In total, 24 patients with HNSCC, which consisted of 15 male and 9 female patients, were evaluated in the current study. The age range of the patients was 21–94 years, with a median age of 64 years. Histological analysis revealed that 95.83% of the patients possessed cancers of a squamous origin. In total, 16 patients were treated using a single treatment modality, including surgery, radiation or chemotherapy, and 8 patients were treated using a combination of various treatment modalities, including surgery with radiation or chemotherapy (Table I). Peripheral blood (10–15 ml) was collected from the 24 patients and also from 24 healthy control donors. Peripheral blood mononuclear cells (PBMCs) were isolated using Ficoll gradient centrifugation, as previously described (15,16). The present study was approved by the Research Ethics Board at the University of Manitoba (Winnipeg, MB, Canada).

Sequential staining was performed to evaluate CD3ζ expression in T cells using flow cytometry. Surface CD3ε, a marker of T cells, was stained using specific fluorescein isothiocyanate (FITC) mouse anti-human CD3 monoclonal antibody (clone, UCHT1; cat no. 55916; dilution, 1:66; BD Biosciences, Franklin Lakes, NJ, USA). Following surface staining, PBMCs were fixed in 1% paraformaldehyde (Sigma-Aldrich, Ontario, Canada)for 10 min at room temperature, followed by two rounds of washing with phosphate-buffered saline (PBS). In total, 2 ml of cell permeabilization buffer (0.1% saponin and 0.05% sodium azide in PBS; Sigma-Aldrich) was added to the PBMCs. The PBMCs were kept on ice for 15 min in the dark, washed once with cell permeabilization buffer and resuspended in 0.2 ml of the cell permeabilization buffer. The expression of CD3ζ was detected intracellularly, using a specific phycoerythrin (PE)-conjugated monoclonal antibody (clone, 2H2D9; ca no. IM3169; dilution, 1:10; Beckman Coulter, Inc., Brea, CA, USA). An immunoglobulin (Ig)-G1 isotype control (eBiosciences, San Diego, CA, USA) was used to distinguish specific CD3ζ binding from non-specific binding during flow cytometry. The optimal amount of antibody was determined by titration and the cells were analyzed by BD FACSCanto™II flow cytometer (BD Biosciences). A total of 10,000 events were acquired and the expression levels of CD3ζ in T cells (gated on CD3ε⁺) were analyzed using FlowJo software, version 9.6.2 (FlowJo, LLC, Ashland, OR, USA) by the mean florescence intensity (MFI).

PBMCs were isolated from the whole blood of healthy donors and HNSCC patients using Ficoll gradient centrifugation, as previously described (15,16), and washed once in complete medium, consisting of RPMI-1640, penicillin/streptomycin/L-Glutamine and 10% fetal bovine serum (Thermofisher Scientific, Inc., Ontario, Canada). Viable cell numbers were determined by a B4005 hemocytometer and trypan blue viable cell exclusion dye (VWR International, Ontario, Canada) (17). PBMCs were cryopreserved at −196°C in aliquots in complete medium, which contained a final concentration of 10% dimethyl sulfoxide, using a Mr. Frosty™ Freezing Container (Thermo Fisher Scientific, Inc., Waltham, MA, USA), according to the manufacturer's protocol. The cryopreserved PBMC was removed from liquid nitrogen, and rapidly thawed in a 37°C water bath prior to the cells being transferred to 10 ml pre-warm complete media. The cell pellet was collected and resuspended in 1 ml of fresh, warm complete media at 37°C. The viable cell count was determined, and the thawed cells were used directly for flow cytometric assays.

The present study collected PBMCs from 5 healthy donors. The PBMCs of the healthy donors were pooled and stored in small aliquots in liquid nitrogen until use. In each flow cytometric assay, one of these vials was thawed (0.5×10⁶ cells) and used in the staining process along with the patient samples. To evaluate the intra and inter-assay variation, the same internal control was thawed and the CD3ζ expression was analyzed 3 times using flow cytometry at 3 time points in weekly intervals. The MFI of the CD3ζ chain expression was measured within an individual experiment for intra-assay variability and between multiple experiments for inter-assay variability.

To determine the cell viability and to exclude the dead cells from the analysis, Fixable Viability Dye eFluor® 780 (eBioscience, Inc., San Diego, CA, USA) was used, according to the manufacturer's protocol. The cells were washed with flow buffer (0.05% FBS, 10% sodium azide in PBS) and surface staining was performed using an anti-CD3ε monoclonal antibody. The cells were then kept in the dark at 4°C for an additional 30 minutes prior to intracellular staining for CD3ζ. Live T cells, gated from the dead cells using either the fixable viability dye or forward scatter/side scatter (FSC/SSC), were analyzed for CD3ζ chain expression.

