Immunofluorescence Microscopy: Immunofluorescence Staining of Paraffin-Embedded Tissue Sections
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Vue d'ensemble
Source: Thomas Chaffee1, Thomas S. Griffith2,3,4, and Kathryn L. Schwertfeger1,3,4
1 Department of Lab Medicine and Pathology, University of Minnesota, Minneapolis, MN 55455
2 Department of Urology, University of Minnesota, Minneapolis, MN 55455
3 Masonic Cancer Center, University of Minnesota, Minneapolis, MN 55455
4 Center for Immunology, University of Minnesota, Minneapolis, MN 55455
Pathologic analyses of tissue sections can be used to obtain a better understanding of normal tissue structure and contribute to our understanding of mechanisms of disease. Tissue biopsies, either from patients or from experimental in vivo models, are often preserved by fixing in formalin or paraformaldehyde and embedding in paraffin wax. This allows for long-term storage and for the tissues to be sectioned. Tissues are cut into thin (5 µm) sections using a microtome and the sections are adhered to glass slides. The tissues sections can be stained with antibodies, which allow for the detection of specific proteins within the tissue sections. Staining with antibodies conjugated to fluorophores (also known as fluorochromes) – compounds that emit light at specific wavelengths when excited by a laser – is known as immunofluorescence. The ability to detect proteins within a section can provide information such as cell type heterogeneity within the tissue, activation of specific signaling pathways, and expression of biomarkers. Depending on the fluorophores used and type of microscope available for analysis, multiple colors can be used, which allows for multiplexed analysis of targets.
The following protocol outlines the basic steps involved in immunofluorescence staining of paraffin embedded tissue sections. It is important to note that this protocol will not include any details on the fixation of tissue, process of paraffin embedding, or sectioning of the tissues. Once tissues have been sectioned and placed on glass slides, they are rehydrated through a series of graded ethanol (EtOH) incubations. The sections are incubated with a blocking reagent to reduce non-specific binding of antibody to the tissue section. The sections are then incubated with a primary antibody that may or may not be directly labeled with a fluorophore. If the primary antibody is not directly labeled, the sections are then incubated with a secondary antibody labeled with a fluorophore. Different antibodies may require different staining conditions, thus suggestions for optimization of antibodies are included. Following washing to remove all unbound antibody, the slides are mounted with media containing DAPI to fluorescently label the nucleus. Once the mounting media has dried, the slides can be imaged using a microscope with lasers that can detect the different fluorophores.
Procédure
1. Set-up
- The typical staining protocol involves the following steps:
- Re-hydrating the tissue sections on the slides using a series of graded ethanols.
- 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.
- Removing the blocking buffer and incubating the section in primary antibody, at which time the antibody will bind its peptide target.
- Removing the primary antibody and washing the sections extensively in wash buffer.
- 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.
- Washing the secondary antibody off the slides.
- Mounting the slides in mounting media and allowing them to dry prior to visualization on a fluorescent microscope.
- The following items are needed: slide holders (glass or plastic), jars, pipettes, pap pen, humid chamber, cover slips, and mounting media with DAPI.
- 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.
- Recipes for buffers, solutions, and reagents
- Graded ethanols
Ethanol – 160 mL 200 proof EtOH and 40mL ddH2O
Ethanol – 140 mL 200 proof EtOH and 60mL ddH2O - Antigen retrieval solution
10 mM sodium citrate, pH 6.0 - 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. - Wash buffer (1X PBS)
- Graded ethanols
2. Protocol
- Rehydration with xylenes and ethanols
- 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.
- 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. - 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. - Incubate slides in 95% EtOH for 2 min.
- Incubate slides in 80% EtOH for 2 min.
- Incubate slides in 70% EtOH for 2 min.
- Incubate slides in 1X PBS for 5 min.
- (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.- 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.
- Place rack in 1000 mL of antigen unmasking solution in 2L beaker – 10 mL antigen unmasking stock to 990mL water.
- Microwave on high for 20 minutes total, make sure slides remain covered with water.
- Cool slides for 20 minutes in beaker.
- 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.
- Incubate slides in 1X PBS for 5 min.
- 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.
- 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. - 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.
- 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.
- 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. - 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.
- 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.
- Dilute the secondary antibody 1:200 in blocking buffer. The dilution can be optimized depending upon the secondary antibody.
- Add the secondary antibody to the all the sections, including the control, and incubate for 1 hour in a humidified chamber protected from light.
- Drain the secondary antibody off the sections, and wash 3 times in PBS for 10 min each time.
- 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.
- Allow the slides to dry overnight in the dark, and image the sections using a fluorescent microscope.
3. Data Analysis and Results
- 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.
