CD8^low CD100⁻ T Cells Identify a Novel CD8 T Cell Subset Associated with Viral Control during Human Hantaan Virus Infection

ABSTRACT

Hantaan virus (HTNV) infection can cause a severe lethal hemorrhagic fever with renal syndrome (HFRS) in humans. CD8⁺ T cells play a critical role in combating HTNV infections. However, the contributions of different CD8⁺ T cell subsets to the immune response against viral infection are poorly understood. Here, we identified a novel subset of CD8⁺ T cells characterized by the CD8^low CD100⁻ phenotype in HFRS patients. The CD8^low CD100⁻ subset accounted for a median of 14.3% of the total CD8⁺ T cells in early phase of HFRS, and this percentage subsequently declined in the late phase of infection, whereas this subset was absent in healthy controls. Furthermore, the CD8^low CD100⁻ cells were associated with high activation and expressed high levels of cytolytic effector molecules and exhibited a distinct expression profile of effector CD8⁺ T cells (CCR7^+/− CD45RA⁻ CD127^high CD27^int CD28^low CD62L⁻). When stimulated with specific HTNV nucleocapsid protein-derived peptide pools, most responding CD8⁺ cells (gamma interferon [IFN-γ] positive and/or tumor necrosis factor alpha [TNF-α] positive) were CD8^low CD100⁻ cells. The frequency of CD8^low CD100⁻ cells among HTNV-specific CD8⁺ T cells was higher in milder cases than in more severe cases. Importantly, the proportion of the CD8^low CD100⁻ subset among CD8⁺ T cells in early phase of HFRS was negatively correlated with the HTNV viral load, suggesting that CD8^low CD100⁻ cells may be associated with viral clearance. The contraction of the CD8^low CD100⁻ subset in late phase of infection may be related to the consistently high expression levels of PD-1. These results may provide new insights into our understanding of CD8⁺ T cell-mediated protective immunity as well as immune homeostasis after HTNV infection in humans.

IMPORTANCE CD8⁺ T cells play important roles in the antiviral immune response. We found that the proportion of CD8^low CD100⁻ cells among CD8⁺ T cells from HFRS patients was negatively correlated with the HTNV viral load, and the frequency of CD8^low CD100⁻ cells among virus-specific CD8⁺ T cells was higher in milder HFRS cases than in more severe cases. These results imply a beneficial role for the CD8^low CD100⁻ cell subset in viral control during human HTNV infection.

INTRODUCTION

CD8 T cells play a critical role in combating viral infections. Dramatic cellular changes occur as T cells transition through the following three characteristic phases of an antiviral response: initial activation and expansion, the death phase, and the formation of memory T cells. However, many aspects of these activation and differentiation processes are inadequately understood, particularly in humans. There are several important cell surface molecules that regulate the interactions between immune cells and, moreover, characterize cell differentiation and activation status.

Sema4D, also called CD100, was the first immune semaphorin discovered (1) and is involved in several aspects of both humoral and cellular immunity (2–6). CD100 exists in both a membrane-bound form and a soluble form. Membrane-associated CD100 is expressed preferentially on T cells and weakly on B cells and antigen-presenting cells (APCs) (2, 7). CD72, the receptor for CD100, is expressed primarily on immune system cells and is present on the surface of most APCs and B cells. Interaction between CD72 and CD100 leads to dendritic cell (DC) maturation and cytokine production and enhances B cell activation (7).

Because of the involvement of CD100 in the immune response, we hypothesized that CD100 may play an important role in the antiviral CD8 T cell response. However, there is limited information available regarding the role of CD100 in infectious diseases. Eriksson et al. (8) investigated the effects of HIV-1 infection on CD100 expression in T cells, and they observed that a subset of CD8⁺ T cells lacking membrane-associated CD100 showed decreased functional capacity. Their findings suggested that loss of CD100 expression would most likely lead to dysfunctional immunity in HIV-1 infection. It is well accepted that there are many differences between acute infections and chronic infections. Acute infections usually result in effective antiviral immune responses, while chronic infections are often associated with suboptimal T cell function. Furthermore, HIV infects CD4⁺ T cells and B cells and has profound pathological effects on the immune system, further confounding the interpretation of T cell dynamics following the acute phase. Therefore, it is important to investigate the functional role of CD100 in the CD8 antiviral response in acute infections.

