Specimen Preparation Using Synthetic Fluorophores and Immunofluorescence
Staining Adherent Cells
with Intermediate Filament Primary Antibodies and Synthetic Fluorophores
A majority of the common fibroblast and epithelial cell lines derived from humans and laboratory animals produce brightly colored fluorescent specimens highlighting specific proteins in the intermediate filament network (such as vimentin and desmin) when stained with a cocktail that includes anti-intermediate filament antibodies conjugated to common low molecular weight synthetic fluorescent probes. Among the most useful fluorescent markers for visualization of intermediate filaments (as well as actin) are rhodamine, fluorescein, the Alexa Fluor series, and the cyanine dyes. Counterstaining for nuclei using a variety of popular DNA-binding dyes follows treatment with the antibodies and filamentous actin probes. This protocol details a generalized procedure for staining adherent cells.
Presented in Figure 1 is a widefield fluorescence image revealing the vimentin intermediate filament and filamentous actin networks present in adherent mink uterus endometrium epithelial cells (GMMe line). The cells were immunofluorescently labeled with mouse anti-vimentin primary antibodies followed by goat anti-mouse secondary antibodies conjugated to Texas Red-X. In addition, the specimen was counterstained with phalloidin conjugated to Alexa Fluor 488 and DAPI, targeting the actin network and nuclei, respectively. Images were recorded in grayscale with a QImaging Retiga Fast-EXi camera system coupled to an Olympus BX-51 microscope equipped with bandpass emission fluorescence filter optical blocks provided by Omega Optical. During the processing stage, individual image channels were pseudocolored with RGB values corresponding to each of the fluorophore emission spectral profiles.
A wide variety of primary antibodies targeting other antigens can be employed in combination with anti-intermediate filament antibodies. In addition, synthetic fluorophores, such as the MitoTrackers, can also be used as counterstains. Because the mixed aldehyde and detergent fixative described below can obscure some antigen markers, primary antibodies to other targets should first be tested alone before mixing into cocktails with intermediate filament antibodies.
- Cytoskeletal Buffer (CB) - Suspend 18.14 grams of the biological buffer PIPES (free base; 60 millimolar) in one liter of double-distilled water. Add 6.50 grams of HEPES buffer (27 millimolar), 3.80 grams of EGTA (10 millimolar), and 0.99 grams of magnesium sulfate (heptahydrate; 4 millimolar). Stir and adjust the pH to 7.0 with concentrated sodium hydroxide. The PIPES buffer crystals will not completely dissolve until the buffer pH nears neutrality, but should then form a clear solution.
- Phosphate Buffered Saline with Calcium and Magnesium (PBS) - Dissolve 0.2 grams of potassium chloride, 0.2 grams of monobasic potassium phosphate, 8.0 grams of sodium chloride, and 1.74 grams of dibasic sodium phosphate (heptahydrate) in one liter of double-distilled water. After dissolving these reagents, add 0.132 grams of calcium chloride dihydrate and 0.10 grams of magnesium chloride hexahydrate. Adjust the pH to 7.2 with concentrated sodium hydroxide. Addition of the divalent alkaline earth metals to the buffer solution is helpful to ensure that uncondensed chromatin remains intact and contained within the nucleus during the staining procedure, dramatically decreasing background fluorescence levels when using DNA probes excited by ultraviolet irradiation (DAPI and Hoechst). This buffer should be used with cryosections.
- Mixed Aldehyde and Detergent Fixative - Dissolve 3 grams of electron microscope grade paraformaldehyde powder in 100 milliliters of CB with heating, and then filter when the solution appears clear. After cooling and filtering, add 1.5 milliliters of 20-percent Triton X-100 (made in CB) and 100 microliters of 50-percent glutaraldehyde. The final concentration of reagents should be 3 percent paraformaldehyde, 0.3 percent Triton X-100, and 0.05 percent glutaraldehyde. Mix the resulting solution well before use. This reagent should be made fresh daily.
