This article details a protocol for visualizing stress granules in osteosarcoma cells using fluorescence microscopy. The method involves cell fixation, permeabilization, and antibody staining to identify specific proteins associated with stress granules.
Take a multi-well plate containing adherent osteosarcoma cells on coverslips.
These bone cancer cells are pre-treated with a stress-inducing chemical, resulting in stress granule formation.
Stress granules are cytoplasmic membrane-less organelles comprising non-translating mRNAs, small ribosomal subunits, RNA-binding proteins, or RBPs, translation-related factors, and signaling proteins.
Discard the media and wash the cells with buffer. Fix the cells to preserve cellular morphology.
Permeabilize the cells to access intracellular targets. Add blocking proteins to prevent non-specific antibody binding.
Introduce primary antibodies that bind to specific RBPs and translation-related factors within stress granules.
Incubate with different fluorophore-labeled secondary antibodies and Hoechst dye.
Secondary antibodies bind to primary antibodies bound to the stress granule-associated proteins, while Hoechst molecules stain the DNA.
Mount the cell-containing coverslips using mounting media.
Using a fluorescence microscope, observe the cells to visualize and quantify stress granules, which appear as fluorescent foci characterized by the co-localization of associated proteins.
To fix the cells, remove the media from the wells, and wash the cells with approximately 250 microliters of PBS. Add approximately 250 microliters of a buffered 4% paraformaldehyde solution to fix the cells, making sure the top of the coverslip is completely covered. Then, leave the plate on a bench rocker at room temperature for 15 minutes. Next, remove and properly discard the paraformaldehyde.
Add approximately 250 microliters of ice-cold methanol to permeabilize and flatten the cells. Incubate the plate for an additional 5 minutes at room temperature on a rocker. Once the permeabilization is complete, discard the methanol and block the cells by applying approximately 250 microliters of a blocking buffer for 1 hour at room temperature. Then, prepare the primary antibody solution by adding 12 microliters of the antibodies against G3BP1, eIF4G, and eIF3b to 3 milliliters of blocking buffer.
Add 250 microliters of the antibody solution to each well of the 24-well plate. Incubate the plate on a rocker for at least 1 hour at room temperature. After 1 hour of incubation, remove the antibody solution and wash the cells with approximately 250 microliters of PBS for 5 minutes. Next, add 12 microliters of anti-mouse Cy2, anti-rabbit Cy3, and anti-goat Cy5 antibodies together with 3 microliters of Hoechst dye to 3 milliliters of blocking buffer to prepare the secondary antibody solution.
Add 250 microliters of the secondary antibody solution to each well, and cover the plate to protect the samples from light. Incubate the plate on a rocker at room temperature for 1 hour. When the incubation is completed, remove the antibody solution, and wash the cells with PBS for 5 minutes. Heat mounting medium in the 37 degrees Celsius heat block for 10 minutes to reduce viscosity. Then, using a pre-cut 200-microliter pipette tip, place 25 microliters of the mounting medium onto a labeled glass slide.
With fine forceps, transfer the coverslip from the well to the slide, inverting the top so that the cells face the mounting medium. Use a clean P200 tip to press the coverslip down. Once all the coverslips are mounted, remove the excess mounting medium by pressing a lab tissue firmly against the slide. Afterwards, flush the slide with water, and immediately blot it with lab tissue once again.
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