Hantaan virus (HTNV), a member of the genus Hantavirus of the family Bunyaviridae, can cause a severe lethal hemorrhagic fever with renal syndrome (HFRS) in humans. More than 100,000 cases of HFRS, over 50% of which are documented in mainland China, occur annually worldwide, with a mortality rate of 2 to 10% (9, 10). People with HFRS are clinically characterized by having sudden fever, hemorrhage, thrombocytopenia, and acute renal failure. Typically, the course of HFRS involves the following five sequential stages: febrile, hypotensive, oliguric, diuretic, and convalescent. Hantaviruses have been documented to infect endothelial cells, causing a viremia that typically clears within the first 2 weeks after the onset of symptoms (11). A Hantaan virus outbreak in China has enabled us to longitudinally study the antiviral T cell response in the setting of a natural acute viral infection. In our previous study, membrane-associated CD100 expression on CD8 T cells was significantly decreased in the early phase of HFRS (12). Whether the change in CD100 expression was associated with cytotoxic T lymphocyte (CTL) function and how the regulation of CD100 expression balances antiviral immunity versus immunopathology during disease progression remain unclear.

In this study, plasma and peripheral blood mononuclear cell (PBMC) samples from 68 HFRS patients and 15 healthy controls were collected. The dynamic changes in CD100 expression on CD8⁺ cells, phenotype, functional markers, the virus-specific cellular immune response, and the relationship between low CD100 expression on CD8⁺ cells and plasma HTNV RNA load during the course of HFRS were analyzed.

MATERIALS AND METHODS

Ethics statement.

The study was approved by the Institutional Review Board of the Fourth Military Medical University. Written informed consent was obtained directly from each adult subject. The aims of the research were explained to the parents and guardians of participating children, and written consent was obtained from the guardians of participating children on behalf of the children for the collection of samples and subsequent analysis.

Patients.

Peripheral blood was collected from 68 hospitalized HFRS patients at Tangdu Hospital of the Fourth Military Medical University (Xi'an, China) from October 2012 to December 2012. The clinical diagnosis of HFRS was confirmed serologically by the detection of IgM and IgG antibodies to HTNV nucleocapsid protein (NP). The disease severity of HFRS could be classified on the basis of clinical and laboratory parameters used in the diagnostic criteria for HFRS in China as follows: (i) mild, mild renal failure without an obvious oliguric stage; (ii) moderate, obvious symptoms of uremia, effusion (bulbar conjunctiva), hemorrhage (skin and mucous membrane), and renal failure with a typical oliguric stage; (iii) severe, uremia, effusion (bulbar conjunctiva and either peritoneum or pleura), hemorrhage (skin and mucous membrane), and renal failure with oliguria (urine output, 50 to 500 ml/day) for ≤5 days or anuria (urine output, <50 ml/day) for ≤2 days; and (iv) critical, for those with ≥1 of the following symptoms during severe disease: refractory shock, visceral hemorrhage, heart failure, pulmonary edema, brain edema, severe secondary infection, and severe renal failure with either oliguria (urine output, 50 to 500 ml/day) for >5 days, anuria (urine output, <50 ml/day) for >2 days, or a blood urea nitrogen level of >42.84 mmol/liter. Healthy controls consisted of 21 uninfected healthy volunteers who had been matched with infected patients with respect to age and sex. Peripheral blood mononuclear cells (PBMCs) were isolated from anticoagulated blood via Ficoll gradient centrifugation. The freshly isolated PBMCs were used in all of the following assays.

