Determination of the Relative Cell Surface and Total Expression of Recombinant Ion Channels Using Flow Cytometry

Summary

Inherited cardiac arrhythmias are often caused by mutations that alter the surface delivery of one or more ion channels. Here, we adapt flow cytometry assays to provide a quantification of the relative total and cell surface protein expression of recombinant ion channels expressed in tsA-201 cells.

Abstract

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these multi-spanning transmembrane proteins, with or without change in channel gating, is often postulated to contribute significantly in this process. It has thus become critical to develop a method to quantify the change of the relative cell surface expression of cardiac ion channels on a large scale. Herein, a detailed protocol is provided to determine the relative total and cell surface expression of cardiac L-type calcium channels Ca_V1.2 and membrane-associated subunits in tsA-201 cells using two-color fluorescent cytometry assays. Compared with other microscopy-based or immunoblotting-based qualitative methods, flow cytometry experiments are fast, reproducible, and large-volume assays that deliver quantifiable end-points on large samples of live cells (ranging from 10⁴ to 10⁶ cells) with similar cellular characteristics in a single flow. Constructs were designed to constitutively express mCherry at the intracellular C-terminus (thus allowing a rapid assessment of the total protein expression) and express an extracellular-facing hemagglutinin (HA) epitope to estimate the cell surface expression of membrane proteins using an anti-HA fluorescence conjugated antibody. To avoid false negative, experiments were also conducted in permeabilized cells to confirm the accessibility and proper expression of the HA epitope. The detailed procedure provides: (1) design of tagged DNA (deoxyribonucleic acid) constructs, (2) lipid-mediated transfection of constructs in tsA-201 cells, (3) culture, harvest, and staining of non-permeabilized and permeabilized cells, and (4) acquisition and analysis of fluorescent signals. Additionally, the basic principles of flow cytometry are explained and the experimental design, including the choice of fluorophores, titration of the HA antibody and control experiments, is thoroughly discussed. This specific approach offers objective relative quantification of the total and cell surface expression of ion channels that can be extended to study ion pumps and plasma membrane transporters.

Introduction

This paper provides a reliable assay to report the relative cell surface expression of membrane proteins such as ion channels expressed in recombinant cells using the existing flow cytometry technology. Ion channels are pore-forming membrane proteins that are responsible for controlling electrical signals by gating the flow of ions across the cell membrane. They are classified by the activation mechanism, nature, and selectivity of ion species transiting through the pore where they are localized. At the cellular and tissue levels, the macroscopic ion fluxes through ion channels are the product of biophysical (gating and permeation), biochemical (phosphorylation), and biogenesis (synthesis, glycosylation, trafficking, and degradation) properties¹. Each of these processes is unique to every type of ion channels and is optimized to fulfill the physiological role of the ion channel. Consequently, alterations in any of these fine-tuned processes through an inherited or a genetic modification, often referred to as "channelopathy", can be detrimental to cell homeostasis. It is important to stress that delivering the "right" amount of ion channels at the cell surface is critical to cell homeostasis. Even small increases (gain-of-function) and small decreases (loss-of-function) in ion channel activity have the potential to cause a serious pathology over a lifetime. Defects in the cell surface delivery of mature ion channels is an important determinant in numerous channelopathies, such as cystic fibrosis (CFTR ion channel)² and cardiac arrhythmias of the long QT syndrome form (cardiac potassium channels)³.

