1. Set-up
2. Protocol
3. Data Analysis and Results
Source: Thomas Chaffee1, Thomas S. Griffith2,3,4, and Kathryn L. Schwertfeger1,3,4
1 Department of Lab Medicine and Pathology, University of Minnesota…
1. Set-up
2. Protocol
3. Data Analysis and Results
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.
View the full transcript and gain access to JoVE Science Education videos
Q1: What is a fluorophore and how does it work in immunofluorescence microscopy?
A fluorophore is a molecule that absorbs light energy at a specific wavelength through excitation, then immediately releases the energy at a different wavelength through emission. Fluorophores are conjugated to antibodies and introduced into tissue samples through immunostaining. When the antibody binds to the target protein, the fluorophore labels it, allowing visualization under a fluorescence microscope.
Q2: How does a fluorescence microscope separate excitation and emission light?
A fluorescence microscope uses specialized optical components to separate wavelengths. Excitation light passes through an excitation filter, then reflects off a dichroic mirror toward the objective lens, which focuses it on the sample. Emitted light from excited fluorophores returns through the objective and dichroic mirror, which allows only emission wavelengths to pass. An emission filter removes unwanted wavelengths before the image reaches the eyepiece or camera.
Q3: Why is rehydration necessary before immunofluorescence staining of paraffin-embedded tissue?
Tissue sections are dehydrated during the paraffin embedding process. Before staining, sections must be rehydrated through graded ethanol incubations and PBS washes to restore tissue hydration and prepare it for antibody binding. This step ensures proper antibody penetration and reduces non-specific binding during the immunostaining procedure.
Q4: What is the purpose of a blocking buffer in immunofluorescence staining?
Blocking buffer reduces non-specific binding of antibodies to the tissue section. The tissue is incubated with blocking buffer for one hour at room temperature before primary antibody application. This step prevents the antibody from binding to sites other than the target protein, improving staining specificity and reducing background fluorescence in the final image.
Q5: Why is a control slide included in immunofluorescence staining procedures?
A control slide is treated identically to experimental slides but receives only blocking buffer instead of primary antibody. This control helps identify any non-specific binding of the secondary antibody to the tissue. By comparing the control to experimental slides, researchers can distinguish true target protein localization from background fluorescence caused by non-specific antibody interactions.
Q6: How can multiple proteins be visualized simultaneously in a single tissue sample?
Multiple different fluorophores with distinct excitation and emission wavelengths can be used to label different proteins within the same sample. DNA binding fluorescent dyes like DAPI label nuclei as a reference point. By using multiple fluorophores, researchers can compare the localization of different proteins and analyze their spatial relationships within the tissue section.
Q7: What information can immunofluorescence imaging provide about tissue samples?
Immunofluorescence imaging provides data on protein intensity and localization within tissue sections, revealing cell type heterogeneity, activation of signaling pathways, and biomarker expression. Researchers can quantify the frequency of specific cell populations by counting positively stained cells and comparing them to the total cell population, calculated using a simple ratio equation.
Chapters in this video
0:01
Concepts
3:12
Rehydrating the Sections
4:46
Staining the Tissue Sections
7:43
Results
Videos from this collection: