Analysis of SAMHD1 Restriction by Flow Cytometry in Human Myeloid U937 Cells

This protocol does not contain any studies involving animals or human participants performed by any of the authors. All the steps of this protocol were performed following the guidelines and codes of practice of the institutions where they were carried out.

1. Transfection of 293T cells for VLP production (both SAMHD1-expression vectors and tester HIV particles)

NOTE: The most significant contributors to the successful production of virus particles are the health of the producer 293T cells, confluency at transfection and time of harvest. Laboratories will typically have their preferred transfection reagents and protocol for retroviral particle production. The following protocol yields good quality infectious particles, but equivalent protocols would also be suitable for this application. To maintain good quality 293T cells, passage at least three times per week to maintain a constant growth rate. Cells should be seeded in the log phase of growth for efficient transfection.

1. Day 1: Seed the required number of 10 cm dishes with a 1/4 split from a near-confluent dish of 293T cells to yield around 60% confluency the following day (approximately 5 x 10⁵ cells/mL). Several densities may need to be seeded to achieve an appropriate density for transfection the following day when working with a new cell stock.
2. Day 2 (morning): Check for approximately 60% confluency by microscopy and gently replace media on cells with 10 mL of fresh Dulbecco's Modified Eagle Medium (DMEM).
3. Day 2 (afternoon, at least 4 h after media change): Transfect for VLP production.
   1. Make up the DNA dilution mix containing 3 µg each of gag-pol expressor plasmid, Long-terminal repeat (LTR)-driven reporter plasmid and Vesicular Stomatitis Virus G protein (VSV-G) expressor plasmid (total 9 µg, see NOTE) in 600 µL of serum-free DMEM. Vortex and centrifuge briefly (15 s, >500 x g).
   2. Add 20 µL of transfection reagent (see Table of Materials), vortex (do not centrifuge) and incubate at 20 °C for 15-20 min. Add the transfection mix dropwise to the plate, swirl gently to mix, and return to the incubator.
      NOTE: For MLV VLP expressing SAMHD1 use KB4^5,6 (gag-pol expressor plasmid), pLGatewaySAMHD1IeYFP¹ (LTR-driven reporter plasmid) and pczVSV-G⁷. For HIV-RFP VLP, use p8.91^8,9 (gag-pol expressor plasmid), SCRPSY¹⁰ (LTR-driven reporter plasmid), and pczVSV-G. Alternatives are discussed in the Table of Materials. Plasmid ratios may need to be optimized for different plasmid/transfection systems.
4. Day 3 (late morning): Remove the media carefully from cells by pipetting, wash with 5 mL of fresh DMEM, and replace with 5 mL of fresh DMEM.
5. Day 4 (morning): Harvest the virus by collecting the supernatant from the cells and replace with 5 mL of fresh DMEM. Pass the VLP-containing supernatant through a 0.45 nm filter, aliquot and transfer to -70 °C for storage. Ensure that the aliquots are of a size suitable for planned experiments (250 µL as a suggestion) as repeated freeze-thaw cycles will result in a loss of viral titer.
6. Day 5 (morning): Harvest the virus as in 1.5 but discard the plates after collecting the supernatant.

2. Titering new VLP prior to use

1. Day 1: Seed 293T at 2 x 10⁵ cells per well of a 24-well plate – 4 wells per virus to be titrated, including a virus of known infectivity for each backbone. Optimize the seeding density for a given cell stock.
2. Day 2: Observe the plate to check confluency (ideally ~70%). Thaw the virus-containing-supernatant from steps 1.5 and 1.6. Add 0, 10, 25, or 100 µL to each well.
3. Day 4 (Morning): Harvest the cells for analysis by flow cytometry.
   1. Remove the media by careful pipetting or aspiration. Wash the cells gently with 250 µL of phosphate buffered saline (PBS). Remove the cells from the plate by pipetting up and down with 250 µL PBS, and transfer to a microcentrifuge tube. Add 250 µL of 4% paraformaldehyde (taking final concentration to 2%).
      CAUTION: Paraformaldehyde is toxic. It should not be mixed with chlorine-based disinfectants.
   2. Pellet the cells by centrifugation at 500 x g for 5 min, discard the supernatant and resuspend the cell pellet in 200 µL of PBS or alternative flow cytometry buffer. Analyze for RFP or YFP as appropriate by flow cytometry. Normalize the infectivity of a given virus stock against values for a stock of known infectivity.
      ​NOTE: VLP can also be normalized by p24 enzyme-linked immunosorbent assay (ELISA); however, this is not necessarily a good indicator of VLP infectivity, especially where VLP stocks are prepared in different transfections. Published median tissue culture infectious dose or multiplicity of infection (TCID50/MOI) methods¹¹ can also provide relative quantitation, although these are not well established for MLV. Relative fluorescence provides a cost-effective alternative to commercial reverse transcriptase ELISA methods.

3. Transduction of U937 with VLP expressing SAMHD1 and YFP

NOTE: Steps 3-6 constitute the restriction experiment. Days are numbered for ease of planning.

