Flow Cytometry-Based Quantification of Therapy-Induced Senescent Cancer Cells

1. Induction of senescence by chemotherapy drugs in cultured cancer cells

NOTE: All cell manipulation steps in this section should be performed in a biosafety cabinet using sterile practices. This section is written for adherent cell types. Suspension cells may be used with appropriate modifications as noted.

1. Grow cancer cell line(s) according to the standard protocol from the supplier or the laboratory that provided the specific cell line(s) used.
   NOTE: Low passage cells (p < 10) are generally preferred as there will be lower levels of replicative senescence, i.e. lower background, in untreated cell samples.
2. One day prior to senescence induction by drugs, harvest cells with trypsin-EDTA 0.25% (or as recommended). Neutralize trypsin by adding an equal volume of complete culture medium, and transfer the cell suspension to a sterile conical tube.
   NOTE: This step is not needed for suspension cells.
3. Count the cells using the standard hemacytometer method and record cells/mL. Plate cells at 1 × 10³-10 × 10³ cells/cm² in standard 6-well plastic culture dishes.
   NOTE: Optimal plating density is dependent on the proliferation rate of cells and should be user-determined. Cells should be in log phase growth at approximately 10%-20% confluency at the time of treatment (i.e., following 18-24 h incubation after plating). The starting density (cells/mL) of suspension cells should be user-determined. Six-well plates typically yield enough senescent cells per well for one standard flow cytometry analysis sample. If flow-sorting, a much larger surface area (e.g., multiple P150 plates) should be used to enable the recovery of sufficient numbers of senescent cells for downstream assays (≥1 × 10⁶).
4. Incubate plated cells overnight (18-24 h) in a 37 °C incubator with 5% CO₂ and a humidity pan.
5. Treat the plated cells with senescence-inducing chemotherapy drug(s). Include at least one positive control, e.g., etoposide (ETO) or bleomycin (BLM). Prepare duplicate wells per drug. Include one set treated with vehicle-only as control.
   NOTE: A dose curve for each experimental agent should be tested by the user to determine the optimal concentration for senescence induction in the cell line(s) being used.
6. Incubate cells for 4 days in a 37 °C incubator with 5% CO₂ and a humidity pan to allow the onset of senescence. Examine daily for expected morphology changes using a light microscope.
   NOTE: Incubation times from 3-5 days may be acceptable depending on the rate of senescence onset. Media can be changed and the agent reapplied (or not), as desired, to promote healthy growth conditions while achieving an acceptable percentage of senescent cells.
7. After the onset of senescence, harvest the cells by adding trypsin-EDTA 0.25% for 5 min at 37 °C. When cells are dissociated into suspension, neutralize trypsin with an equal volume of complete medium.
   NOTE: This step is not needed for cells growing in suspension. If surface marker staining will be conducted, avoid the use of trypsin-EDTA as it can temporarily destroy surface antigens on cells. Instead, gently dissociate the monolayer using a sterile plastic cell scraper (or an alternate dissociation reagent designed to preserve surface antigens).
8. Count the cells in each sample using a hemacytometer. Calculate the cells/mL for each sample.
   NOTE: Trypan blue can be added to evaluate the percentage of dead cells at this point (i.e., due to drug treatment), but cell death will also be determined with a fluorescent viability dye during DDAOG staining workflow.
9. Aliquot ≥0.5 × 10⁶ cells per sample into 1.7 mL microcentrifuge tubes.
   NOTE: The number of cells per sample should be standardized across all samples.
10. Centrifuge the tubes for 5 min at 1,000 × g in a microcentrifuge at 4 °C. Remove the supernatant.
    NOTE: If a refrigerated microcentrifuge is unavailable, it may be acceptable to perform centrifugations at ambient temperature for certain resilient cell types.
11. Proceed to DDAOG staining in section 2.

2. DDAOG staining of SA-β-Gal in cell samples

1. Dilute 1 mM Bafilomycin A1 stock at 1:1,000x into DMEM medium (without FBS) for a final concentration of 1 µM.
2. Add prepared Baf-DMEM solution to the cell pellet samples (from step 1.11) for a concentration of 1 × 10⁶ cells/mL.
   NOTE: For example, if using 0.5 × 10⁶ cells per sample, add 0.5 mL of Baf-DMEM.
3. Incubate for 30 min at 37 °C (without CO₂) on a rotator/shaker set at a slow speed.
   NOTE: Avoid CO₂ incubators for the staining process, which can acidify solutions and thereby interfere with Baf and DDAOG staining.
4. Without washing, add the DDAOG stock solution (5 mg/mL) at 1:500x (10 µg/mL final) to each sample. Pipette to mix. Replace on a rotator/shaker at 37 °C (without CO₂) for 60 min. Protect from direct light.
5. Centrifuge the tubes for 5 min at 1,000 x g at 4 °C. Remove the supernatant.
6. Wash with 1 mL of ice-cold 0.5% BSA per tube and pipette to mix. Centrifuge the tubes for 5 min at 1,000 x g at 4 °C and remove the supernatant. Repeat this step 2x to thoroughly wash the cells. Remove the supernatant and proceed.
   NOTE: It is important to perform the wash steps in step 2.6 to remove uncleaved DDAOG, which can exhibit undesired fluorescence emission (460/610 nm).
7. (Optional) Fixation and storage of DDAOG stained cells for later analysis
   1. Add 0.5 mL of ice-cold 4% paraformaldehyde dropwise to each washed sample. Pipette to mix.
   2. Incubate for 10 min at room temperature.
   3. Wash the cells 2x with 1 mL of PBS.
   4. Store the samples for up to 1 week at 4 °C prior to flow cytometry analysis.
      NOTE: For fixed samples, skip step 2.8.
8. Dilute Calcein Violet 450 AM stock (1 mM) at 1:1,000x into 1% BSA-PBS (1 µM final). Add 300 µL (for cultured cell samples) to the washed cell pellets from step 2.6. Incubate for 15 min on ice in the dark.
9. Proceed to flow cytometry setup (section 3).

