I have been trying to image live endothelial cells (contact mode) with our atomic force microscope (AFM) for the past few weeks. I can image fixed cells perfectly fine, but living cells are much less stiff and therefore harder to image. I am currently employing two cell culture surfaces - BD petri dishes with gelatin coating and poly-lysine coated cover slips. The cover slips yield better images because of (I'm assuming) increased stiffness which increases cell stiffness. I use CO2-independent media and keep the cells at 37 degrees during this process. I allow them to adhere for 48 hours or more, as longer adhesion times have been demonstrated to increase cell stiffness (for ECs). Finally, I use Olympus/Asylum TR400PB probes (long lever, less stiff), and have an Asylum MFP-3D-BIO AFM with the BioHeater stage for maintaining cells at 37 degrees.
I am curious what protocols, techniques, culture surfaces, surface treatments, tip selection, buffer selection, etc. other researchers are employing to successfully image ANY living cell type with an AFM in contact mode. Please be as specific as possible, because everything seems to influence image quality on an AFM. Also, if it will influence your answer, I plan to do simultaneous fluorescence microscopy of these cells with the Zeiss Axio Observer A1 upon which the AFM is mounted.
please check this book http://www.springerprotocols.com/BookToc/doi/10.1385/1592596479
Hi,
In the following papers we have been using Asylum MFP-3D-BIO AFM to image living endothelial cells, similar circumstances as you discribe:
http://www.ncbi.nlm.nih.gov/pubmed/17115151
http://www.ncbi.nlm.nih.gov/pubmed/17921584
http://www.ncbi.nlm.nih.gov/pubmed/22038122
The main difference might be the cantilevers: we were using Bio-lever, BL-RC150 VB-C1, Olympus. The TR400PB is rather for AC mode in liquid, or contact in air. The Biolever is the only one wich is working for us in contact in liquid (spring constant 0.03N/m).
One other idea could be the silanization of the cantilevers. Protocol briefly:
"The silanized tips are strongly hydrophobic, hindering the tip–cell interaction. The procedure consists of vapor deposition of a mixture of 10 ml bicyclohexane, 1–4 drops of octaldecyltrichlorosilane and 4–8 drops of carbon tetrachloride. The tips were incubated overnight in a closed chamber together with the above mixture."
Best wishes,
Zoltán
I don't know why you have to imaging under contact mode, but I would say tapping mode is much better for imaging live cells. Be careful when you use very soft cantilever, cause they have quite low resonant frequency, so you may need choose second harmony to drive your tip to oscillate. Yeah, Veeco DNP-10, tip C or D is working well.
I imaging my cells ( osteoblasts, trabecular meshwork cells) in PBS, which provide physiologically environment, since keeping live cells staying on the AFM for maximum 3h anyway.
Hi!
Have a look at this webniar: http://www.asylumresearch.com/Webinars/index.shtm
It will be available as a record soon. If you have any urgent questions, I would recommend to contact the speaker Irene Revenko via eMail ([email protected])
Hi,
i also did a lot of cell measurements with an AFM from Agilent (Model 5500). I always used silicone nitride AFM tips with a low spring constant (approx. 0.06 N/m). You have to check your set-point before you start with the image. So you have to adjust the set-point at the lowest possible value in order to apply a force as low as possible on the cell surface. The imaging conditions i used for my experiments were: Normal buffer solution (PBS), room temperature, living cells grew on "ibidi-dishes" or on fibronectin coated PDMS membranes. After two days of incubation living cells (osteoblasts, lung epithelial cells) became adherent and could be used for AFM imaging. I did not image for a longer period than 2 hours, then cells often loosed their contact to the substrate.
Good luck!
I agree with Baiyun Liu, tapping mode in PBS with DNP probes should work well as I used that method to image live bacterial cells on poly-lysine coated cover slips successfully. Your tip get contaminated in contact mode. I also suggest to trap your cells in polycarbonate membrane with the pore size similar to your cell size instead of gelatine coating method.
Hi Navid, We are able to image live cells, human, microbes, fungi, in contact mode. We use cantilevers with very low spring constants (0.02 - 0.05 nN/nm), but it is not trivial. We do, however, get better images of live cells in contact mode than we do in either non-contact or intermittent contact mode. It certainly helps to use some sort of physical or chemicals immobilization method. Best of luck! Tanya
Try a softer cantilever such as MSCT (0.01 N/m) or OTR4 (0.02 N/m). Note, OTR4, not OTR400.
After gaining much more experience on AFM, the answer is quite simple - softer probe (Bruker MLCT or similar), slower scan, smaller area. Also, if the cells are too active, try imaging them in PBS instead of their own media - that tends to slow them down. Thank you all for your help. Please see my new AFM question if you can help with imaging of Carbon Nanotubes. Enjoy the attached image of a living endothelial cell monolayer.
Hello Navid,
I am going to image fixed neuroblastoma cells with AFM (Multi mode AFM, possibly with contact or tapping modes) in a few days. As you are expert in this field, do you know any working protocol for cell fixation and sample preparation for AFM?
(1) wash the cells on glass slide or plastic dish with HBSS buffer (with Ca & Mg);
(2) fix the cells in 4% paraformaldehyde for 15min - one day; Or, fix the cells in 0.5-2% glutaraldehyde for 1-60min;
(3) wash the cells;
(4) treat the cells with 50mM ethanolamine for 10min to inactivate the free aldehyde;
(5) wash the cells.
Step (1) is optional - you can also fix cells in medium. Step (4) and (5) are also optional. For topography imaging, inactivation of free aldehyde is not necessary. But for force measurements and recognition imaging, it is essential. For fluorescence imaging, it can reduce autofluorescence.
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