Channelopathies are associated with cardiac sudden death⁴. The current worldwide prevalence of all cardiac channelopathies is thought to be at least 1:2,000-1:3,000 per individual⁵ and are responsible for about half of sudden arrhythmic cardiac death cases⁶. Dysfunction in cardiac voltage-gated sodium-, potassium-, and calcium- selective ion channels are known to play a key role in this process. The L-type Ca_V1.2 voltage-gated calcium channel is required to initiate synchronized heart muscle contraction. The cardiac L-type Ca_V1.2 channel is a multi-subunit protein complex composed of the main pore-forming Ca_Vα1 subunit and Ca_Vß and Ca_Vα2δ1 auxiliary subunits^7-12. Note that the full complement of auxiliary subunits is required to produce functional Ca_V1.2 channels at the plasma membrane and dynamic interactions between these subunits are essential to support the normal electric function of the heart¹³. Ca_Vß promotes the cell surface expression of Ca_V1.2 channels through a non-covalent nanomolar hydrophobic interaction¹⁴. Co-expression of the Ca_Vα2δ1 subunit with Ca_Vß-bound Ca_Vα1 stimulates peak current expression (5 to 10-fold) and promotes channel activation at more negative voltages. Gain-of-function mutations of the pore-forming subunit Ca_V1.2 have been associated with a form of ventricular arrhythmias called the long QT syndrome¹⁵ whereas a host of point mutations in the three main subunits forming the L-type Ca_V1.2 channel have been identified in subjects suffering from arrhythmias of the short QT syndrome form^16,17. Ion channels are membrane proteins that can be investigated from a biochemical perspective (protein chemistry) or using electrophysiological tools (current-generating machines) and often using these complementary approaches. Electrophysiology, in particular whole-cell patch-clamping, is a suitable approach to elucidate the function of ion channels¹⁵ but cannot resolve modifications in protein trafficking from changes in their biophysical properties. Protein chemistry has, however, often limited use due to the relatively low expression of large membrane proteins relative to smaller soluble proteins. Robust high-throughput methods using fluorescence readout need to be developed in order to specifically address defects in protein biogenesis causing changes in the cell surface expression of ion channels.

Flow cytometry is a biophysical technology employed in cell counting, sorting, biomarker detection, and protein engineering¹⁸. When a sample solution of live cells or particles is injected into a flow cytometer, the cells are ordered into a single stream that can be probed by the machine's detection system (Figure 1). The first flow cytometer instrument produced in 1956¹⁹ detected only one parameter but modern flow cytometers have multiple lasers and fluorescence detectors that allow the detection of more than 30 fluorescent parameters^20,21. Filters and mirrors (emission optics) direct the light scatter or fluorescent light of cells to an electronic network (photodiode and detectors) that convert the light proportionally to its intensity. Digital data are analyzed using specialized software and the primary output is displayed as a dot plot²¹.

Figure 1: Biophysical principles of flow cytometry sorting. Single cells are pushed through a nozzle under high pressure within a stream of sheath fluid which moves them across one or more laser interrogation points. The light beam is deflected by the passing cells and the light collected in the forward direction (Forward Scatter, FCS) is sent to a photodiode that converts the light into a signal proportional to the size of the cell. The light is also collected at a 90° angle to the laser path and sent to detectors (also called photomultipliers (PMT)). This light is routed through dichroic mirrors that permit the detection of the side scatter signal (SSC), which reflects the granularity within the cells, and the fluorescent emissions if excited fluorochromes are present in the cell. Three detectors (Green, Yellow, and Red) are represented with different wavelength bandpass filters, allowing the simultaneous detection of different fluorochromes. The different signals are digitized by an external computer and converted into data that will be analyzed to quantify the characteristics of the cells. Please click here to view a larger version of this figure.