- Blocking Buffer - 10 percent normal goat serum (NGS) in PBS containing 0.05 percent Triton X-100 (add 2-3 milligrams sodium azide per 100 milliliters of blocking buffer to eliminate the growth of microorganisms). If secondary antibodies to a host other than goat are being used, prepare the Blocking Buffer with normal serum from that species.
- PBS-Triton Wash Buffer - For wash sequences immediately before blocking and once again before staining with nuclear dyes, use PBS containing 0.05 percent Triton X-100.
- PBS-Triton Wash Buffer with Blocking Serum - For wash sequences between the primary and secondary antibody incubations and immediately after the secondary antibody treatment, use PBS containing 0.05 percent Triton X-100 and 1 percent normal host (goat) serum.
- Primary Antibody Cocktail - Add the appropriate volume of concentrated primary antibody (for example, anti-vimentin) stock solution to Blocking Buffer diluted 50-percent with PBS-Triton Wash Buffer (to yield a final concentration 5-percent normal goat serum). Several primary antibodies from different hosts can be mixed into a cocktail. Adherent cells on coverslips should be treated with 100 microliters of primary antibody cocktail.
- Secondary Antibody/Phalloidin Cocktail - Add the appropriate volume of secondary antibody conjugated to the selected fluorophore (for example, 8 microliters of goat anti-mouse heavy and light chain secondary IgG with Alexa Fluor 350 at 2 milligrams per milliliter) to 1 milliliter of Blocking Buffer diluted 50-percent with PBS-Triton Wash Buffer (to yield a final concentration 5-percent normal goat or other host serum). If the cells are to be simultaneously counterstained with phalloidin, add 20-40 microliters of concentrated stock solution (6.6 micromolar) to 1 milliliter of the Blocking Buffer. As with the primary antibody mixture, adherent cells on coverslips should be treated with 100 microliters of primary antibody cocktail.
- Nuclear Dye - Prepare fresh dilutions of the nuclear dye immediately prior to staining. When using SYTOX, DRAQ5, and the cyanine nuclear stains, background fluorescence can be significantly reduced (especially in the cytoplasm) by adding 10 milligrams of RNase to the Blocking Buffer. In this case, block the cells or tissues at 37 degrees Celsius, rather than at room temperature, in order to activate the enzyme.
- Nuclear Dye Wash Buffer - Hoechst and SYTOX dyes require Hanks Balanced Salt Solution (Hanks BSS), while DAPI, as well as the monomeric and dimeric cyanine nuclear stains, can be used with PBS.
Nuclear Counterstain Dilutions
- Hoechst (33342 and 33258) - Dilute 5 microliters of 10 milligram/milliliter stock solution in 150 milliliters of Hanks BSS (treat for 30 minutes).
- SYTOX Green and Orange - Dilute 10 microliters of concentrated stock solution (5 millimolar in dimethyl sulfoxide) in 250 milliliters of Hanks BSS (treat for 30 minutes).
- DAPI - Dilute 5 microliters of 10 milligram/milliliter stock solution in 150 milliliters of PBS diluted 50-percent with double-distilled water (treat for 5 minutes).
- Monomeric and Dimeric Cyanine Dyes - Dilute the concentrated stock solution (for example, TO-PRO-3; usually 1 millimolar) as recommended by the manufacturer (1:20 to 1:1000) into PBS (treat for 5 to 30 minutes).
- DRAQ5 - Dilute the concentrated stock solution (usually 1 millimolar) as recommended by the manufacturer (1:20 to 1:1000) into PBS (treat for 5 to 30 minutes).
Aspirate the growth medium from a Petri dish containing healthy cells adhered to coverslips and replace with pre-warmed (37 degrees Celsius) CB buffer to remove medium and serum proteins (use 3 milliliters of buffer for 60-millimeter Petri dishes). Wash the cells twice for 5 minutes (each wash) with the pre-warmed CB buffer, and incubate the cells at 37 degrees Celsius during the wash cycles. Each Petri dish should be individually marked with the primary antibodies and other stains used for the coverslips in that dish. Coverslips should remain with the original Petri dish for each step in the entire procedure.
