Confocal Fluorescence Microscopy: A Technique to Determine the Localization of Proteins in Mouse Fibroblasts
Source: Dominique R. Bollino1, Eric A. Legenzov2, Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland School of Medicin…
1. Materials
Buffers
- Wash buffer: 1 X sterile phosphate-buffered saline (PBS) without calcium or magnesium
- Fixation buffer: 1% paraformaldehyde in PBS
- Permeabilization buffer: 0.1% Triton X-100 in PBS
- Blocking buffer: 1% bovine serum albumin in PBS
- Cell growth medium: DMEM supplemented with 10% fetal bovine serum (FBS), penicillin/streptomycin, and L-glutamine
Equipment
- Laminar flow hood
- Humidified incubator (37°C, 5% CO2)
- Confocal laser scanning microscope; here, Nikon Eclipse Ti laser
Materials and Reagents
- Chambered cell culture slides
- Anti-fade mounting media with DAPI (for staining nuclei)
- Microscope cover glass
- Delicate task wipes
- Pipettors and tips
Assay Specific Reagents
- Adherent cells (primary cells or cell lines); here, mouse fibroblasts transfected with CD1d were used (LCD1).
- Primary antibodies to detect cellular targets; here, rat anti-mouse CD107a (LAMP-1) and mouse anti-mouse CD1d were used.
- Fluorophore-conjugated secondary antibodies specific to primary antibody isotypes; here anti-rat IgG conjugated to Alexafluor 488 and anti-mouse IgG conjugated to Alexafluor 647 were used.
2. Protocol
Preparing for Antibody Staining
Seeding cells
- Resuspend the cells of interest in growth media.
- Then, seed 500 µL of the cell suspension/per well into the wells of a 4-well chamber slide. (Here, LCD1 cells were seeded at 2.5x105 cells/chamber in 500 µL of growth media. The seeding density may vary between cell lines).
- Incubate the chamber slide overnight in a 5% CO2 incubator at 37°C, to allow cells to adhere to the glass.
- Next day, aspirate the media from each well and then wash the cells 1X with 500 µL PBS.
Fixation
- To fix the cells, add 500 µL 1% paraformaldehyde solution into each well and incubate for 15 min at room temperature.
- After the incubation, collect the paraformaldehyde into an appropriate hazardous liquid waste container.
- Then, wash the cells 3 times with 500 µL PBS to remove any remnants of the fixative.
Permeabilization
- Permeabilize the cells by incubating with 500 µL permeabilization buffer/well for 15 minutes at room temperature.
- Then, wash the cells briefly 3 times with 500 µL of PBS.
Blocking
- Incubate the cells in each well with 500 µL blocking buffer for 1 hour at 4°C, to block non-specific antibody binding.
Primary Antibody Incubation
- Aspirate the blocking buffer from the slide chambers.
- Then, add 500 µL of diluted primary antibody solution to cells. (Here, anti-CD107a (LAMP-1) was diluted 1:500 and anti-CD1d was used undiluted (1H6 monoclonal antibody was kindly provided by Dr. Randy Brutkiewicz)).
- Incubate the slides overnight at 4°C.
Note: If probing for more than one target, make sure that the primary antibodies are different isotypes. Recommended antibody concentrations vary between manufacturers and should be titrated prior to use.
Secondary Antibody Incubation
- Aspirate the primary antibody solution from the wells.
- Wash the well chambers 4 times with 500 µL PBS.
- Then, add 500 µL of the diluted secondary antibody solution to each well. (Here, both the secondary antibodies- anti-mouse IgG Alexafluor 647 and anti-rat IgG Alexafluor 488 were diluted 1:2000 in the blocking buffer).
- Incubate at room temperature for 1 h in the dark.
- After the incubation, aspirate the secondary antibody solution.
- Wash the chambers 4 times with 500 µL PBS to remove any unbound secondary antibody.
Note: Recommended antibody concentrations vary between manufacturers and should be titrated prior to use. If probing for more than one target, secondary antibodies must be conjugated to different fluorophores with unique excitation/emission spectrums. Also keep in mind the excitation/emission of the nuclear counterstain (i.e. DAPI) while selecting the fluorophores. Fluorophore selection may be impacted by the laser configuration of the confocal microscope used. The laser configuration of the machine will dictate which fluorophores are suitable for the experiment.
3. Mounting Coverslips
- First, carefully remove the chambers from the slide.
- Then, hold the slide at angle over a delicate task wipe and remove the fluid from the edges without touching the cells.
- Add 1 drop of antifade mounting medium, containing the nuclear stain DAPI, onto each section of cells.