The high-throughput capacity of flow cytometers was exploited to quantify the relative membrane expression of recombinant wild-type and trafficking-deficient voltage-gated L-type Ca_V1.2 channels and associated subunits in live cells. cDNA constructs coding for the proteins were doubly tagged to simultaneously carry an extracellular non-fluorescent epitope that can be detected by an impermeable fluorescent conjugated antibody and an intracellular fluorophore that is constitutively fluorescent. Both the extracellular epitope, inserted in an extracellular loop of the protein, and the intracellular fluorophore, inserted after the C-terminus, are translated with the protein. In this series of experiments, the Ca_Vα2δ1 protein was engineered to express an extracellular hemagglutinin (HA) epitope (YPYDVPDYA) detected by an impermeable FITC (Fluorescein isothiocyanate)-conjugated anti-HA and mCherry as the intrinsic intracellular fluorophore. To determine the relative cell surface expression level of the mCherry-Ca_Vα2δ1 HA-tagged protein, recombinant cells expressing the fusion protein were harvested after transfection, and stained with the FITC-conjugated mouse monoclonal anti-HA epitope tag antibody (Figure 2). FITC is an organic fluorescent compound that is considerably smaller than enzyme reporters and therefore not as likely to interfere with biological function. mCherry- Ca_Vα2δ1-HA overexpressed in tsA-201cells, produces a significant 3-log increase in the FITC fluorescence and mCherry fluorescence on two-dimensional plots²². Given that the HA epitope is located in the extracellular portion of the protein, the fluorescence intensity for FITC obtained in the presence of intact cells reflect the relative index of the cell surface expression of HA-tagged protein. The accessibility of the HA epitope in the constructs is systematically validated by measuring the FITC signal after cell permeabilization. This measure also serves to corroborate the normalized total protein expression since the relative fluorescence intensities for FITC estimated in permeabilized cells are qualitatively comparable to the relative fluorescence values for mCherry measured under permeabilized and non-permeabilized conditions^22,23. It is important to note that the intrinsic fluorescence spectrum is shifted toward higher values after permeabilization but that the only value being reported is the change in fluorescence intensity as compared to the control construct. Relative changes in the fluorescence intensity for the test constructs are estimated using the ΔMean Fluorescence Intensity (ΔMFI) values for each fluorophore (mCherry or FITC). Experiments are designed to measure the fluorescence intensity of the test construct relative to the fluorescence intensity of the control construct expressed under the same conditions to limit experimental variations in the intrinsic fluorescence of the fluorophore-conjugated antibody. Two membrane proteins were successfully studied using this assay: the pore-forming subunit of the L-type voltage-gated calcium channel Ca_V1.2^14,22 and in a different series of experiments, the extracellular auxiliary Ca_Vα2δ1 subunit^22,23. The following protocol was used to determine the cell surface expression of the Ca_Vα2δ1 subunit of the cardiac L-type Ca_V1.2 channel under control conditions and after mutations affecting the posttranslational modification of the ion channel. Under standardized experimental conditions, the cell surface fluorescence of FITC increases quasi-linearly with the expression of cDNA coding for the mCherry-Ca_Vα2δ1-HA proteins (Figure 5 from reference²²).

Figure 2: Schematic representation of total and membrane labeling in the flow cytometry experimental protocol. The scheme outlines some of the main steps necessary to quantify the relative total and cell surface expression of recombinant ion channels by flow cytometry. Cells are transfected with the double-tagged construction mCherry-Ca_Vα2δ1-HA in tsA-201 cells (1) and stained before or after permeabilization (2). Multiparameter data are acquired in a flow cytometer (3) for multivariate analysis (4). Please click here to view a larger version of this figure.

Protocol

1. Doubly Tagged DNA constructs

1. Insert the HA epitope (YPYDVPDYA) in the extracellular linker of Ca_Vα2δ1 between D676 and R677 by site directed mutagenesis (Figure 3B)²⁰. Use Forward primer gccggattatgcgGGAAAACTCCAAACAACC and Reverse primer acatcatacggataTCAATAAATTCATTGAAATTTAAAAGAAATTC.
2. Subclone the cDNA sequence of the tagged HA Ca_Vα2δ1 into the mammalian pmCherry-N1 expression vector designed to express the protein fused to the N-terminus of mCherry between the SacI and SalI sites (Figure 3B)²⁰.
   NOTE: Appropriate channel function needs to be tested with the control construct using standard electrophysiological methods²⁴.

