Confocal Fluorescence Microscopy: A Technique to Determine the Localization of Proteins in Mouse Fibroblasts

1. Materials

Buffers

1. Wash buffer: 1 X sterile phosphate-buffered saline (PBS) without calcium or magnesium
2. Fixation buffer: 1% paraformaldehyde in PBS
3. Permeabilization buffer: 0.1% Triton X-100 in PBS
4. Blocking buffer: 1% bovine serum albumin in PBS
5. Cell growth medium: DMEM supplemented with 10% fetal bovine serum (FBS), penicillin/streptomycin, and L-glutamine

Equipment

6. Laminar flow hood
7. Humidified incubator (37°C, 5% CO₂)
8. Confocal laser scanning microscope; here, Nikon Eclipse Ti laser

Materials and Reagents

9. Chambered cell culture slides
10. Anti-fade mounting media with DAPI (for staining nuclei)
11. Microscope cover glass
12. Delicate task wipes
13. Pipettors and tips

Assay Specific Reagents

14. Adherent cells (primary cells or cell lines); here, mouse fibroblasts transfected with CD1d were used (LCD1).
15. Primary antibodies to detect cellular targets; here, rat anti-mouse CD107a (LAMP-1) and mouse anti-mouse CD1d were used.
16. 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

1. Resuspend the cells of interest in growth media.
2. 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).
3. Incubate the chamber slide overnight in a 5% CO₂ incubator at 37°C, to allow cells to adhere to the glass.
4. Next day, aspirate the media from each well and then wash the cells 1X with 500 µL PBS.

Fixation

5. To fix the cells, add 500 µL 1% paraformaldehyde solution into each well and incubate for 15 min at room temperature.
6. After the incubation, collect the paraformaldehyde into an appropriate hazardous liquid waste container.
7. Then, wash the cells 3 times with 500 µL PBS to remove any remnants of the fixative.

Permeabilization

8. Permeabilize the cells by incubating with 500 µL permeabilization buffer/well for 15 minutes at room temperature.
9. Then, wash the cells briefly 3 times with 500 µL of PBS.

Blocking

10. 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

11. Aspirate the blocking buffer from the slide chambers.
12. 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)).
13. 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

14. Aspirate the primary antibody solution from the wells.
15. Wash the well chambers 4 times with 500 µL PBS.
16. 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).
17. Incubate at room temperature for 1 h in the dark.
18. After the incubation, aspirate the secondary antibody solution.
19. 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

1. First, carefully remove the chambers from the slide.
2. Then, hold the slide at angle over a delicate task wipe and remove the fluid from the edges without touching the cells.
3. Add 1 drop of antifade mounting medium, containing the nuclear stain DAPI, onto each section of cells.
4. 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).
5. 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.

1. To begin imaging the cells, open the 'NIS Elements Advanced Research software' by clicking on the 'NIS software icon.'
2. Next, on the control window click on 'TiPad' tab and choose the desired objective for imaging. (Here, first 40x objective was used).
3. Load the slide with cells onto the stage and center it beneath the lens.
4. 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.
5. Then, select the appropriate emissions in the dropdown menu under the first dichroic mirror.
6. 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).
7. 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).
8. 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.
9. 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.
10. 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.
11. Now, click the 'Run now' tab on the 'ND Acquisition menu' in order to start imaging the sample.
12. 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.