Fix the cells by adding the appropriate volume of pre-warmed (37 degrees Celsius) mixed aldehyde fixer to each Petri dish and rapidly transfer the dishes to the incubator. Fix for 10 minutes.
Remove the Petri dishes from the incubator and wash once with CB buffer for 5 minutes and twice with PBS-Triton Wash Buffer (5 minutes each wash) before blocking.
Remove the PBS-Triton Wash Buffer and block nonspecific secondary antibody binding sites with 10-percent normal host serum Blocking Buffer. Treat the adherent cells for 60 minutes at room temperature with the Blocking Buffer and slowly rotate the Petri dishes as the cells are being blocked on an orbital shaker at 5-10 revolutions per minute.
During the blocking step, prepare antibody treatment supports by covering 2 × 3-inch microscope slides with Parafilm, as illustrated in Figure 2. Secure the Parafilm so that it adheres tightly and is smoothly distributed along the glass surface (no blisters). After blocking, carefully remove the coverslips from the Petri dishes and place them cell-side down on a 100 microliter drop of diluted primary antibody cocktail in 50-percent blocking buffer deposited on a Parafilm-covered slide. Between 3 and 6 coverslips (depending on size) can be placed on a single slide. Next, place the slides in a humidity chamber (see Figure 3) and incubate the coverslips in the humidity chamber for 1.5 hours at 37 degrees Celsius. If the primary antibodies are not conjugated to fluorophores, it is not necessary to protect the coverslips from light at this point.
After primary antibody treatment, return the coverslips to the original Petri dishes and wash three times at room temperature (5 to 10 minutes for each wash) with PBS-Triton Wash Buffer with Blocking Serum to remove unbound primary antibodies. Slowly rotate the Petri dishes as the cells are being washed on an orbital shaker at 5-10 revolutions per minute.
After washing, carefully remove the coverslips from the Petri dishes and place them cell-side down on a 100 microliter drop of diluted secondary antibody cocktail in 50-percent blocking buffer deposited on a Parafilm-covered slide. Once again, place the slides in a humidity chamber (see Figure 3) and incubate the coverslips in the humidity chamber for 1 hour at 37 degrees Celsius if smaller secondary antibody fragments are being used, or 1.5 hours for entire antibody molecules. It is important to cover the humidity chamber with aluminum foil during this step to protect the fluorophores from light.
After secondary antibody treatment, return the coverslips to the original Petri dishes and wash three times at room temperature (5 to 10 minutes for each wash) with PBS-Triton Wash Buffer with Blocking Serum to remove unbound secondary antibodies. Slowly rotate the Petri dishes as the cells are being washed on an orbital shaker at 5-10 revolutions per minute.
In preparation for nuclear staining, wash the cells twice with PBS-Triton Wash Buffer for 5 minutes (each wash). Slowly rotate the Petri dishes as the cells are being washed on an orbital shaker at 5-10 revolutions per minute.
For DAPI and cyanine nuclear counterstains, add the diluted dye in PBS (dilute PBS to 50 percent with double-distilled water for DAPI) to the Petri dish and treat the coverslips for the recommended time: 5-10 minutes for DAPI; 15-30 minutes for cyanine dyes (protect from light with aluminum foil). When using Hoechst or SYTOX stains (30 minute incubation), first wash the slides in Hanks Balanced Salt Solution for two buffer exchanges prior to counterstaining.
Wash the counterstained coverslips with either PBS or Hanks Balanced Salt Solution (depending upon the nuclear dye) for three times at 5 minutes for each wash. Protect from light with aluminum foil.
In order to remove excess salt, wash the cells three times for 2 to 3 minutes (each wash) in distilled water. Note that this step is only necessary if the coverslips are to be air-dried overnight before mounting.
After the final distilled water washing step, carefully remove the coverslips from the Petri dish with tweezers and wipe excess water from the back and edges. Lean the coverslips on their sides against the labeled Petri dish cover and allow them to dry overnight. Protect the drying coverslips from light with an aluminum baking tray. After drying, mount the coverslips (cell-side down) on clean microscope slides using the appropriate mounting medium.