2. Liposome-mediated Transient Transfection (30 min, All Steps are Performed Under Laminar Flow Hood)

1. Day 1: Plate half a million of tsA-201 cells (or HEKT) in 35 mm culture dishes with 2 mL of Dulbecco's high-glucose minimum essential medium (DMEM-HG) supplemented with 10% Fetal Bovine Serum (FBS) and 1% penicillin-streptomycin (PS) culture medium. Count cells using a standard hemacytometer. Assess cell viability from a fraction of the cell sample using Trypan Blue. Plate enough cells to reach 90% confluence at the time of transfection.
2. Day 2: Change culture medium with 2 ml of fresh pre-warmed (37 °C) culture medium without PS.
3. For each transfection sample, prepare two 1.5 ml tubes. In tube 1, dilute 4 µg of DNA in 250 µl of reduced serum medium. In tube 2, mix 10 µl of the liposome-mediated transfection reagent with 250 µl serum-reduced culture medium. Mix gently the transfection reagent before use.
4. Incubate for 5 min at room temperature.
5. Combine the contents of tube 1 and tube 2, mix gently and incubate at least 20 min at room temperature.
6. Add the liposomes/DNA complexes to the cultured cells and gently rock the culture dish to mix.
7. Incubate at 37 °C under 5% CO₂ atmosphere for 24 hr.

3. Staining of Cells for Flow Cytometry (3 hr)

1. Preparing Cell Samples
   1. Day 3: Remove medium from the culture dish carefully and wash the cells with 400 µl of pre-warmed (37 °C) 0.05% trypsin-1x EDTA (ethylenediaminetetraacetic acid).
   2. Add 400 µl of trypsin-EDTA and incubate the dish at 37 °C under 5% CO₂ atmosphere for 5 min to allow cells to detach from the dish.
   3. Stop the enzyme digestion by adding 1 ml of cold culture medium without PS and wash out all the cells from the surface by pipetting gently 4-5 times. Avoid over-digestion and over-pipetting to reduce cell death.
   4. Collect cells in 1.5 ml tubes and place immediately on ice. Use ice cold solutions and keep the cells at 4 °C to prevent the internalization of surface antigens. Decrease lighting to limit photobleaching of the fluorescent signal.
   5. Centrifuge tubes at 400 x g for 5 min at 4 °C. Carefully aspirate and discard the supernatant.
   6. Re-suspend the pellet in 1 ml of phosphate-buffered saline 1x (PBS) to prepare a single cell suspension.
   7. Briefly vortex the tubes very gently and repeat steps 3.1.5 and 3.1.6 to completely remove culture medium.
   8. Re-suspend the pellet in 600 µl of 1x PBS and adjust the cell concentration to a minimum of 3 x 10⁶ cells/ml.
   9. Divide the cells in two new 1.5 ml tubes for extracellular and intracellular staining. Include appropriate controls to discriminate specific staining from non-specific staining.
      NOTE: The isotype control antibody helps assessing the level of background staining and should ideally match each primary antibody's host species, isotype and fluorophore. Use isotype control and conjugated antibody at the same protein concentration.

Table 1: Flow-cytometry experiment control samples for non-permeabilized and permeabilized cells. Each experiment needs to include the following negative controls: (1) Nontransfected cells (without antibody, with the isotype or with the conjugated antibody). (2) Transfected cells with the protein of interest subcloned in a plasmid without constitutive fluorescent intracellular fluorochrome (pCMV- Ca_Vα2δ1-HA) or with the doubly tagged construction (pmCherry-Ca_Vα2δ1 and incubated without antibody, with the isotype or with the conjugated antibody). Single color controls are used for compensation of fluorochrome emission overlap. The same controls are run for non-permeabilized and permeabilized conditions in each series of experiments.