3. Flow cytometer setup and data acquisition

1. Transfer the cell samples to tubes compatible with the flow cytometry instrument. Place the tubes on ice and keep them protected from light.
   NOTE: If aggregates are observed in the cell suspensions, pass the suspension through 70-100 µm cell strainers prior to analysis. Do not use 40 µm strainers because they can exclude some of the larger senescent cells.
2. In the referenced software (see the Table of Materials), open the following plots: 1) FSC-A vs SSC-A dot plot, 2) violet channel histogram, 3) far-red channel (e.g., APC-A) versus green channel (e.g., FITC-A) dot plot.
   NOTE: Doublet exclusion plots and single-channel histograms can also be used but are not strictly required.
3. Initiate cytometer data acquisition.
   1. Place the vehicle-only control sample stained with DDAOG on the intake port. At a low intake speed, begin to acquire sample data.
   2. Adjust FSC and SSC voltages so that >90% of events are contained within the plot. If cells do not fit well on the plot, lower the area scaling setting to 0.33-0.5 units.
   3. Remove the vehicle-only sample without recording data.
   4. (Optional) Add one droplet of rainbow calibration microspheres to a cytometer tube with 1 mL of PBS. Place the tube on the cytometer intake port. Begin to acquire sample data.
   5. Adjust violet, green, and far-red channel voltages so that the top peak of the rainbow microsphere is in the range of 104-105 units of relative fluorescence in each channel and all peaks are well separated in each channel. Record 10,000 events. Remove the tube.
4. Place the positive control sample (e.g., BLM, ETO) stained with DDAOG on the intake port. At a low speed, acquire sample data. Observe the events in FSC, SSC, violet, green, and far-red channels to ensure that over 90% of events are contained within all plots. Look for an increase in AF and DDAOG signal versus vehicle-only control.
5. If using a sorting cytometer, initiate sorting at this step.
   1. For record-keeping purposes, record 10,000 cells for the control sample and each sorted sample.
   2. Sort the desired amount of cells (≥1 × 10⁶ is typically suitable) into an instrument-appropriate collection tube with 3-5 mL of culture medium.
   3. After sorting, proceed to downstream culture or analysis.
      NOTE: For the calibrated flow cytometer used herein, optimal channel voltages typically fell between 250 and 600 (mid-range), but the optimal voltages and channel voltage ranges will vary across instruments. Avoid using voltages at very low or high ranges, which may suppress signal or amplify noise.
6. After completing steps 3.1-3.5 and making adjustments to the cytometer settings as necessary, record data for all the samples. Ensure that the settings remain uniform for all sample recordings. Record ≥10,000 events per cultured cell sample.
   NOTE: Although gating and analysis can be performed using data acquisition software (e.g., FACSDiva), a complete gating and analysis workflow to be conducted post-acquisition using separate analysis software (FlowJo) is described in section 4 below. Post-acquisition analysis is preferred to reduce time at the cytometer workstation and take advantage of additional tools included in the dedicated analysis software.
7. Save sample data in .fcs file format. Export the files to a workstation computer equipped with flow cytometry analysis software (e.g., FlowJo).

4. Flow cytometry data analysis

NOTE: The workflow presented uses FlowJo software. Alternative flow cytometry data analysis software may be used if the key steps described in this section are similarly followed.

1. Using FlowJo software, open .fcs data files from step 3.7.
2. Open the layout window.
3. Drag and drop all samples into the layout window.
4. Gate viable cells.
   1. First double-click on the sample data for the vehicle-only control to open its data window.
   2. Visualize the data as a violet channel histogram. Identify the viable cells stained by CV450 based on their brighter fluorescence than the dead cells.
   3. Draw a gate using the single-gate histogram tool to include viable cells only. Name the gate viable.
   4. Then, from the sample layout window, drag the viable gate onto the other cell samples to apply the gate uniformly.
   5. In the layout window, visualize all samples as violet channel (viability) histograms. Verify that viable cell gating is appropriate across samples before proceeding; if not, make adjustments as needed.
      NOTE: Viability staining can exhibit variations across treatments.
5. Gate senescent cells.
   1. Double-click on the gated viable cell data for the vehicle-only control to open its data window.
   2. Visualize the data as a dot plot for far-red channel (DDAOG) vs green channel (AF).
   3. Draw a gate using the rectangle gating tool to include <5% of cells that are DDAOG+ and AF+ (upper right quadrant). Name the gate senescent.
   4. Then, from the sample layout window, drag the senescent gate onto the viable subsets of the other cell samples to apply the gate uniformly.
   5. Into the layout window, drag and drop all viable cell subsets gated in section 4.4. Visualize all viable samples as far-red (e.g., APC-A) versus green channel (e.g., FITC-A) dot plots.
   6. Ensure that the senescent gate drawn in step 4.5.3 is visible on all plots and that the gate for the vehicle-only control exhibits ≤5%-10% senescent cells.
6. Once the percentage of senescent cells has been determined using the steps above, present the resulting data using the FlowJo plots, summarized in a data table, and/or statistically analyzed using standard software.