Live cell imaging is often a race against the clock in which stress must be minimized to maintain cell viability. In microscopy, stress is caused largely by light exposure— -- ultraviolet light damages DNA, infrared light causes localized heating and fluorescence excitation results in phototoxicity to tissues and cells. There is generally a tradeoff between capturing a fabulous image and collecting enough quantitative data with a high enough signal-to-noise (S/N) ratio.
“You need light to get the image [data], but this damages the sample … so think in terms of what you can give up,” says Paul Herzmark, imaging specialist, Department of Molecular and Cell Biology at the University of California, Berkeley. “Do you really need all those z-sections, multiple colors and so many time points? By eliminating one element, you gain light to use elsewhere.”
In parallel with deciding exactly what imaging is required, culture medium, temperature control and pH are important factors. Any adjustment can alter your cellular processes of interest. Herzmark and DeLaine Larsen, imaging specialist, Nikon Imaging Center (NIC) at the University of California, San Francisco, suggest the following four tips for finessing image-acquisition conditions with live cells.
Optimize the sample environment
Mammalian cells in bicarbonate-based media require 5% CO2 to maintain physiological pH. Without a CO2 supply, cells are adversely affected within five minutes. To minimize pH change, you can supplement media with 25 mM HEPES buffer when possible. (Cells can proliferate without CO2 if the medium is buffered with 25 mM HEPES at pH 7.4, but this environment is viable for no more than 10 hours and is highly dependent upon your cell line and cell concentration.)
“Work out your growth conditions beforehand,” says Larsen. “You need to know what keeps your cells happy before you sit down to image. This is especially important the longer you image, when it’s more problematic to keep cells healthy.”
Most of the live cell imaging at the NIC is performed on the spinning-disk confocal and epifluorescence microscopes and runs no longer than three days. Both microscopes have a CO2-controlled stage-top incubation chamber for working with mammalian cells. (Mammalian cells typically require a stable 37°C environment.) However, you can also construct a do-it-yourself incubation chamber using packing material and an egg-incubator heater, see Construction of a Heated Incubation Chamber around a Microscope Stage for Time-Lapse Imaging from Cold Spring Harbor Protocols [1]. To further control temperature, it’s important to use an objective heater when using an immersion lens. A temperature sink is created between the immersion objective and sample. Additional objectives should be removed from the turret in such cases, to avoid heat transfer. Temperature differences in the room also can cause problems with sample drift and focus, as heat causes parts of the microscope to gradually expand and contract. Cover vents in the room if air blows on or close to a microscope.
Test for autofluorescence
Grappling with autofluorescence cuts into precious imaging time and can lead to bleaching or phototoxicity. Riboflavin, tryptophan and phenol red are well-known culprits. To improve S/N, nonspecific background fluorescence can be reduced by using phenol-red-free medium, for instance.
“Unless the cells have very specific growth requirements, try switching to a different growth medium that has less autofluorescence at the wavelengths you need,” says Herzmark. “Otherwise, experiment with different filters. If the autofluorescence is slightly different in color, you can switch the fluorescence filters in some cases.”
Herzmark specializes in two-photon microscopy, which is best suited for thick samples. Regardless of your fluorescence imaging technique, he recommends testing for autofluorescence beforehand. If you’re using GFP, for example, set up the experiment under the scope with the cells but without the GFP. Illuminate with the GFP filter set, look for any visible fluorescence and adjust accordingly.
Monitor cell health regularly
Check the health of your cells throughout imaging, especially in longer experiments or when using UV-light sources. Some obvious signs of cell stress include: bulging (‘blebbing’) of plasma membranes, large vacuoles, enlarged and isolated mitochondria and clustering of fluorescent proteins. For more subtle changes, cell health can be monitored using live-cell stains. It’s advisable to use the smallest amount of fluorescent compound for labeling—often 10 to 100 times more dilute in concentration than recommended—to reduce cell toxicity caused by dyes. Adding 5% BSA in the labeling medium can help minimize nonspecific binding, as well.
Improve light throughput efficiency
Finally, fine-tune light throughput to collect as much fluorescent light as possible so incident light may be decreased, thereby reducing phototoxic effects. For multicolor experiments, use filters and mirrors designed for single dyes and collect images sequentially to optimize light collection for each fluorophore. Collect images in all detection channels using control samples that contain single dyes. Imaging these samples in the same manner as the multicolored samples lets you correct for cross talk and bleed-through beforehand.
For the sake of cell preservation, live cell images typically have low S/N and never use the full dynamic range of the camera (usually ≤20%). To generate higher S/N images, reduce the intensity of the light and increase camera exposure time. However, all this depends on your specific cell type and experiment—keep practicing, and good luck!
Reference
[1] “Construction of a Heated Incubation Chamber around a Microscope Stage for Time-Lapse Imaging,” Cold Spring Harb Protoc, 2007, doi:10.1101/pdb.prot4792 (http://cshprotocols.cshlp.org/content/2007/7/pdb.prot4792.full)
The image at the top of the page is from the Nikon Imaging Center at UCSF.