- 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.
The function of a protein in a cell is largely dependent on its proper localization within the cell. Immunofluorescence microscopy is a method by which a protein can be visualized inside cells using fluorescent dyes. A fluorescent dye is a fluorophore, that is a molecule that absorbs light energy at a specific wavelength by a process called excitation, and then immediately releases the energy at a different wavelength, known as emission.
Fluorescent dyes are conjugated to a target-specific antibody and introduced into cultured cells or tissue by immunostaining. When this primary antibody binds to the protein of interest, the protein gets labeled with the fluorescent dye. Alternatively, the fluorescent dye can be conjugated to a secondary antibody, instead of the primary antibody, and the secondary antibody binds to the protein primary antibody complex to label the target. After that, the sample is sealed in an antifade mounting medium to preserve the fluorescence labeling and is then ready for imaging on a fluorescence microscope.
A fluorescence microscope is equipped with a powerful light source. The light beam first passes through an excitation filter, which allows only the light at the excitation wavelength to pass through. The excitation light then reaches a specialized mirror, called a dichroic mirror or a beam splitter, which is designed to selectively reflect the excitation wavelength towards an objective lens. The lens then focuses the light onto a small region in the sample. Upon reaching the sample, the light excites the fluorophores, which then emit the light energy at a different wavelength. This light is transmitted back through the objective lens to the dichroic mirror. Since the emission wavelength is different from the excitation wavelength, the dichroic mirror allows the emission light to pass through. Then, it goes through a second filter, called the emission filter, which eliminates light from any other wavelengths that may be present. After that, the light rays now reach the eyepiece or camera, where they present a magnified image created from the emitted light from the specific fluorophores. This image represents the location of the protein of interest within the cell.
DNA binding fluorescent dyes are often used along with immunostaining to label nuclei as a point of reference within the cells. Multiple different fluorophores, with different excitation emission wavelengths, can be used for different proteins within the same sample to compare localization of the proteins.
This video demonstrates the procedure for immunofluorescent staining of a protein of interest in a tissue sample followed by imaging the sample on a fluorescence microscope.
Before beginning the staining process, the sections, which were dehydrated during the embedding process, need to be rehydrated. To do this, first, place the slides into a slide holder and then completely submerge the slides in 100% Xlene isomers. Allow the slides to incubate for three minutes. Then, remove the slides from the container, wipe off any excess Xylene with a paper towel, and place them into a new Xylene bath in a fresh container, for a further three minutes. Repeat this incubation each time in a new container with fresh Xylene and wiping down the slides with paper towels before transferring to the new container, for a total of three incubations. Next, incubate the sections in a series of graded ethanol solutions, starting with 100% ethanol for two minutes. Wipe off the slide rack with a paper towel, and transfer the slides to a new container of 100% ethanol for another two minutes. Continue the cycle of washing, wiping excess ethanol with a paper towel, and transferring the slides to a new bath, following the indicated concentrations of ethanol for the specified time. After the final ethanol wash, shake off the excess solution and incubate the slides in 1X PBS for five minutes.
To begin the staining process, first, circle the sections with a PAP pen to identify the minimal area that the buffers need to cover. Once the sections are clearly marked on the slide, add 100 microliters of blocking buffer to each slide, making sure to cover the entire section surface. After the tissues are covered in blocking buffer, place the slides in a humidified chamber. Leave the slides to incubate for one hour at room temperature.
Following the desired incubation time, remove the blocking buffer by draining it off the slide. Next, dilute the primary antibody in blocking buffer. For a 1:100 dilution, add 990 microliters of blocking buffer to a 1.5 milliliter centrifuge tube, followed by 10 microliters of the primary antibody to the same tube. Label one slide as a control and then add 100 microliters of blocking buffer. This control will help identify any non-specific binding of the secondary antibody. Now, add 100 microliters of primary antibody buffer to the remaining slides. Incubate the sections overnight in a humidified chamber at four degrees celsius, in the dark.
Following the overnight incubation, remove the sections from the chamber and drain the primary antibody off each slide and the blocking buffer from the control. Place the slides into a slide rack and then wash them three times in 1X PBS for ten minutes each. While the slides are washing in 1X PBS, dilute the secondary antibody in blocking buffer. For a 1:200 dilution, add 995 microliters of blocking buffer to a 1.5 milliliter tube, followed by five microliters of the secondary antibody to the same tube. Add the secondary antibody to all of the sections, including the control, and incubate them for one hour in a humidified chamber protected from light. When the timer sounds, remove the slides from the incubator. Drain the secondary antibody off the sections. Once the secondary antibody is removed, place the slides in a slide rack and then completely submerge the slides in 1X PBS for 10 minutes, protected from light. Repeat this wash three times, using fresh 1X PBS for each wash. Following the washes, add two to three drops of mounting media containing DAPI to each slide and place a glass coverslip on the samples. Allow the slides to dry overnight in a dark place before imaging the sections using a fluorescent scope.