2. Cell Surface Staining of Intact Live Cells
   1. Aliquot 1 x 10⁶ cells/100 µl in 1.5 ml tubes.
   2. Add the FITC-conjugated monoclonal anti-HA antibody at 5 µg/ml and, vortex before incubating the cells on a rocker platform (200 rpm) in the dark at 4 °C for 45 min.
      NOTE: The optimal concentration of antibody was determined in preliminary titration experiments (Figure 4).
   3. Remove the cells from the dark and add 900 µl of 1x PBS/tube. Centrifuge at 400 x g for 5 min at 4 °C.
   4. Aspirate the supernatant and resuspend the pellet in 1 ml of 1x PBS, vortex and centrifuge at 400 x g for 5 min at 4 °C.
   5. Repeat the wash (step 3.2.4) twice to remove any unbound antibody. If an unconjugated primary antibody is used, incubate with the appropriate secondary antibody.
   6. After the final wash, resuspend the cells in 500 µl of 1x PBS and transfer the single cell suspension in 5 ml flow cytometry tubes. Keep the cells in the dark at 4 °C until running the sample.
   7. Run the samples on a flow cytometer. For best results, analyze the cells on the flow cytometer as soon as possible and no later than 24 hr after.
3. Intracellular Staining: Fixation, Permeabilization, and Staining
   1. Aliquot 1 x 10⁶ cells/100 µl in 1.5 ml tubes and centrifuge at 400 x g for 5 min at 4 °C.
   2. Discard supernatant and resuspend cells in 100 µl of fixation-permeabilization solution directly from stock.
   3. Incubate in the dark at 4 °C for 20 min.
   4. Add 100 µl of freshly prepared 1x permeabilization-washing buffer (dilute 10x permeabilization-washing buffer in distilled H₂O). Vortex and sediment cells using a table centrifuge at 400 x g for 5 min at 4 °C.
   5. Aspirate and discard the supernatant.
   6. Repeat steps 3.3.4 and 3.3.5.
   7. Add FITC-conjugated monoclonal anti-HA antibody at 5 µg/ml in 100 µl of 1x permeabilization-washing buffer and, vortex before incubating cells in the dark at 4 °C for 30 min.
      NOTE : The intracellular staining is performed following the same procedure as the one used for cell surface staining. Saponin-mediated cell permeabilization is however, a quickly reversible process, therefore it is important to replace 1x PBS with 1x Perm/Wash buffer to keep the cells in the constant presence of saponin during intracellular staining.
   8. Remove the cells from the dark and add 100 µl permeabilization-washing buffer. Centrifuge at 400 x g for 5 min at 4 °C.
   9. Aspirate carefully the supernatant and resuspend the pellet in 100 µl permeabilization-washing buffer, vortex and centrifuge at 400 x g for 5 min at 4 °C.
   10. Repeat the wash (step 3.3.9) one more time to remove any unbound antibody.
   11. After the final wash, resuspend the cells in 500 µl of 1x PBS and transfer the single cell suspension to 5 mL flow cytometry tubes. Keep the cells in the dark at 4 °C until injecting the sample into the flow cytometer.
   12. Run the samples on a flow cytometer. Run the fixed samples on the cytometer as soon as possible but no later than 1 week after staining. Run the non-permeabilized and permeabilized cells on the same day.