1. Day 1 (afternoon): Thaw VLP for transduction including MLV-Wild-type SAMHD1-YFP (positive control), MLV-SAMHD1(HD206-7AA)-YFP (negative control) and variant MLV-SAMHD1-YFP (experimental samples). Dilute the normalized VLP to a final volume of 500 µL with Roswell Park Memorial Institute 1640 media (RPMI) in a microfuge tube, add 0.5 µL of polybrene (10 mg/mL), and mix by flicking.
   1. If VLP cannot be titrated beforehand, then use 100 µL in the first instance. Use a maximum 1:1 ratio of VLP to RPMI to limit VLP-induced toxicity. Aim for approximately 30% transduction.
2. Aliquot 5 x 10⁵ U937 cells into a sterile microfuge tube for each transduction condition. Pellet by centrifugation at 500 x g for 5 min, and remove the supernatant (include two additional tubes as untransduced controls for flow cytometry). Resuspend the cell pellet in 500 µL of diluted VLP (or RPMI for untransduced controls) and transfer to one well of a 24-well plate.
3. Spinoculate in a tabletop centrifuge at 800 x g for 90 min at 20 °C. Add 1 mL of 37 °C RPMI to the well and allow to recover in a 37 °C incubator for 3 days.

4. Differentiation of transduced U937

1. Day 4 (morning): Re-suspend the cells by gently pipetting and transfer 350 µL into each of 4 wells of a 12-well plate. Add 700 µL of 37 °C RPMI containing 150 nM PMA to each well giving a final concentration of 100 nM and allow to differentiate for 3 days.
   ​NOTE: PMA is bought as a powder and should be resuspended to 1 mM in DMSO and stored in the dark. A working stock of 100 µM should be prepared by diluting 1/10 from the 1 mM stock with DMSO.

5. Infection of differentiated, SAMHD1-expressing U937 with HIV-RFP

1. Day 7: Observe the cells by microscopy. Ensure that the cells are adherent. Aspirate the media from all the wells, add 1 mL of RPMI to well 1 of each set of 4 allowing for untransduced, uninfected control as well as single color controls. Add back 0.5 mL of RPMI containing roughly 10 µL of HIV-1-RFP (approximately 10 ng p24 as a guide) to all the other wells. Aim for approximately 50% infection.
2. Day 8 (morning): Add 0.5 mL of RPMI to each of the infected wells.

6. Flow cytometry analysis

NOTE: It is good practice to include a live-dead stain in flow cytometry pipelines. However, if the U937 cells are of good quality, this step can be omitted without negative consequences for downstream analysis. Gentle pipetting of trypsinized cells should easily result in a single cell suspension.

1. Day 10 (morning): Aspirate the media from the wells and wash once with PBS. Add 300 µL of cell dissociation enzyme (or trypsin) for 5-10 min. Add an additional 300 µL of PBS, resuspend thoroughly making sure all the cells have come off the plate, and transfer to flow cytometry tubes.
2. Add 300 µL of 4% paraformaldehyde (taking final concentration to 2%). Pellet the cells by centrifugation at 500 x g for 5 min, discard the supernatant and resuspend the cell pellet in 200 µL of PBS or alternative flow cytometry buffer. Analyze by flow cytometry.
   ​NOTE: The steps below are for a Fortessa analyzer, but equivalent analysis can be achieved using any analyzer capable of reading RFP and YFP fluorescence. Consult local flow cytometry support or other experienced researcher before setting up any flow cytometry analysis. When many samples need to be screened at the same time, a high throughput sampler may be appropriate. In this case, a larger cell number and volume are advised.

7. Flow cytometry analysis

1. Turn on the cytometer 10 min before it is required and turn on the computer. Check that the waste is empty and that the sheath tank is full.
2. Log in to the machine and open the analysis software.
3. Move the arm to the side, take the water tube off, set the flow rate to High and press Prime. Wait for the light to go out and repeat three more times.
4. Put the water back on and run on high for 3 min and then switch to Low and press Standby until proceeding to acquisition.
5. Meanwhile set up the experiment within the software:
   1. Select Experiment/ New Experiment (name according to local guidance). In the inspector, delete the unrequired fluorochromes, leaving Blue Laser 530/30 and Yellow Laser 610/20 or the recommended laser and filter combinations for YFP and RFP for the machine.
   2. Right click on the experiment and choose Cytometer Settings/Application Settings/Create Worksheet. On the worksheet, create a Forward Scatter-Area (FSC-A)/Side Scatter Area (FSC-A) dot blot and YFP histogram, RFP histogram, and GFP/YFP dot blot by clicking on the corresponding icon and changing the axes by clicking on the axis label.
   3. Right click again on Experiment and add New Specimen. Rename as appropriate. Open the specimen by clicking the plus sign to reveal a new tube.
6. Open the acquisition dashboard from View, load the uninfected, untransduced control and press Acquire. Adjust FSC and SSC voltages in the dashboard to position cells in the lower-left quadrant (no cells on the axes). Gate around this population P1. Right click on other plots and select show P1 to remove debris from subsequent analysis.
7. Load the single-color control for YFP (wild-type SAMHD1 only) and acquire. Adjust the YFP voltage in the dashboard so that the most fluorescent cells are within the range of the detector (not on the edge of the axis). Repeat for single color RFP control.
8. Run automatic compensation using the untransduced, uninfected and single-color controls.
   1. Select Experiment > Compensation Setup > Create Compensation Controls from the menu to toggle to a normal worksheet. Select unstained normal worksheet, press on Run and acquire for the unstained control.
   2. Draw a gate around the intact cells (P1) and record. Remove the tube and put the machine on Standby. Right click on P1, click on Apply to All Compensation Controls.
   3. Switch to the RFP normal worksheet, and press Run. Record the single-color RFP control, and put the machine on Standby. The software should automatically select the positive population.
   4. Repeat for YFP single color control. Select Experiment > Compensation Setup > Calculate Compensation > Link and Save.
9. Switch to the global worksheet. Acquire the cells transduced with wild type SAMHD1, infected with HIV-RFP and check that four distinct populations can be seen in the corners of the quadrants that are aligned vertically and horizontally for YFP and RFP, respectively. Remove the tube and press Standby.
10. Reload the untransduced, uninfected sample, press Run and Record 30,000 events with P1 as the stopping gate. Repeat for the remaining samples (including other controls). Rename the samples while running or post hoc. Once exported, files cannot be renamed.
11. Export the data as .fcs files and clean the fluidics according to local instructions.

