We receive frequent requests for staining protocols for fixed tissue. While we cannot provide specific protocols that cover the enormous variety of samples that users bring to our laboratory, we do have several general recommendations for sample preparation for widefield and confocal imaging. If you want to prepare samples for light-sheet imaging, please email Pablo to discuss your proejct before you begin preparing samples. Please note that MSL does not officially endorse any of the products mentioned below.


We are often asked: “is this staining real”? This question is unanswerable without proper controls imaged with exactly the same settings as the experimental sample. Here are some of the most common controls we recommend:

Autofluorescence control

Prepare a slide where you don’t stain anything, or don’t include any reagent with fluorophores (fluorescent proteins, injected dyes, etc.). Tissues and cells can have considerable autofluorescence, particularly (but not exclusively) in the green, cyan and blue region of the spectrum. Including this control can be very useful when determining whether any signals that are visualized in the experimental samples can be attributed to autofluorescence. Ideally, “real” staining will be significantly brighter than autofluorescence. If staining in your experimental sample is similar in intensity and distribution to the autofluorescence control, your sample does not have what you are staining for and/or the staining did not work.

No primary control

When performing immunostaining, prepare a slide where you don’t include the primary antibody but do include the secondary. Secondary antibodies can bind non-specifically to tissue or, if aggregated, lead to punctate staining throughout a sample. Including this control helps determine whether a signal visualized in the experimental sample can be explained by a problem with the secondary antibody. Ideally, “real” staining will be significantly brighter than the no primary control, and, if punctate, no puncta (or significantly fewer and dimmer puncta) will appear in the no primary control. If the staining in the experimental sample is similar in intensity and distribution to the no primary control, which is in turn brighter than the autofluroescence control, your sample does not have what you are staining for and/or your secondary antibody is binding non-specifically and/or aggregating.

No target control

Even if there is no non-specific binding of secondary antibodies, and autofluorescence is comparatively low, a strong signal in your experimental sample does not guarantee you are looking at your protein of interest. Primary antibodies are not necessarily as specific as advertised; the best control for this is to remove the putative target protein from the sample and test staining there. If there is still significant staining in this no target control (assuming autofluorescence and non-specific binding of secondary antibodies are not a problem), then the primary antibody is detecting something other than what it should. No target controls are frequently extremely expensive as they can involve making (or purchasing) knockdown or knockout cell lines or animals. In fact, they are sometimes impossible to procure because removing the target protein kills the animal or cells in which it is removed. Nevertheless, they are the only way to prove an antibody is staining what it is advertised to stain, and nothing else.

How to grow/mount cells and tissue

Slides and coverslips

Make sure to use #1.5 coverslips (0.17mm average thickness). This is what almost all of the objectives on our microscopes are designed for; other thicknesses will lead to noticeably worse images, particularly using high-resolution objectives. If you are not using #1.5 coverslips, this is one of the cheapest things you can do to improve how your samples look under a microscope.

Optical performance will be optimal at the coverslip/sample interface. Therefore, if applicable, try to grow your cells on coverslips (not slides).

Use a coverslip that does not extend all the way to the edge of the slide. Large coverslips prop up one edge of the slide and lead to images tilted in the z dimension on inverted microscopes (all of the confocals at MSL are inverted). In addition, they are difficult to seal properly with nail polish, which can lead to various problems (see below).


If you want to grow your cells in dishes, for best results use dishes with #1.5 glass coverslip bottoms. MatTek and ibidi are examples of companies that sell these products. If your cells have trouble growing on glass, coverslips coated with various substances are available. Alternatively, some dishes made by ibidi have proprietary plastic bottoms which are optically similar to #1.5 glass coverslips and to which certain cell types adhere to better than glass.

Chambered slides or coverglass

If you are using chambered coverglass holders (like the Nunc Lab-Tek II), ensure the coverglass is #1.5. If you are using chambered slides, remember to use a #1.5 coverslips at the end. Note that the added convenience of chambered slides comes with two problems:

  • Because cells are grown on the slide, and there are spacers left around the wells when the chambers are removed, the cells can be quite far from the coverslip after staining and mounting. As a result, optical performance is not optimal (see above). In some cases, the distance from the coverglass can be so large that it exceeds the working distance of some of our high resolution objectives.
  • Because the chambers for cells fill up most of the slide, you will need large coverslips that extend all the way to the edge of the slide. As a result, this may lead to the sample being tilted in an inverted microscope (like both of our confocals), as well as making the samples hard to seal (because there is little room between the edge of the coverslip and the edge of the slide).

Multiwell plates

If you want high resolution (for example, subcellular structures), use plates with #1.5 coverglass bottoms. If you don’t mind sacrificing some resolution for the convenience of a plastic-bottom multiwell plate, you will only be able to use certain lower-resolution objectives (low magnification, air only). If you decide to use plates, contact the MSL staff for further guidance.


Newer fluorophores tend to be brighter and more photostable than older ones (for example, AlexaFluor 488 is better than FITC).

Dyes that are good for flow cytometry (PE, APC) are not the best options for microscopy, due to their broad excitation spectra.

Keep in mind that there is more autofluorescence in the blue-green region of the spectrum. If your tissue is very autofluorescent, staining in the red or far red may be advantageous.

Avoid far red dyes (like AlexaFluor 647) if staining rare structures or proteins that will require a lot of hunting down to find in your sample. Far red dyes are not visible by eye. Therefore, you will need to find your structure of interest using the microscope detectors, which have a smaller field of view, slower refresh rate and lower dynamic range than the human eye-brain combination. This will make it harder and slower to find the particular things you are looking for. Thus, for these cases it is best to label those rare structures with a visible fluorophore.

Ask the MSL staff for help deciding among dyes and fluorescent proteins before starting your experiments. This could save you a lot of time and money in the long run.

Mounting media

If possible (when using coverslips and slides), use a mounting media to reduce bleaching. Our users have had good results with Prolong Gold, Prolong Diamond, Vectashield and Fluoromount, among others. Avoid mounting media with DAPI. Instead, stain for DAPI separately. Using DAPI in the mounting media can lead to significantly higher background overall (particularly in widefield microscopy) and problems in the DAPI staining. Note that Prolong Gold is not appropriate for use with fluorescent proteins (i.e. GFP, RFP, etc.), and Vectashield is not good for AlexaFluor 647.

Mounting media typically come in hardening or non-hardening formulations. The first tend to preserve the fluorescence for longer and flatten the samples as they harden. If you are not interested in the detailed 3D characteristics of your sample, this is typically not a problem, and makes samples thinner, and easier to visualize on a widefield microscope. If you DO care about the arrangement of cells or subcellular components in three dimensions do not use a hardening media OR do not let the hardening media actually harden. To preserve 3D information in the sample, put on the coverslip immediately after adding the mounting media, wipe off the excess mounting, and seal with nail polish. Give your nail polish time to harden before bringing samples to MSL (ie, don’t seal them 5 minutes before coming down). If coverslips are loose, this can cause mounting media to deposit on the objective making it difficult to image samples and even damage the objective.

Do not use mounting media in open containers such as multiwell plates, dishes or chambered coverglass; you will need a–comparatively–enormous volume of reagent, making it very expensive. Just keep your samples in a buffer (like PBS), in the dark and at 4 degrees celsius until imaging.

Light microscopy sample preparation services

There are several excellent cores on campus that can help you prepare samples:

Translation Pathology Laboratory

Animal Histopathology and Laboratory Medicine Core

Histology Research Core Facility