4. Flow Cytometry

1. Flow Cytometer Cell Sorter Daily Setup
   1. Turn on the flow cytometry software. Prior to experiment, calibrate and setup the flow cytometer cell sorter to ensure optimal instrument performance (i.e. laser and optics are performing to specification, the laser and flow cell are properly aligned) by using instrument setup beads.
   2. Use the 100 µm nozzle with 20 psi sheath pressure.
      NOTE: The nozzle does not have to be changed on a bench flow cytometer.
   3. Set the cytometer's flow rate according to the manufacturer specification. Exceedingly high flow rates will decrease sensitivity in the detection of variations in fluorescence.
   4. Select blue (488 nm to excite Fluorescein Isothiocayanate or FITC) and yellow-green (561 nm to excite mCherry) lasers. Collect FITC and mCherry fluorescence levels with a 530/30 nm and with a 610/20 nm bandpass filter respectively.
   5. Acquire the forward scatter (FCS) versus side scatter (SSC) dot plot for unstained cells using linear scale. Adjust each detector's amplification to visualize cells in the lower left quadrant of the dot plot.
2. Sample Reading of Intact Non-permeabilized Cells
   1. Set the P1gate for live non-permeabilized cells by delineating a free form around the cells to be analyzed excluding cell debris and cell aggregates, thus limiting the fluorescence signal to intact cells.
      NOTE: Live/dead exclusion dyes can be used to facilitate gate placement on live cells. Set 10,000 events to record in the stopping gate P1. Set this to a higher number of events if need be.
   2. Acquire mCherry versus FITC two-parameter contour plot to detect baseline autofluorescence of unstained cells. Use bi-logarithmic scale to show negative values and improve resolution between populations²⁵. Adjust each detector's voltage to set the unstained negative cells within the lower portion of the first ten units of the log fluorescence intensity plots.
   3. Acquire all intact non-permeabilized samples using settings established in 4.1.5 and 4.1.6 and collect FSC, SSC and signals in the fluorescence detectors.
   4. Export and save *.fcs files to be used for analysis using flow cytometry analysis software.
3. Sample Reading of Permeabilized Cells
   1. Move the P1 gate to select live cells in the permeabilized samples and adjust FSC and SSC voltage as shown in 4.1.5 and 4.1.6.
   2. Acquire all permeabilized samples and collect FSC, SSC and signals in the fluorescence detectors.
   3. Export and save *.fcs files to be used for analysis using flow cytometry analysis software.
4. Data Analysis
   1. Launch the flow cytometry analysis software and import *.fcs files saved in 4.2.4 and 4.3.3.
   2. Click on the first sample listed in the workspace window. A new window named after the tube I.D. number opens automatically. Start the gating process in the plot of SSC versus FSC. Draw a gate (P1) using the Ellipse icon around live cells and eliminate any debris, dead cells, or aggregates which have different forward scatter and side scatter than live cells
   3. To draw the two-parameter contour plot of the mCherry (y-axis) versus FITC (x-axis) fluorescence intensity of the live cells, click first on the x-axis and choose the FITC-A channel and then click on the y-axis and choose the PE-mCherry-A channel. Click on the "Quad" icon to position the quadrant marker at the edge of autofluorescent cells in each fluorescence channel.
      NOTE: The gate set around the FITC and mCherry positive cells is the P2 gate. The fluorescence negative cell population is referred to as the P3 gate. See Figure 5 for the representative gating method used in this article.
   4. Select P2 and P3 gates and click on the "Add Statistics" icon in the original workspace window. Click on "Count" (number of positive cells) and click on "Mean" (Mean Fluorescence Intensity of each fluorochrome) or "Median" (Median Fluorescence Intensity of each fluorochrome) statistics among the list of options. Click on the "Add Statistics" icon again. All these values are automatically transferred to the original workspace window.
      NOTE: The "Mean" is used only if the fluorescence intensity follows a normal distribution. In every other case, click on the "Median" tab. MFI thus could refer to Mean Fluorescence Intensity or Median Fluorescence Intensity.
      NOTE: The next step is to apply the gates' parameters and statistics to all samples probed by the cytometer.
   5. In the workspace window, use the mouse to drag and drop the gates and statistics parameters onto the line marked ALL SAMPLES.
   6. Generate a batch report of two-dimensional contour plots (mCherry vs FITC) and histograms (cell count versus fluorescence intensity) for non-permeabilized and permeabilized cells (Figures 6A-B).
   7. From the statistics generated in step 4.4.4, calculate the mean fluorescence intensity (MFI) for each fluorochrome for the stained cells. From this value, subtract the MFI value obtained from unstained cells to quantify the surface and total expression of the protein of interest.
   8. Report the ΔMFI values for each fluorophore (mCherry and FITC) (Figure 6C-D). Normalize the ΔMFI measured for the Ca_Vα2δ1 construct mutants to the ΔMFI value obtained for FITC and mCherry with the WT construct.
      NOTE: The absolute value of fluorescence intensity could vary sharply depending upon the batch of antibodies and the technical abilities of each lab worker, hence the need to normalize fluorescence intensity of the mutant construct using the WT construct.

