Immunofluorescence Microscopy: Immunofluorescence Staining of Paraffin-Embedded Tissue Sections

1. Set-up

1. The typical staining protocol involves the following steps:
   1. Re-hydrating the tissue sections on the slides using a series of graded ethanols.
   2. Incubating the tissue sections with a blocking buffer, which will help to block non-specific binding of antibody to the tissue and reduce background fluorescence.
   3. Removing the blocking buffer and incubating the section in primary antibody, at which time the antibody will bind its peptide target.
   4. Removing the primary antibody and washing the sections extensively in wash buffer.
   5. Incubating the sections with secondary antibody to allow binding to primary antibody, if the primary antibody is not directly labeled with a fluorophore and a secondary antibody is required.
   6. Washing the secondary antibody off the slides.
   7. Mounting the slides in mounting media and allowing them to dry prior to visualization on a fluorescent microscope.
2. The following items are needed: slide holders (glass or plastic), jars, pipettes, pap pen, humid chamber, cover slips, and mounting media with DAPI.
3. Xylene is used for the rehydration of slides. Xylene is hazardous and should be used in a fume hood along with appropriate PPE, including gloves and a lab coat.
4. Recipes for buffers, solutions, and reagents
   1. Graded ethanols
      Ethanol - 160 mL 200 proof EtOH and 40mL ddH₂O
      Ethanol - 140 mL 200 proof EtOH and 60mL ddH₂O
   2. Antigen retrieval solution
      10 mM sodium citrate, pH 6.0
   3. Blocking buffer
      100 µL Serum from host animal from which secondary antibody was made
      900 µL 1X PBS
      Note: This is the volume for 10 sections; adjust the volume for roughly 100 µL buffer per section, as needed.
   4. Wash buffer (1X PBS)

2. Protocol

1. Rehydration with xylenes and ethanols
   1. Place slides into a slide holder, and then submerge the slides in the 100% Xylene isomers solution, ensuring that the slides are entirely covered with solution.
   2. Incubate slides in the 100% Xylene isomers for 3 min. Repeat two times for a total of 3 separate incubations. Be sure to wipe off the slide rack with a paper towel prior to transferring to a new solution to minimize contamination.
      Note: It is recommended that these incubations are performed in three separate containers.
   3. Incubate slides in 100% EtOH for 2 min. Repeat two times for a total of 3 separate incubations.
      Note: It is recommended that these incubations are performed in three separate containers.
   4. Incubate slides in 95% EtOH for 2 min.
   5. Incubate slides in 80% EtOH for 2 min.
   6. Incubate slides in 70% EtOH for 2 min.
   7. Incubate slides in 1X PBS for 5 min.
2. (Optional) Antigen retrieval to unmask epitopes recognized by the primary antibody
   Note 1: This procedure is highly dependent upon the antibody used and it is recommended that initial optimization procedures are performed to determine the requirement for antigen retrieval.
   Note 2: This procedure was not performed with the F4/80 staining shown below. Requirement for antigen retrieval should be optimized with each new antibody.
   1. Place slides in a heat-resistant plastic or glass slide holder, and ensure the rack is filled with slides to ensure even heat distribution. Blank slides can be used if there are fewer samples than slots in the rack.
   2. Place rack in 1000 mL of antigen unmasking solution in 2L beaker - 10 mL antigen unmasking stock to 990mL water.
   3. Microwave on high for 20 minutes total, make sure slides remain covered with water.
   4. Cool slides for 20 minutes in beaker.
   5. Wash slide rack in slide holders containing ddH₂O for 5 min each. Repeat two times for a total of 3 separate incubations using fresh ddH₂O each time. The slides can be washed in the same slide holder during each wash.
   6. Incubate slides in 1X PBS for 5 min.
3. Circle sections with a pap pen. This will allow for the use of a minimal volume of buffers needed to cover the tissue sections. Do not allow tissue sections to dry out.
4. Add blocking buffer to each section. The amount of buffer needed to cover the section will vary depending upon the size of the section but could range from 25-500 µL. Enough buffer should be used to form a bead that covers the entire section surface including the edges.
   Note: The choice of blocking buffer may vary depending upon the antibody being used. For example, 10% serum from the host animal in which the secondary antibody was raised may be used to reduce non-specific binding of the secondary antibody (for example, normal goat serum can be used if the secondary antibody was raised in goat). Optimization of blocking buffer should be performed for each primary antibody. To stain mammary tumor sections with F4/80, 0.1 ml of 10% normal goat serum in PBS was used.
5. Incubate the sections in blocking buffer in a humidified chamber for 1 hour at room temperature or 4°C up to 24 hours. The humidified chamber ensures the sections do not dry out.
6. Remove the blocking buffer by draining it off the slide. Alternatively, the blocking buffer can be removed using a pipette, although take care not to touch the section with the pipette tip.
7. Dilute the primary antibody in blocking buffer.
   Note: The correct dilution will need to be determined for each antibody and sample type. Optimization would include performing a series of dilutions to identify optimal staining. To stain mammary tumor sections with F4/80, tissue sections were incubated overnight at 4°C in 0.1 mL rat anti-F4/80 antibody diluted 1:100 in 1% normal goat serum in PBS.
8. Add the primary antibody to each section and incubate in a humidified chamber for up to 16 hours (overnight) at 4°C. Leave at least one section in blocking buffer without primary antibody to serve as a control, which will help identify non-specific binding by the secondary antibody.
9. Drain the primary antibody or blocking buffer off the section, and wash sections with PBS by placing slides into slide rack and placing into a slide holder with 1x PBS. Wash 3 times for 10 min each time.
10. Dilute the secondary antibody 1:200 in blocking buffer. The dilution can be optimized depending upon the secondary antibody.
11. Add the secondary antibody to the all the sections, including the control, and incubate for 1 hour in a humidified chamber protected from light.
12. Drain the secondary antibody off the sections, and wash 3 times in PBS for 10 min each time.
13. Add 2-3 drops of mounting media containing DAPI to the slides and place a coverslip on the samples. If the mounting media is not a rapid-drying type of media, it may be necessary to seal the edges of the slide with melted paraffin wax or fingernail polish to prevent leakage and maintain the samples long-term.
14. Allow the slides to dry overnight in the dark, and image the sections using a fluorescent microscope.

3. Data Analysis and Results

1. Stained sections are analyzed using a fluorescent microscope. The specific details of image capture and analysis will depend upon the specifications of the microscope and the software used to perform the analysis. Typically, images can be taken as single color images and overlapped to generate multi-color images. Percent positive cells can be quantified by counting the number of positively stained cells and dividing by the total number of DAPI-stained cells within a section. For F4/80 staining of mammary tumor sections, using the software Leica Application Suite, version 3.8, under the Acquire tab and in Image Overlay acquisition mode, DAPI and RFP were both enabled. Exposure, Gain, and Gamma were adjusted (20.0 ms, 1.0x, and 1.53 respectively for DAPI and 944.2 ms, 1.0x, and 1.08 respectively for RFP), though this varies between experiments. The Acquire Overlay button was used to create overlay images of the DAPI and RFP exposures.
2. The image below (Figure 1) shows an example immunofluorescence image of a tumor section stained with F4/80, which detects an antigen on macrophages and other myeloid cells. The section was mounted using DAPI-containing mounting media and the nuclei are shown in blue.