- Next, place a 20 mm x 60 mm coverslip over the slide by holding it over the edges using the fingertips. (Avoid bubble formation over the cells, as they interfere with imaging).
- Wipe off any extra mounting medium on the sides with a delicate task wipe and store the slides in the dark at room temperature up to a week.
4. Confocal Imaging
Image samples on a confocal laser scanning microscope. For data shown in Figure 2, the Nikon Eclipse Ti A1R was used with NIS Elements Advanced Research software. The following section details the procedure for capturing images using the aforementioned software.
- To begin imaging the cells, open the 'NIS Elements Advanced Research software' by clicking on the 'NIS software icon.'
- Next, on the control window click on 'TiPad' tab and choose the desired objective for imaging. (Here, first 40x objective was used).
- Load the slide with cells onto the stage and center it beneath the lens.
- Now, on the 'A1plus Compact GUI' tab, set up the lasers appropriate for the fluorophores used. Click on the gear symbol to open the dye and spectral settings menu and select the channels needed and set the laser for each channel.
- Then, select the appropriate emissions in the dropdown menu under the first dichroic mirror.
- Next, under 'A1plus Compact GUI' window, click on 'Ch. Series' to set up the line channel series, which sets up whether the lasers used will fire on the sample simultaneously or sequentially. (Here, sequential passes were chosen, starting with channel 1, followed by channel 2, then 4).
- After that, start scanning by clicking the 'Arrow tip' icon on the top. At this point while the imaging is live, under 'A1plus Compact GUI' window, click on the sliding scale and modify the pinhole size to assure limiting out of focus light. (Here, the lowest available setting (0.5), was used).
- Next, adjust the 'high voltage' and 'offset' settings under each laser to appropriate levels, by using the sliding scales to enable detection of the specific staining while limiting any potential background staining. If a positive-staining sample is available, start by imaging this sample for each channel to make sure the laser settings yield optimal signal to noise ratios.
Caution: high laser intensity for extended periods can cause photobleaching. - After setting the optimal HV and offset values for each laser, click on the 'ND Acquisition tab'and then select the 'Z' icon to set up the parameters for the z-series. Next, while acquiring a live image of the sample, first set the bottom by finding the bottom of the image and clicking the 'bottom' button then find the top position of the sample and click the 'top' button. Set the step size either by specifically typing the preferred step size in µm for each step or by specifying how many total steps are needed.
- Once the z-series parameters have been set, select the desired size/pixel resolution of the image. To do so, click the 'A1plus Compact GUI' window and under the 'size' icon select the desired resolution. To decrease the noise of the image, one can select the dropdown menu next to the 'ø' symbol to average the selected number of images.
- Now, click the 'Run now' tab on the 'ND Acquisition menu' in order to start imaging the sample.
- After imaging is complete, save the image by clicking 'File', then 'Save As', which will export the image file with the extension '.nd2'. Finally, repeat the process for each of the other samples.
Confocal fluorescence microscopy is a specialized imaging technique for localization of a protein or antigen of interest in a cell or tissue sample by labeling the antigen with an antibody-conjugated fluorescent dye and detecting the fluorescent signal. It offers higher spatial resolution than wide-field fluorescence microscopy, with the help of two pinholes placed at the focal planes of the objective lens, giving it the name confocal. It enables users to visualize the staining at a subcellular level, such as the differentiation between surface membrane staining from intracellular staining.
A confocal microscope follows a similar basic principle as a classic fluorescence microscope. The beam from a light source, usually a laser for confocal, is reflected by a dichroic mirror and focused by an objective lens on the sample. This light excites the fluorophores to emit a different wavelength, which travels back through the objective lens and dichroic mirror to a camera or eyepiece.
The enhanced resolution of a confocal microscope is mainly due to the presence of two pinholes, which are very small holes for light to pass through on the excitation and emission light paths. The pinholes are strategically placed at the focal plane of the objective lens. Now, let's switch to a side view schematic of the microscope arrangement to review the light path. After passing through the excitation pinhole, the excitation light beam has the effect of originating from a focal point, which enables the objective lens to then focus the light to a point on the sample as well. The emission beam from this focal point converges at the emission pinhole, which allows it to pass through. Now during excitation, fluorophores within the light path, above and below the focal point, are also slightly excited. While the emission light originating from the focal point passes through the pinhole, the emissions from the out-of-focus points converge before or after the emission pinhole, and are hence blocked, resulting in reduced background fluorescence.