8. Data analysis

NOTE: The steps that follow correspond to analysis in FlowJo v10; however, equivalent steps can easily be performed in other analysis software.

1. Open the software and drag all .fcs files into the dashboard. Select the compensation controls and drag them into the compensation subfolder. Open the file for the untransduced, uninfected tube by double-clicking, which will bring up the FSC-A/SSC-A plot.
2. Select the polygon tool and gate on the intact cell population, excluding debris in the lower-left corner. Name this Cells. Drag this gate onto the whole population of tubes in the All Samples bar. Scroll through each individual sample using the horizontal arrow buttons to ensure gating is appropriate for each tube.
3. Double-click on the Cells population for the untransduced, uninfected sample and adjust the axes to FSC-Height (H) vs FSC-Area (A). Select the single large population using a rectangular gate to exclude doublets. The software will automatically suggest naming this Single Cells.
4. Drag this gate onto all the Cells populations and again, check that the gating is appropriate for all samples. Select the Single Cells population for the uninfected, untransduced sample, change the axes to compensated blue laser (y, YFP) versus compensated yellow laser (x, RFP). Press T on the axis and select Biexponential scale for x and y.
5. Select the Quadrant Gating Tool and click at the upper-right extreme of the negative cell population. Apply this preliminary gating to all Single Cell populations by dragging into the All Samples bar. Where the quadrant gates do not separate the populations satisfactorily, it may be necessary to gate individual quadrants using the polygon tool, as in Figure 2.
6. Scroll to the wild-type SAMHD1 only sample (YFP only) and check that the gating between the untranduced (YFP negative) and YFP positive is correct. It may help to switch to contour view to discriminate negative from dim YFP cells. If the upper-most YFP +ve cells are spread widely, adjust the upper-vertical gate to the right.
7. Scroll through all samples and check the gating with respect to YFP positivity. Use the best division between YFP negative and positive that is valid for all samples.
   NOTE: The compensation will have been calculated based on the wild-type SAMHD1 YFP expression level. If infection levels are very different between different SAMHD1-YFP transduced cells, the compensation may need to be adjusted. It is therefore preferred that YFP VLP are normalized prior to use.
8. Scroll to the untransduced, HIV-RFP infected tube and check whether the gating between uninfected RFP negative and infected RFP positive is correct. Adjust as appropriate, using the contour view if required and scroll through the remaining samples to check the gating is valid for all samples.
9. Each sample requires double negatives (Q4: lower left, untransduced, uninfected), YFP positive (Q1: upper left, SAMHD1 expressing, uninfected), RFP-positive (Q3: lower right, untransduced, HIV infected), and double positive (Q2: upper right, SAMHD1 expressing HIV-infected). Open the table editor by clicking on the icon and drag the four quadrant gates into the dashboard. Select Excel Export and click on Create Table. Save the generated spreadsheet according to local file naming conventions.
10. To export representations of plots and gating strategies, click on the Layout Editor icon. Select each population and drag into the editor with the axes, labeling and spacing required.
11. To create a layout with the same plots shown for all samples, select one column and press on Batch. Press on Scale to Width and then Avoid Page Breaks. Save in the file format required.
12. In the exported spreadsheet, calculate the restriction ratio by generating columns of percentage RFP of YFP-ves (RFP+ve / (double negatives + RFP+ve)) and percentage RFP of YFP+ves (double positives / (double positives + YFP+ve)) and then a final column of (%RFP of YFP+ve / %RFP of YFP-ve), see Figure 1. Average the replicate data for each SAMHD1 construct and calculate whether restriction values for tested SAMHD1 variants are significantly different from wild-type and/or 206-7AA using appropriate data analysis software.