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For Best Results with Confocal Microscopy
Which microscope should I use?In short, Theresa, the Facility Manager, will help you determine this after discussing the details of your experiments. Each system has different strengths. The nature of the information you need from the images, the type of manipulations you need to do (in the case of live cells), and the fluorophores you choose will determine the microscope you'll need. The deconvolution system (Inovision) is able to collect the three most common fluorescence emission wavelengths (red, green, and blue) with a minimum of complexity. It also has the most sensitive camera, which is important when collecting images for deconvolution. If you do not need 3D imaging, but just want a nice single image, the Inovision system is the fastest and simplest solution. With the optional digital deconvolution, images from the Inovision system can also achieve 3D resolution and greater clarity. A thin sample with a punctate or linear distribution of fluorescence seems to show the greatest improvement. The Inovision system is an inverted microscope: the objective is below the sample. This makes it good for live-cell studies with a specialized culture dish, but chamber slides cannot be used without removing the chambers. Chambered coverslips can be used. Standard microscope slides are appropriate for all of our microscopes. The confocal microscopes (the LSM 410 and 510) are more versatile in 3D imaging capabilities, but they are complicated to use and their detectors are somewhat less sensitive. If you have a thick sample with high signal but also high background, the confocal microscopes will probably give you the best images.
The two main differences between the LSM 410 and 510 are in the microscope configurations and the available excitation wavelengths. The LSM 410 is an inverted microscope with the live-cell capabilities listed above for the Inovision system; the LSM 510 is an upright microscope outfitted for electrophysiology. Because triple labeling is increasingly common, both confocal systems can resolve three fluorescent labels: green (as in GFP or FITC) and red (as in rhodamine, Cy3, or DsRed), plus a third label. The LSM 410 uses a higher wavelength to visualize a far-red label such as Cy5 or TOTO-3. The LSM 510 (in 2-photon mode) can excite shorter-wavelength dyes such as DAPI or coumarin, and it also possesses the greatest flexibility in changing filter sets for multi-labeled samples. In fact, the LSM 510 can handle up to six labels, if the correct filters are available.
What kinds of labeling can I look at?Most three-dimensional optical microscopy techniques use fluorescence imaging. The Facility's microscope systems are therefore optimized for looking at fluorescence.We can also collect DIC (differential interference contrast, also called Nomarski) and black-and-white phase-contrast images. However, these images are not confocal.1 Fortunately, fluorescence imaging is versatile and powerful, especially when combined with 3-D resolution. Background is low, spatial resolution is high, the method is largely non-destructive, and samples can be co-labeled with two or three different colors. The best-known methods of fluorescent labeling for microscopy are probably immunofluorescence (using fluorescent antibodies to bind directly or indirectly to a molecule of interest in a fixed sample) and GFP fusion (DNA encoding green fluorescent protein is inserted in the coding sequence of a gene of interest, producing a fusion protein visible in live or fixed samples). Other types of fluorescent labeling include the following:
Fluorescence can show much more than just the location of a molecule or structure of interest; here are some creative variations.
Notes
How should I prepare the sample?Because of the immense diversity of samples that can be looked at with fluorescence microscopy, no one can deduce the ideal conditions for your particular sample. The best guide is the experience of other researchers who have prepared samples similar to your own. Following are some general points that I hope will be helpful in optimizing your protocol.FixationSome types of imaging do not require fixation. If you need to preserve the sample in its current state, or if you need to do antibody staining, generally you must fix and permeabilize it. In brief, fixation either aggregates or cross-links molecules, especially proteins, to keep dead cells and tissues from falling apart. Permeabilization opens holes in cell membranes, allowing antibodies and other large molecules to get in (and any unfixed molecules to escape).
Techniques vary widely with the nature of the sample and the type of labeling desired. Two general references I have found quite useful are Polak and Van Noorden (1997) and Lane and Harlow (1999).