CD3ζ expression (expressed as the MFI) was determined in the PBMCs of the internal control, healthy controls and patients using an individual flow cytometric assay. Data normalization was performed by transforming the MFI values of the test samples (patients and healthy donor) to a common scale using the following equation: Final relative fluorescence intensity = MFI of the test sample/MFI of the internal control.

Normality and statistical analyses of the data were performed using the GraphPad Prism 5 software (GraphPad Software, Inc., La Jolla, CA, USA), termed the normality test. The D'Agostino-Pearson omnibus test was performed to investigate whether the data passed the normality test. An unpaired t-test was applied to the data that passed the normality test. The Mann-Whitney test was used on data that did not pass the normality test.

To evaluate the specificity of anti-CD3 antibodies in flow cytometry, fluorochrome-conjugated isotype controls, consisting of the PE-mouse IgG1/κ antibody for CD3ζ detection and the FITC-IgG1/κ antibody for CD3ε detection, were used in the surface and intracellular staining of PBMC. The present study demonstrated that, when compared to the isotype controls, anti-CD3ζ and anti-CD3ε antibodies were specific (Fig. 1).

Using a commercial fixable viability dye, the present study observed that the percentage of dead cells in the thawed samples were in the range of 2–23% (Fig. 2). To measure CD3ζ chain expression using MFI in live cells, the present study separated live cells from dead cells using the viability dye prior to the analysis of CD3ζ expression in the CD3ε⁺ T cells. In addition, the present study compared the differences in CD3ζ expression when the live cells were gated by either the viability dye or the FSC/SSC properties of the total cells acquired. The present study observed that there were no significant differences in the MFI determined by these two methods of live cell gating (Fig. 2).

Subsequently, the present study calculated the standard deviation for the inter-assay and intra-assay variations in MFI of CD3ζ expression using various aliquots of the identical PBMC sample thawed and assayed at various time points (weekly intervals for 3 consecutive weeks) for the inter-assay, and triplicate measurements of 1 aliquot of PBMC sample thawed and assayed in a single flow cytometric assay for the intra-assay. The ranges of the variations observed within an assay and between multiple assays were 407–412 and 407–477, respectively. The standard deviations for intra-assay and inter-assay were 2.16 and 28.6, respectively (Fig. 3).

Despite efforts to perform flow cytometric assays under identical conditions, the present study observed larger inter-assay variations in the MFI values of CD3ζ expression during the last 36 months. The present study noted that the MFI values of CD3ζ expression of the identical internal control varied between 378 and 1,106. It was reasoned that an internal control provided not only a quick way to examine MFI variations acquired using various equipment or the same equipment under identical photomultiplier (PMT) voltages, but also a reference value for data normalization in the inter-assays (Table II). Therefore, the present study applied a normalization procedure to the data sets of CD3ζ expression in the PBMC samples of the patients with HNSCC, which represented a low range (MFI, 378; Table III) and a high range of CD3ζ MFI (MFI, 1,106; Table IV). In addition, the present study incorporated a data set collected under an extreme MFI (MFI, 3,106; Fig. 4; Table V) to additionally test the normalization procedure. As revealed in Tables III–V, normalization transformed the data in the low, high and extreme MFI groups into more consistent values.

Previous small cross-sectional studies have reported a reduced level of the CD3ζ chain in the peripheral blood lymphocytes of patients compared with healthy controls (12,13). Therefore, the present study compared the MFI of CD3ζ expression in the healthy controls and the HNSCC patients prior to and following the normalization process. The present study measured the MFI of CD3ζ expression in the CD3ε⁺ T lymphocytes of the healthy control individual and patient samples and normalized the data using the internal control under identical conditions in each experiment. Prior to normalization, no significant difference was observed between the MFI of CD3ζ expression in the healthy controls and the patients with HNSCC. By contrast, following normalization, CD3ζ expression was observed to be significantly decreased in the patients with HNSCC compared with the healthy controls (Fig. 5). Subsequently, the present study validated the normalization procedure in the analyses of CD3ζ expression in a longitudinal study. The present study used longitudinal PBMC samples from a single patient to illustrate the feasibility of using the normalization procedure in a longitudinal study. As revealed in Fig. 6, the use of the internal control during normalization standardized the flow cytometric CD3ζ MFI data acquired at various time points, which in turn allowed each subsequent follow-up sample to be compared to the previous sample, including the baseline data obtained prior to the patient receiving any type of clinical treatment (Fig. 6).