During imaging, the details of image capture will depend upon the specific microscope and software available. However, in this particular example, the software Leica Application Suite, Version 3. 8, is used to perform the analysis. Using this program, click on the Acquire tab and in Image Overlay Acquisition mode, enable both DAPI and RFP. Next, adjust the Exposure, Gain, and Gamma for both DAPI and RFP, by taking an initial using the default settings defined by the software, and then optimizing for brightness by modifying the exposure time and the gain, keeping in mind that minimal optimal settings are desirable to avoid image saturation and photobleaching of the samples. Gamma can be modified to optimize darker areas of an image.
Once the settings are adjusted, press the Acquire Overlay button to create overlay images of the DAPI and RFP exposures. This example image, captured using the demonstrated technique, shows a mouse mammary tumor section, stained with the antibody F4/80, which detects an antigen, depicted in red, on macrophages and other myeloid cells. Since DAPI-containing mounting media was used, nuclei are shown in blue. The data from the imaging will provide information regarding the intensity and localization of the protein within the tissue section.
For example, in the image of the tumor stained with F4/80, cell surface staining of this antigen is observed. These data can also provide information regarding the frequency of specific cell populations within the tissue section. This can be quantified by counting the number of positively stained cells, here shown in red, and comparing that with the total cell population, shown in blue, and calculating the frequency using the following equation.
Résultats
Figure 1: F4/80 staining of a mammary tumor section. Following fixation, a mouse mammary tumor was sectioned and stained with anti-F4/80 and mounted using a DAPI-containing mounting media. Staining is shown by cell surface F4/80 staining in red. Please click here to view a larger version of this figure.
The data obtained from the imaging will provide information regarding the intensity and localization of expression of the protein-of-interest within the tissue section. Depending upon the protein being examined, these data could also provide information regarding the frequency of specific cell populations within the tissue section. This can be quantified by counting the number of positively stained cells and comparing with total cell population.
Applications and Summary
Immunofluorescence allows for the investigation of protein expression and localization within the context of a tissue section. This technique can be used to understand how tissues change in the context of disease by examining protein localization or cell number in normal and diseased tissues. Changes in localization or in expression patterns can be determined and linked to specific attributes of the samples.
References
- Im K, Mareninov S., Diaz MFP, Yong WH. An Introduction to Performing Immunofluorescence Staining. Yong W. (eds) Biobanking. Methods in Molecular Biology. 1897, Humana Press, New York, NY (2019)
- Ramos-Vara JA. Principles and Methods of Immunohistochemistry. Gautier JC. (eds) Drug Safety Evaluation. Methods in Molecular Biology. 1641, Humana Press, New York, NY (2017)
- Donaldson JG. Immunofluorescence Staining. Current protocols in Cell Biology. 69 (1):1 4.3.1-4.3.7. (2015)
Transcription
The function of a protein in a cell is largely dependent on its proper localization within the cell. Immunofluorescence microscopy is a method by which a protein can be visualized inside cells using fluorescent dyes. A fluorescent dye is a fluorophore, that is a molecule that absorbs light energy at a specific wavelength by a process called excitation, and then immediately releases the energy at a different wavelength, known as emission.
Fluorescent dyes are conjugated to a target-specific antibody and introduced into cultured cells or tissue by immunostaining. When this primary antibody binds to the protein of interest, the protein gets labeled with the fluorescent dye. Alternatively, the fluorescent dye can be conjugated to a secondary antibody, instead of the primary antibody, and the secondary antibody binds to the protein primary antibody complex to label the target. After that, the sample is sealed in an antifade mounting medium to preserve the fluorescence labeling and is then ready for imaging on a fluorescence microscope.
A fluorescence microscope is equipped with a powerful light source. The light beam first passes through an excitation filter, which allows only the light at the excitation wavelength to pass through. The excitation light then reaches a specialized mirror, called a dichroic mirror or a beam splitter, which is designed to selectively reflect the excitation wavelength towards an objective lens. The lens then focuses the light onto a small region in the sample. Upon reaching the sample, the light excites the fluorophores, which then emit the light energy at a different wavelength. This light is transmitted back through the objective lens to the dichroic mirror. Since the emission wavelength is different from the excitation wavelength, the dichroic mirror allows the emission light to pass through. Then, it goes through a second filter, called the emission filter, which eliminates light from any other wavelengths that may be present. After that, the light rays now reach the eyepiece or camera, where they present a magnified image created from the emitted light from the specific fluorophores. This image represents the location of the protein of interest within the cell.
