Fluorescent IHC Staining of Frozen Tissue Protocol

This protocol is intended as a guide only, for full experimental details please read the reference provided.

Introduction

Fluorescent Immunohistochemistry/immunofluorescence is a technique which utilizes fluorescently labeled antibodies to visualize protein expression in a tissue section. Unlike immunocytochemistry (ICC) which utilizes fluorescence to visualize protein expression in cells, fluorescent immunohistochemistry/immunofluorescence uses fluorescent detection to determine protein expression in tissue, not cells. Fluorescent visualization allows easier multiplex analysis of multiple proteins by immunohistochemistry over traditional chromogenic detection system, such as DAB. The fluorescent immunohistochemistry/immunofluorescence protocol below is intended for the fluorescent visualization of protein expression in frozen tissue sections.

 

Tissue Preparation

 

Perfusion and Fixation  

Note: This portion of the protocol can be skipped if you are working with pre-mounted tissue slides. The technique described below utilizes formaldehyde-based fixation before the tissue is frozen and sectioned. Tissues can also be fixed following snap freezing and sectioning. See the next section of this protocol for more information on cryopreservation.

Optional: Alternate fixation methods may be used and result in better performance depending on the protein target. Please see Application Notes section on the datasheet of the product you are using for further information, if available.

  1. Fix tissue by perfusing the animal with freshly prepared 4% paraformaldehyde or by immersing it in 4% paraformaldehyde for 4-24 hours at room temperature. Fixation temperature and time may require optimization depending on the tissue type and size.

  2. Cryoprotect the tissue by directly perfusing a sucrose solution or by first dissecting the tissue and allowing it to sink overnight in a 30% sucrose/ 70% fixative solution.

  3. Embed tissue in OCT cryostat sectioning medium and store at -80° C until ready for sectioning. Tissue can be safely stored for 6-12 months.

  4. When ready for sectioning, move the embedded tissue directly into the cryostat and use OCT medium to mount it to the chuck. Allow the temperature of the tissue to equilibrate with the cryostat.

  5. Cut the tissue in 5-20 µm thick sections. Mount tissue sections onto gelatin or poly-L-lysine coated slides by placing the cold sections onto warm slides. Slides can be safely stored for 6-12 months at -80° C until ready for staining.

 

Cryopreservation

Note: This portion of the protocol can be skipped if you are working with pre-mounted tissue slides. The technique described below utilizes frozen tissues that are fixed after snap freezing and sectioning with a cryostat.

 

  1. After dissection, immediately snap freeze tissue with isopentane cooled by liquid nitrogen. To do this, prepare a small dewar of liquid nitrogen. Take an aluminum can cut in half and fill with isopentane. Float the can in the liquid nitrogen until the isopentane is cooled. Quickly dissect the tissue, wrap in aluminum foil, and place in the cooled isopentane. After the tissue is frozen, place it in dry ice and move to -80° C until ready for cutting.

  2. Embed tissue in OCT compound by slowly layering the compound so that the tissue does not thaw. Move the embedded tissue directly into the cryostat and use OCT medium to mount it to the chuck. Allow the temperature of the tissue to equilibrate with the cryostat.

  3. Cut the tissue in 5-20 µm thick sections. Mount tissue sections onto gelatin or poly-L-lysine coated slides by placing the cold sections onto warm slides. Slides can be safely stored for 6-12 months at -80° C until ready for fixing. Uncut tissue can be restored at -80°C.

  4. Remove slides from freezer and fix with cold fixative (acetone or methanol) for 10 minutes. Proceed to staining


Blocking Non-specific Binding 

  1. Warm slides to room temperature and wash slides twice with PBS. Note: Fixation may result in epitope masking and non-specific background that can impact specific labeling. If necessary, a protocol for antigen retrieval can be performed at this time. However, many antigen retrieval methods are too harsh on cryostat cut tissue sections.

  2. Draw a circle on the slide around the tissue with a hydrophobic barrier pen.

  3. To permeabilize the tissue/cells, wash the sections twice for 10 minutes each with permeabilization buffer containing 1% animal serum and 0.4% Triton X-100 in PBS (PBS-T).

  4. Block non-specific binding by incubating the tissue sections with 5% serum in PBS-T for 30 minutes at room temperature.

Tip: The species of the animal serum used in permeabilization and blocking buffers is dependent on the host of your secondary antibody. (e.g. when using a goat anti-mouse secondary, use goat serum).



Antibody Staining 

  1. Add primary antibody diluted in 1% animal serum PBS (with or without 0.05-0.1 % Triton X 100) to the sections and incubate at room temperature for 1-2 hours. Then store overnight at 4°C in humidified chamber. Use the recommended dilution of the antibody specified on the datasheet. If not specified, the typical starting dilution can be 2-5 µg/ml. For more information on primary antibody selection, please read our IHC Primary Antibody Selection Guide.

  2. Wash sections twice with 1% serum PBS-T for 10 minutes each.

  3. Add a fluorescent label conjugated secondary antibody diluted with 1% serum in PBS (with or without 0.05-0.1% Triton X 100) and incubate at room temperature for 1-2 hours. Use the recommended dilution of the antibody specified on the datasheet. Note: For help selecting the optimal secondary antibody, please read our Secondary Antibody Handbook.

  4. Wash sections twice with 1% serum PBS-T for 10 minutes each.



Double or Nuclear Labeling (Optional)

  1. Double Labeling: If using a second primary antibody and appropriately matched secondary, repeat step 5-8.

  2. Nuclear Labeling: After application of all primary antibodies, DNA binding dyes such as DAPI can be applied to the slides. After dye incubation, wash the slides once for 5 minutes with PBS.



Detection

  1. Tap off excess wash buffer and apply one drop of ant-fade mounting medium to the slide.

  2. Place a coverslip on the tissue sections and circle the edges of the coverslip with clear fingernail polish to prevent the cells from drying. Allow the polish to air dry.

  3. Slides may now be examined under a microscope with the appropriate fluorescent filter sets. Be sure to limit slide exposure to light to prevent photobleaching.

  4. Slides can be stored between -20°C and 4°C in a dark slide box or slide book.
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