An Immunofluorescence Assay to Characterize and Quantify Mammalian Stress Granules

1. Cell Preparation

1. Add autoclaved coverslips to 12 wells of a 24-well plate, as indicated in Figure 1A; this can be done using a sterile Pasteur pipette attached to a vacuum to pick up each coverslip. Gently tap the coverslip on the side of the well to release the suction, allowing the coverslip to fall into the well.
2. Plate 1 x 10⁵ U-2 OS (U2OS) osteosarcoma cells per well at a final volume of 500 µL of medium. Move the plate side to side and up and down several times to ensure that the cells are evenly distributed.
3. Gently press the coverslips down with a clean P1000 tip to ensure that the coverslip is on the bottom of the well and that there are no bubbles under the coverslip.

2. Stressing Cells

1. The following day, visually check the plate to ensure that the cells are evenly dispersed and evenly confluent across the coverslip, as variations between samples may adversely impact the reproducibility of results. Use a 10X objective on an inverted tissue-culture microscope.
2. Preheat the medium to 37 °C. Avoid applying cold or hot media to the cells, as these cause cold shock or heat shock, respectively.
3. Dilute drugs in preheated media. For each different drug concentration, prepare 2 wells containing 500 µL of medium. Prepare an additional 500 µL to allow for loss while pipetting. Aliquot 4 tubes, each containing 1.5 mL.
   1. For sodium arsenite: to each well containing 500 µL, add 1.5 µL of 100 mM sodium arsenite (final concentration: 100 µM) or add 0.75 µL of 100 mM sodium arsenite (final concentration: 50 µM).
   2. For vinorelbine: to each well containing 500 µL, add 22.5 µL of 10 mM vinorelbine (final concentration: 150 µM) or add 18.75 µL of 10 mM vinorelbine (final concentration: 100 µM).
      NOTE: Sodium arsenite is a well-characterized and commonly used stress granule inducer and is therefore classically used as a positive control.
4. Remove and discard the media from wells B1 – 4 and C1 – 4. Add the media with drugs and wait for 60 minutes (Figure 1A).
   1. Add 500 µL of medium with 100 µM sodium arsenite to wells B1 and B2. Add 500 µL of medium with 50 µM sodium arsenite to wells B3 and B4.
   2. Add 500 µL of medium with 150 µM vinorelbine to wells C1 and C2. Add 500 µL of medium with 125 µM vinorelbine to wells C3 and C4.
5. 30 min prior to fixation, treat the wells in column 2 with 5 µL of cycloheximide (1 mg/mL stock) and the wells in column 4 with 2 µL of puromycin (1.25 mg/mL stock). Gently rock the plate to mix before placing the plate back in the 37 °C incubator.

3. Cell Fixation and Immunofluorescence

NOTE: Before the experiment, ensure that the methanol is cooled to -20 °C. Make buffers, including Phosphate-buffered Saline (PBS), 5% Normal Horse Serum (NHS) in PBS, and 4% paraformaldehyde (PFA) in PBS.

1. Discard the media and wash the wells containing coverslips with PBS. Depending on the drugs used, it might be important to properly discard the media-containing drugs; contact the local environmental safety office for specific instructions. To wash efficiently, fill a squeeze bottle with PBS, cut back the tip in order to allow a gentle flow rather than a tight stream, and use this to add PBS to each well after aspirating the original PBS.
2. Fix the cells using ~250 µL of 4% PFA for 15 min at room temperature, RT under gentle agitation. Completely cover the top of the coverslip with PFA; 250 µL should be sufficient, but if not, add more. Always avoid pipetting directly onto the coverslips, as some cells (or stress treatments of cells) can alter attachment, and the force from pipetting can dislodge the cells.
3. Remove the PFA and discard properly. Contact the local environmental safety office for details on how to properly discard paraformaldehyde.
4. Add ~250 µL of methanol (-20 °C) and incubate for 5 min at RT under gentle rocking; this both permeabilizes and flattens the cells.
5. Remove and discard the methanol and block the cells by applying ~250 µL of 5% NHS for 1 h at RT overnight, O/N, at 4 °C. Contact the local environmental safety office for details on how to properly discard methanol.
6. For primary antibodies, dilute the antibodies in 5% NHS.
   NOTE: The use of multiple SG markers is key to confirming whether the granules observed are bona fide SGs. The number of SG markers that can be used depends on the filters available in the microscope. If standard green, red, and far-red filter sets are available, use G3BP1, eIF4G, and eIF3b at a 1/250 dilution.
   1. For this experiment (12 coverslips at 250 µL each-Total: 3 mL of 5% NHS), add 12 µL of each of the following: anti-G3BP1, anti-eIF3b, and anti-eIF4G. Incubate the primary antibodies using gentle agitation for at least 1 h at RT or O/N at 4 °C.
7. Wash the wells 3x with PBS, and incubate each wash for 5 min.
8. For secondary antibodies, dilute all secondary antibodies (1/250 dilution) and Hoechst dye (1/1,000 dilution) in 5% NHS.
   1. For this experiment (12 coverslips at 250 µL each-Total: 3 mL of 5% NHS), add 12 µL of each of the following: Cy2-Mouse, Cy3-Rabbit, and Cy5-Goat (a 1/250 dilution of each) and add 3 µL of Hoechst dye.
   2. Incubate the coverslips with the secondary antibodies for 1 h at RT. During this and the following steps, protect the samples from light by covering the plate with a box or by placing foil over the top of the tissue culture dish.
9. Wash the wells three times with PBS for 5 min each.
10. Mount the coverslips onto labeled glass slides using a mounting medium. Heat the mounting medium in a 37 °C heat block for 10 min; this decreases the viscosity and makes it easier to pipette. With a cut P200 tip, pipette 25 µL of mounting medium per coverslip onto a glass slide; 4-8 coverslips can fit on one glass slide, assuming that the microscope stage allows this. Using fine forceps, transfer the coverslip from the well to the slide, making sure to put the cell side down on the mounting medium. Using a clean P200 tip, press the coverslip down.
11. Once all the coverslips have been mounted, use folded lab tissue to clean up the excess mounting medium by pressing the lab tissue very firmly onto the slide. Use a squeeze bottle containing H₂O to flush off the excess mounting medium and then repeat the blotting with folded lab tissue to remove the water.