Results

This article describes a reliable protocol to quantify total and cell surface of recombinant ion channels expressed in tsA-201cells by a two-color flow cytometry assay. As an example, the relative cell surface and total protein expression was quantified for the Ca_Vα2δ1subunit. In order to perform the two-color flow cytometry assay, Ca_Vα2δ1 was doubly tagged to express an extracellular non-fluorescent epitope HA that can be detected by an anti-HA FITC-conjugated antib...

Discussion

This flow cytometry-based assay was successfully applied to the measurement of relative total and cell surface levels of fluorescently-labelled pore-forming and associated subunits of voltage-gated calcium channels^14,22,26. It is best used when investigating the impact of genetic mutations and thus requires that the intrinsic fluorescence intensity of the fluorescently-labelled tagged wild-type construct be at least 10 to 100-fold larger than for the fluorescence intensity of the fluorescently-labelled untagge...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Q5 Site-Directed Mutagenesis Kit	New England Biolabs	E0554S	Can be substitute with QuickChange site-directed mutagenesis Kit (Agilent, #200523).
Tubes 1.5 ml	Sarstedt	72-690-001
Tubes 15 ml	Sarstedt	62-554-002
Disposable graduated Tranfer Pipets	VWR	160001-192
100 mm culture dish	Corning	430167	For standard culture of HEKT cells.
35 mm culture dish	Falcon	353001	For standard culture of HEKT cells.
Serological pipette 1 ml	Sarstedt	86.1251.001
Serological pipette 5 ml	Sarstedt	86.1253.001
Serological pipette 10 ml	Sarstedt	86.1254.001
Serological pipette 25 ml	Sarstedt	86.1285.001
Dulbecco's high-glucose medium	Life Technologies	12100-046	Warm in 37 °C water bath before use.
Fetal Bovine Serum, qualified, heat inactivated, US origin	Life Technologies	16140-071
Penicillin-Streptomycin (10,000 U/ml)	Life Technologies	15140-122
Lipofectamine 2000	Life Technologies	11668-019	For liposomal transfection. Can be substituted with calcium phosphate transfection.
Opti-MEM I Reduced Serum Medium	Life Technologies	31985-070	Warm in 37 °C water bath before use.
Trypsin-EDTA (1x) 0.05%, phenol red	Life Technologies	25300-062
1.5 ml microtubes	Sarstedt	72.690.001
Phosphate Buffered Saline 1x	Fisher	BP661-10	Can be "home-made".
Anti-HA FITC conjugated antibody	Sigma	H7411
IgG1−FITC Isotype Control antibody	Sigma	F6397
BD Cytofix/Cytoperm Fixation/Permeabilization Solution Kit	BD Biosciences	554714	Fixation/Permeabilization. Permeabilization/Wash solution, store at 4 °C.
Hemacytometer	Fisher	49105161
Trypan Blue	Fisher	15250061	To access cell viability.
Refrigerated Microcentrifuge, 5430R	Eppendorf	A14H172200
Forma Steri-Cycle CO2 Incubator	Fisher	370
Laboratory Platform Rocker	Fisher	545034
Water Bath	VWR	89032-216
BD FACSARIA III	BD Biosciences	648282	Flow cytometer.
FlowJo Software v10	FlowJo	FlowJo v10 Dongle	For data analysis.