite robust and highly mobile. You can monitor the cells with brightfield illumination, or, better, use time-lapse fluorescent imaging to monitor the CD14+ cells (basically, the migrating cells in the brightfield channel will be the green fluorescent ones).

It would be best to use fresh blood from healthy donors - 10-15 ml is more than enough. You can freeze and thaw cells, but the quality will be questionable - done this before for Ca2+ imaging, but fresh cells were much better.

Dear Ulrike,

Please forgive me if there are repeats in my answer but there are so many responses.

As I am working on chemotaxis in the last 25 years I think there are 3 points which you should consider as very important in Boyden of transwell etc. assays:

1. size of the pores

2. time of incubation

3. optimal concentration range

From your description I could learn that the size of the pores (3 microM) is too small. Perhaps the assay works with this filter in THP-1 but not in these monocytes.

I could not find the concentration range of MCP-1. As far as I know the preferred conc. for monocytes in 10-12-10-9 M range.

Well, I have left the incubation time to the end of the list. I feel it is OK, however, If you have the 3 microM pore filters for assay, it is a choice also to oncrease the incubation time to 5-6 hours.

Best regards from,

Laszlo

Thank you all very much for your replies! Your comments really help a lot.

I started with the mentioned assay conditions according to a technical bulletin of BD: http://www.biocompare.com/Application-Notes/43112-Optimized-Chemotaxis-Conditions-For-Primary-Blood-Monocytes-Or-THP-1-Cells-Using-BD-Falcon-FluoroBlok-96-Multiwell-Insert-Plates/.

Next we are going to perform the test with freshly isolated monocytes, replace the buffer by RPMI and add fMLP as a chemoattractant. If that doesn’t work, we’ll try out plates with bigger pores.

Dear Peter,

It was interesting to see that you use the Ibidi u-slides. I had tired this out before during the optimisation of my cell culture set-up, but could never get the optics correct. I have now moved to a glass bottom dish set-up, provided by MatTek Corps, and resolution were vastly improved. But just out of interest, what microscope set-up are you using for the best optics, and how did you overcome the issue of media drying out?

Best,

Chris

1. Frozen cells are a bad idea. Systems as complex as actin-based movement are very badly affected by any alterations in underlying cell physiology, and monocytes are fragile. Use fresh cells if possible.

2. Transwell assays are rubbish. They are studded with artifacts, most of which can be corrected if you have a will to do so (google "checkerboard assays") but the worst problem is you can't see the cells so you can't judge their health. Ibidi slides or Zigmond chambers are way better. Dunn chambers are better still. Insall chambers are for some reason my favourite. You could also consider under-agar assays, which allow a very high quality of observations. Ibidi are the easiest to use. But ANY of these are miles better than a transwell.

Dear Chris,

We observe the cells in the observation area (2000 x 1000 x 70 µm channel connecting the 2 reservoirs) of an Ibidi 2D chemotaxis slide using phase-contrast (and fluorescence) microscopy via a 10x/0.3 objective. The slide is pre-filled with medium. You add 15 µl chemoattractant-containing medium (plus color dye, as visual indicator) to one of the reservoirs, then, straightaway, you add the monocyte/collagen suspension (10 µl) to the connecting channel. All 4 plugs are inserted, so the system is water-tight (unable to dry). This is the approach for 3D chemotaxis assays using the "2D" chemotaxis slide (we prefer this to the Ibidi 3D chemotaxis slide).

When I've used primary human monocytes that were frozen and thawed, even though the viability looked good (Annexin V and Trypan blue staining were negative) I could never get the cells to chemotax anymore. If possible, freshly isolated cells is best. I use a microchemotaxis assay from NeuroProbe for primary human monocytes and it works beautifully. The system consists of an upper and lower chamber separated by a polycarbonate membrane with 5 um pores. I haven't coated the membrane with anything, the cells stick to it perfectly fine. Here's the link to the chemotaxis chamber I use. Good luck!

http://www.neuroprobe.com/products/ap48.html

I am using the Dunn Chamber assay on U937 cell line activated with a phorbol ester and stimulated with EGF. And I could nor notice any movement of the cells when running the time lapse movie for 5 hours. It demands a lot of patience. And I couldn't find any good protocol for this assay.

Hi everyone, here’s our update: Now that we are using freshly isolated monocytes, RPMI and inserts with 8 µm pores (fluoroblok inserts with 5 µm are not available) the chemotaxis assay works quite well. Thank’s again for your input! Ulrike

I'm working with BMDM, and they seem to resist freezing/thawing, but I am not sure if their ability to chemotaxis will be unchanged. Thanks to all these valuable suggestions, I'll try some under agarose assays and possibly Ibidi slides in the coming weeks instead of transwell assays which don't seem to be as good to evaluate chemotaxis.

Dear Ulrike,

I see that you have tons of answers - let me contribute also with some questions or suggestions:

1. Why do you use 3um pore size. We used THP-1 also but 5um pore worked much better - the cells still had to migrate actively.

2. The incubation time is nice - 3hrs -, however, sometimes it requires more. As I could learn from 15 years chemotaxis research first of all a time course experiment is required when you start an experiment.

3. What was the concentration of MCP-1? Reguraly it works well in 10-8 - 10-6 M conc.

As you are working with thawed cells - have you tested the viability of cells?

If you need any more practical hints please contact me on [email protected].

Regards from,

Laszlo

I am afraid that any functional assay on previously frozen monocytes will not work, since a lot of chemokine and activation receptors are lost. If you have fresh blood you should not have a problem doing migration assays.