The excitation-emission-detection cycle needs to be repeated for each imaging point in the region of interest, which can be done in a few different ways. For example, laser scanning confocal uses galvanometer scanning mirrors, which deflect the excitation light at different angles. Hence, sweeping the light beam across the specimen in the XY plane. Spinning disc confocal uses a disc with an array of pinholes, which rotates to shift the arrangement of the pinholes. This enables users to illuminate multiple small imaging points in the sample each time, gradually covering the whole area as the disc rotates. As a result of the pinholes, the XY image at the detector represents a narrow Z plane. Therefore, images can be collected from a series of consecutive Z planes, often referred to as a Z stack. From these images, an appropriate software can generate a 3D depiction of the fluorescence signal pattern in the sample.
In this protocol, you will observe immunostaining of mouse fibroblasts, followed by imaging on a confocal microscope to differentially visualize a cell surface protein and a lysosomal protein.
To begin, using sterile techniques, resuspend the cells of interest in 500 microliters of growth media per well, and then seed them into the wells of a four-well chamber slide. Here, we are using mouse fibroblasts that were transfected to express the antigen-presenting molecule, CD1d. To allow cells to adhere to the glass, place the chamber slide in a 5% carbon dioxide incubator at 37 degrees Celsius, and incubate overnight. In the morning, aspirate the media from each well, and then wash the cells once with 500 microliters of PBS for a few seconds.
To fix the cells, add 500 microliters of 1% paraformaldehyde solution into each well, and incubate for 15 minutes at room temperature. After the incubation, collect the paraformaldehyde into an appropriate hazardous liquid waste container, and then remove any remnants of the fixative by washing the cells three times with PBS for a few seconds.
To allow antibody penetration into the cells, add 500 microliters of permeabilization buffer to each well, and incubate on the bench for 15 minutes at room temperature. After permeabilization, wash the cells briefly three times with 500 microliters of PBS. Next, add 500 microliters of blocking buffer to each well, and incubate for one hour at four degrees Celsius to prevent nonspecific antibody binding.
Prepare the primary antibodies, anti-CD1d and anti-LAMP-1, at appropriate working concentrations. Then, aspirate the buffer from the wells and cover the cells in each well with 500 microliters of diluted primary antibody solution and then incubate the slide on a flat surface overnight at four degrees Celsius. The next morning, dilute the secondary antibodies, in this case an anti-mouse and anti-rat antibody with distinct fluorescent tags, in blocking buffer to appropriate working concentrations. Next, aspirate the primary antibody solution from the wells and then wash the cells four times with 500 microliters of PBS. Then, add 500 microliters of the diluted secondary antibody solution to each well, and incubate at room temperature for one hour in the dark. After the incubation, aspirate the secondary antibody solution and wash the wells four times with 500 microliters of PBS to remove any unbound secondary antibody.
To mount the samples after the final wash, carefully detach and remove the chambers from the slide. To remove the residual PBS, hold the slide at an angle over a delicate task wipe, and remove the fluid from the edges without touching the cells. Once the excess PBS is removed, add one drop of antifade mounting medium, containing the nuclear stain DAPI, onto each section of cells. Next, take a 20-by-60-millimeter coverslip, and using just fingertips start lowering the coverslip slowly on either edge, taking care to avoid bubble formation over the cells. Wipe off any extra mounting medium on the slides with a delicate task wipe and store the slides in the dark at room temperature for up to a week.
To begin imaging the cells, first click on the NIS software icon on the desktop. Once in the control window, click on the TiPad tab at the top, and choose the desired objective for imaging. Then, load the slide with cells onto the stage, and center it beneath the lens. Next, on the A1plus Compact GUI tab next to the TiPad tab, set up the lasers appropriate for the fluorophores used. Click on the gear symbol to open the dye and spectral settings menu. Once the dye and spectral settings menu is open, select the channels needed and set the laser for each channel. Then, select the appropriate emissions in the drop down menu under the first dichroic mirror. Next, under A1plus Compact GUI window, click on Ch.Series to set up the line channel series, which sets up whether the lasers used will fire on the sample simultaneously or sequentially.
After that, start scanning by clicking the arrow-tip icon on the top. At this point, while the imaging is live, under A1plus Compact GUI window, click on the sliding scale, and modify the pinhole size to assure limiting out-of-focus light. Next, adjust the high voltage and offset settings under each laser to appropriate levels by using the sliding scales to enable detection of the specific staining while limiting any potential background staining. If a positive staining sample is available, start by imaging this sample for each channel to make sure the laser settings yield optimal signal-to-noise ratios. After setting the optimal HV and offset values for each laser, click on the ND Acquisition tab, and then select the Z icon to set up the parameters for the z series.
Next, while acquiring a live image of the sample, first set the bottom by finding the bottom of the image and clicking the bottom button. Then, find the top position of the sample and click the top button. Set the step size either by specifically typing the preferred step size in microns for each step or by specifying how many total steps are needed. To select the desired size/ pixel resolution of the image, click the Aiplus Compact GUI window, and under the size icon, select the desired resolution.