SectioningA major advantage of confocal microscopy is the lack of need for thin tissue sections. In my experience, a 10-µm or 20-µm section under the confocal microscope gives images as good as, or better than, a 2-µm section viewed by conventional fluorescence microscopy. It is easier on the tissue as well as on the researcher, and gives better anatomical context.With thicker sections, up to several hundred microns, confocal imaging is still successful, but a few caveats arise. First, antibodies tend to diffuse into the center of the tissue very slowly or not at all. Second, spherical aberration2 reduces signal from deep regions of the sample. Water-immersion objectives can alleviate the problem but cannot eliminate it. Deeper regions also suffer signal loss from light scattering, a function of the amount of tissue that must be traversed by both the excitation and the emission light beams before the signal emerges. Despite the abovementioned limitations, we have successfully imaged embryonic rat kidney, brain slices, large cellular aggregates formed in culture, whole-mount rodent gut, flattened retina, and other thick samples on both the LSM 410 confocal and the LSM 510 multiphoton systems. Larger tissue samples allow observation of macroscopic structures such as embryonic organs , ganglia, and neural networks. Some model systems (e.g. brain slices) do not function well in very thin slices. For these reasons, the physiological advantages of thick samples often outweigh the technical difficulties. Fortunately, those difficulties are constantly being alleviated by better labels, better objective lenses, and better imaging technology. Multiphoton microscopy (as with our LSM 510 system) has proved very useful in overcoming the last problem, scattering. It excels at imaging of intrinsic labels such as GFP fusions, or of injected and easily absorbed labels such as neural tracers and indicator dyes.
MountingMounting Medium and Antifade ReagentsIf a sample is allowed to dry after staining, bright artifacts will result from precipitation of fluorescent antibody. Mounting medium used between slide and coverslip prevents drying of the sample and can greatly influence the quality of imaging. A glycerol-PBS mixture is suitable for most fixed immunofluorescence and GFP-labeled samples, using from 50-90% glycerol, 1x final PBS*, and pH 8-9. *Recipe for 10X PBS: For 1 L, dissolve in 800 mL distilled H2O: 80g NaCl, 2 g KCl, 14.4g Na2HPO4, 2.4g KH2PO4. Adjust pH to 7.2. Bring volume to 1 L. Autoclave. Source: Lane and Harlow (1999). An antifading additive is highly recommended. Time and the act of observation both accelerate the loss of fluorescence from any sample. This inevitable process can be slowed by adding various reagents, chiefly antioxidants and radical scavengers, to the mounting medium. Commercial products are available, such as (in no particular order):
Slides and Coverslips
For high-magnifcation work, a glass coverslip is essential. The oil-immersion objectives (40x, 63x oil, 100x) have short working distances owing to their extremely high numerical apertures (a necessity for the highest resolution). Therefore, you will be able to focus a maximum of ~50 microns into the sample (beyond the coverslip) with the 100x objective, and 75-100 microns with the 40x. A small excess of mounting medium can make the difference between visibility and invisibility of an object on the slide. Thus, it is preferable to grow cells on the coverslip rather than on the slide. If you use the chamber slides with removable walls, make sure you remove the walls completely so that the coverslip can sit abolutely flat. If possible, use a glass slide or rigid plastic. Bending of plastic slides under oil-immersion objectives impairs focus stability. However, if the plastic slide is of standard length, a glass slide can be placed under it to prevent bending. Coverslip Placement
Fading is minimized if the samples are kept in the darkand at low temperature: 4°C if the medium is mainly aqueous; -20°C if enough glycerol is present to prevent freezing. Other things to remember when mounting samples:
After the coverslip is mounted and sealed, household window cleaner (Windex, Glass Plus, Sparkle, etc.) is recommended for removing oil, buffer salts, and other material (scientifically referred to as 'schmutz') from slides and coverslips. The gentlest method is to squirt the cleaner on a cotton swab (Q-tip) and wipe the side of the swab, with very little pressure, across the center of the surface. Excessive pressure or large amounts of cleaner will dissolve the nail polish and damage the sample. Use a fresh swab to dry the coverslip, and repeat as necessary. Do not use window cleaner on microscope optics!
Recommended references
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