DNA binding fluorescent dyes are often used along with immunostaining to label nuclei as a point of reference within the cells. Multiple different fluorophores, with different excitation emission wavelengths, can be used for different proteins within the same sample to compare localization of the proteins.
This video demonstrates the procedure for immunofluorescent staining of a protein of interest in a tissue sample followed by imaging the sample on a fluorescence microscope.
Before beginning the staining process, the sections, which were dehydrated during the embedding process, need to be rehydrated. To do this, first, place the slides into a slide holder and then completely submerge the slides in 100% Xlene isomers. Allow the slides to incubate for three minutes. Then, remove the slides from the container, wipe off any excess Xylene with a paper towel, and place them into a new Xylene bath in a fresh container, for a further three minutes. Repeat this incubation each time in a new container with fresh Xylene and wiping down the slides with paper towels before transferring to the new container, for a total of three incubations. Next, incubate the sections in a series of graded ethanol solutions, starting with 100% ethanol for two minutes. Wipe off the slide rack with a paper towel, and transfer the slides to a new container of 100% ethanol for another two minutes. Continue the cycle of washing, wiping excess ethanol with a paper towel, and transferring the slides to a new bath, following the indicated concentrations of ethanol for the specified time. After the final ethanol wash, shake off the excess solution and incubate the slides in 1X PBS for five minutes.
To begin the staining process, first, circle the sections with a PAP pen to identify the minimal area that the buffers need to cover. Once the sections are clearly marked on the slide, add 100 microliters of blocking buffer to each slide, making sure to cover the entire section surface. After the tissues are covered in blocking buffer, place the slides in a humidified chamber. Leave the slides to incubate for one hour at room temperature.
Following the desired incubation time, remove the blocking buffer by draining it off the slide. Next, dilute the primary antibody in blocking buffer. For a 1:100 dilution, add 990 microliters of blocking buffer to a 1.5 milliliter centrifuge tube, followed by 10 microliters of the primary antibody to the same tube. Label one slide as a control and then add 100 microliters of blocking buffer. This control will help identify any non-specific binding of the secondary antibody. Now, add 100 microliters of primary antibody buffer to the remaining slides. Incubate the sections overnight in a humidified chamber at four degrees celsius, in the dark.
Following the overnight incubation, remove the sections from the chamber and drain the primary antibody off each slide and the blocking buffer from the control. Place the slides into a slide rack and then wash them three times in 1X PBS for ten minutes each. While the slides are washing in 1X PBS, dilute the secondary antibody in blocking buffer. For a 1:200 dilution, add 995 microliters of blocking buffer to a 1.5 milliliter tube, followed by five microliters of the secondary antibody to the same tube. Add the secondary antibody to all of the sections, including the control, and incubate them for one hour in a humidified chamber protected from light. When the timer sounds, remove the slides from the incubator. Drain the secondary antibody off the sections. Once the secondary antibody is removed, place the slides in a slide rack and then completely submerge the slides in 1X PBS for 10 minutes, protected from light. Repeat this wash three times, using fresh 1X PBS for each wash. Following the washes, add two to three drops of mounting media containing DAPI to each slide and place a glass coverslip on the samples. Allow the slides to dry overnight in a dark place before imaging the sections using a fluorescent scope.
During imaging, the details of image capture will depend upon the specific microscope and software available. However, in this particular example, the software Leica Application Suite, Version 3. 8, is used to perform the analysis. Using this program, click on the Acquire tab and in Image Overlay Acquisition mode, enable both DAPI and RFP. Next, adjust the Exposure, Gain, and Gamma for both DAPI and RFP, by taking an initial using the default settings defined by the software, and then optimizing for brightness by modifying the exposure time and the gain, keeping in mind that minimal optimal settings are desirable to avoid image saturation and photobleaching of the samples. Gamma can be modified to optimize darker areas of an image.
Once the settings are adjusted, press the Acquire Overlay button to create overlay images of the DAPI and RFP exposures. This example image, captured using the demonstrated technique, shows a mouse mammary tumor section, stained with the antibody F4/80, which detects an antigen, depicted in red, on macrophages and other myeloid cells. Since DAPI-containing mounting media was used, nuclei are shown in blue. The data from the imaging will provide information regarding the intensity and localization of the protein within the tissue section.
For example, in the image of the tumor stained with F4/80, cell surface staining of this antigen is observed. These data can also provide information regarding the frequency of specific cell populations within the tissue section. This can be quantified by counting the number of positively stained cells, here shown in red, and comparing that with the total cell population, shown in blue, and calculating the frequency using the following equation.