To decrease the noise of the image, you can select the drop-down menu next to the theta symbol to average the selected number of images. After this, click the Run Now tab on the ND Acquisition menu in order to start imaging the sample. After imaging is complete, save the image by clicking file, then save as, which will export the image file with the extension dot-nd2. Finally, repeat the process for each of the other samples.
In this experiment, mouse fibroblasts expressing the surface glycoprotein gene CD1d were fixed, immunostained, and imaged on a confocal microscope. This image shows a single section of a Z stack at 40X magnification, where CD1d is stained in red. The sample was costained with LAMP-1, a lysosomal marker, in green. Nuclear stain DAPI was used to show the nuclei of the cells.
In a composite image where the three different channels are merged, the appearance of yellow results from overlap of the red and green channels, and indicates an area where CD1d and LAMP-1 are co-localized in the lysosomes. Areas where only one color is present indicate the presence of CD1d or LAMP-1 without co-localization. This image shows a 3D rendering of the cells constructed from images captured in the z-stack and this method enabled the construction of a side view of this group of cells. This following image shows a slice out of the z-stack at 100X magnification, demonstrating the expression patterns of these two proteins in greater detail. The pink outlined box on the right side of the image displays the cross-section of the x-coordinate designated by the pink line in the image, which represents the side view at the pink line. Similarly, the blue outlined box on the bottom of the image shows the cross-section of the y-coordinate designated by the blue line in the image, which represents the front view at the blue line. The 3D rendering of the z-stack image enables users to view the image in 3D, visualizing all of the x, y, and z planes. This can be used to study co-localization of the different stains at different regions within the cell.
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Q1: How does confocal microscopy achieve better resolution than wide-field fluorescence microscopy?
Confocal microscopy uses two strategically placed pinholes at the focal planes of the objective lens to block out-of-focus light. While wide-field microscopy detects fluorescence from above and below the focal plane, creating hazy background, confocal pinholes allow only light from the focal point to reach the detector. This eliminates background fluorescence and dramatically improves both spatial resolution and image contrast.
Q2: What role do pinholes play in confocal microscope imaging?
Pinholes are small holes positioned on both the excitation and emission light paths at the focal plane of the objective lens. The excitation pinhole makes the light beam appear to originate from a focal point, enabling precise focusing. The emission pinhole blocks divergent light from out-of-focus regions, allowing only in-focus fluorescence to pass through to the detector, resulting in reduced background and improved image clarity.
Q3: How can confocal microscopy distinguish between surface and intracellular protein localization?
Confocal microscopy achieves subcellular resolution by collecting images from narrow Z planes using pinholes that eliminate out-of-focus light. This enables visualization at the subcellular level, allowing researchers to differentiate between surface membrane staining and intracellular staining. The technique can reveal whether proteins are located on the cell surface or within specific organelles like lysosomes.
Q4: What is a Z-stack and how is it used in confocal microscopy?
A Z-stack is a series of consecutive images collected from adjacent focal planes throughout the depth of a sample. Because confocal pinholes restrict detection to narrow Z planes, images can be acquired sequentially from top to bottom of the specimen. Specialized software then processes these Z-stack images to generate a three-dimensional depiction of the fluorescence signal pattern, enabling visualization of protein localization in all spatial dimensions.
Q5: Why is sample fixation preferred over live cell imaging for confocal microscopy?
Fixed cells are preferred for confocal Z-stack imaging because acquiring multiple images from numerous Z planes is time-consuming and challenging with live cells. Live cells are difficult to maintain healthy during extended imaging, and their organelles constantly move, compromising image quality and 3D reconstruction accuracy. Fixed cells preserve cellular morphology and remain stationary, allowing reliable collection of high-quality image series.
Q6: How do secondary antibodies with distinct fluorescent tags enable multi-protein visualization?
When imaging multiple cellular targets, primary antibodies from different species are used to label each antigen. Fluorophore-conjugated secondary antibodies, each with unique excitation and emission spectra, then bind to their respective primary antibodies. This allows the confocal microscope to distinguish signals from different proteins using separate laser channels, enabling simultaneous visualization and co-localization analysis of multiple antigens within the same cell.
Q7: What does co-localization indicate when visualizing multiple stained proteins in confocal images?
Co-localization occurs when two or more proteins occupy the same cellular region, appearing as color overlap in merged confocal images. For example, when red and green fluorescent signals overlap, they produce yellow, indicating that both proteins are present in the same subcellular compartment. This technique reveals functional relationships between proteins, such as whether CD1d traffics through lysosomal compartments marked by LAMP-1.