The following protocol has been developed and optimized by R&D Systems IHC/ICC laboratory for fluorescent IHC experiments using paraffin-embedded tissue samples.
This protocol provides a basic guide for the fixation, microtome sectioning, and staining of paraffin-embedded tissue samples. Each investigator must determine the precise experimental conditions required to generate a strong and specific signal for each antigen of interest. If R&D Systems primary antibodies are employed, please refer to the product data sheets to obtain the recommended working dilutions. In this protocol, signal visualization is achieved using R&D Systems NorthernLights™ range of fluorescent secondary antibodies and reagents. For all other reagents, please follow the manufacturer's instructions.
Please read protocol in its entirety before beginning.
Protocol for Fixing and Sectioning Paraffin-embedded Tissues
Formaldehyde Fixative Solution: 85 mM Na2HPO4, 75 mM KH2P04, 4% paraformaldehyde, and 14% (v/v) saturated picric acid, pH 6.9. (Picric acid is optional, enhances preservation of morphology in some tissues)
Ethanol: 100%, 90%, 70%
Wash Buffer: 1X PBS (0.137 M NaCl, 0.05 M NaH2PO4, pH 7.4)
Xylene (mixed isomers)
To preserve tissue morphology and retain the antigenicity of the target molecules, fix the tissue by vascular perfusion with 500 - 700 mL of Formaldehyde Fixative Solution. Note: When it is not possible to fix by perfusion, dissected tissue may be fixed by immersion in a 10% formalin solution for 4-8 hours at room temperature. It is commonly accepted that the volume of fixative should be 50 times greater than the size of the immersed tissue. Avoid fixing the tissue for greater than 24 hours since tissue antigens may either be masked or destroyed. Note: R&D Systems scientists perfuse fix all rodent tissue with the exception of lung, spleen, and embryonic tissue, which are immersion fixed.
Dehydrate Tissues. Because paraffin is immiscible with water, tissue must be dehydrated before adding molten paraffin wax.
Immerse the tissue in 70% ethanol three times for 30 minutes each at room temperature.
Immerse the tissue in 90% ethanol two times for 30 minutes each at room temperature.
Immerse the tissue in 100% ethanol three times for 30 minutes each at room temperature.
Immerse the tissue in xylene (mixed isomers) three times for 20 minutes each at room temperature.
Embed the tissue in paraffin at 58 °C. Tissues can be embedded into paraffin using specialized automated tissue processing systems.
Cut 5 - 15 µm thick tissue sections using a rotary microtome.
Immerse the slides in xylene (mixed isomers) 2 times for 10 minutes each.
Immerse the slides in 100% ethanol 2 times for 10 minutes each.
Immerse the slides in 95% ethanol for 5 minutes.
Immerse the slides in 70% ethanol for 5 minutes.
Immerse the slides in 50% ethanol for 5 minutes.
Rinse the slides with deionized H2O.
Rehydrate the slides with wash buffer for 10 minutes. Drain the excess wash buffer.
Note: Excessive fixation may result in the masking of an epitope and strong non-specific background signal that can obscure specific labeling. If necessary, an antigen retrieval protocol can be performed at this time.
Surround the tissue with a hydrophobic barrier using a barrier pen.
Block non-specific staining between the primary antibodies and the tissue, by incubating in blocking buffer (1% horse serum in PBS) for 30 minutes at RT.
Apply primary antibodies diluted in Incubation Buffer according to manufacturer’s instructions. For fluorescent IHC staining of paraffin-embedded tissue sections using R&D Systems antibodies, it is recommended to incubate overnight at 2-8 °C. This incubation regime allows for optimal specific binding of antibodies to tissue targets and reduces non-specific background staining. These variables may need to be optimized for your system. Note: Appropriate controls are critical for the accurate interpretation of IHC/ICC results. All IHC/ICC experiments should include a negative control using the incubation buffer with no primary antibody to identify non-specific staining of the secondary reagents. Additional controls can be employed to support the specificity of staining generated by the primary antibody. These include absorption controls, isotype matched controls (for monoclonal primary antibodies), and tissue type controls.
Wash slides 3 times for 15 minutes each in wash buffer.
Incubate with the NorthernLights secondary antibody diluted in Incubation Buffer according to the manufacturer’s instructions. Recommended incubation with NorthernLights secondary antibody is for 30 - 60 minutes at room temperature. From this step forward, samples should be protected from light. Note: R&D Systems NorthernLights fluorescent secondary antibodies and streptavidin conjugates are bright, resistant to photobleaching and are ideal for multi-color fluorescence microscopy. Note: If a biotinylated antibody was used in step 4, apply the appropriate NorthernLights Streptavidin conjugate in step 6.
Wash slides 3 times for 15 minutes each in wash buffer.
Add 300 µL of the diluted DAPI solution to each well and incubate 2-5 minutes at room temperature. DAPI binds to DNA and is a convenient nuclear counterstain. It has an absorption maximum at 358 nm and fluoresces blue at an emission maximum of 461 nm. Note: DAPI counterstain can obscure visualization of targets localized in cell nuclei.
Rinse 1 time with PBS.
Mount with an anti-fade mounting media.
Visualize using a fluorescence microscope. Note: Initial IHC/ICC studies often require further optimization and/or additional troubleshooting steps.