Antibodies.

Peridinin chlorophyll protein (PerCP)-Cy5.5-CD3 monoclonal antibody (MAb), fluorescein isothiocyanate (FITC)-CD8α MAb, PerCP-Cy5.5-CD8α MAb, phycoerythrin (PE)-HLA-DR MAb, PerCP-Cy5.5-CD38 MAb, Ki67-PE MAb, FITC-CD127 MAb, PE-CD45RA MAb, PerCP-Cy5.5-CCR7 MAb, FITC-CD27 MAb, PE-CD62L MAb, PerCP-Cy5.5-PD-1 MAb, FITC-granzyme MAb, PE-perforin MAb, FITC-gamma interferon (IFN-γ) MAb, and PE-tumor necrosis factor alpha (TNF-α) MAb were purchased from BioLegend (San Diego, CA, USA), FITC-CD95 (Fas) MAb and PE-CD8β MAb were purchased from BD (San Diego, CA, USA), and CD100 MAb was prepared by our laboratory previously (13). Allophycocyanin (APC)-CD100 MAb (CD100–FMU-2.3) was labeled with APC using an DyLight antibody labeling kit (84535; Thermo Inc., USA).

Flow cytometry.

Fresh PBMCs were isolated from whole blood by density gradient centrifugation using standard procedures. For surface-expressed antigens, PBMCs (approximately 2 × 10⁶ cells/ml) were incubated with antibodies for 30 min at 4°C in the dark. For intracellular staining, cells were permeabilized using BD FACS-Perm2 (BD Biosciences) according to the manufacturer's instructions. After an additional wash, PBMCs were analyzed with four-color fluorescent-antibody staining. Flow cytometric acquisition and analysis were performed on at least 50,000 acquired events (gated on lymphocytes) on a FACSCalibur flow cytometer using CellQuest software (BD Pharmingen). Data analysis was performed using FlowJo software.

In vitro cytokine production.

To assess HTNV-specific responses, HTNV NP 9-mer peptide pools were used to stimulate the PBMCs of HFRS patients in vitro during the early phase. The peptide pools contained 14 peptides that were confirmed to be HTNV NP CTL epitopes in our previous study (14–16). Detailed information regarding the 14 epitopes can be found in Table 1. The freshly isolated PBMCs (2 × 10⁶ cells) were stimulated with HTNV NP 9-mer peptide pools (5 μg/ml) or were left unstimulated as controls. After 1 h of incubation, brefeldin A was added (10 μg/ml), and the cells were incubated at 37°C and 5% CO₂ for an additional 5 h. The T cell expression levels of IFN-γ and TNF-α were evaluated by intracellular cytokine staining.

TABLE 1.

HLA-A02, B07, and other class I molecules restricted by the HTNV nucleocapsid protein peptide pool

Amino acid position in HTNV NP	9-mer peptide sequence	HLA class I restriction
129–137	FVVPILLKA	HLA-A02
131–139	VPILLKALY	HLA-B35
247–255	LPDTAAVSL	HLA-B35
167–175	DVNGIRKPK	HLA-A33
277–285	ETKESKAIR	HLA-A33
197–205	RYRTAVCGL	HLA-A11
245–253	KLLPDTAAV	HLA-A24
258–266	GPATNRDYL	HLA-B07
133–141	ILLKALYML	NDa
176–184	HLYVSLPNA	ND
214–222	QMISPVMSV	ND
246–254	LLPDTAAVS	ND
247–255	LPDTAAVSL	ND
277–285	ETKESKAIR	ND

^aND, not defined.

HLA class I pentamer staining.

HLA-A2 refolded with the CD8⁺ T-cell 9-mer epitope FVVPILLKA (NP positions 129 to 137) was customized from Proimmune (Oxford, United Kingdom). The PBMCs of the patients or the healthy controls were stained with PE-labeled HLA class I pentamer for 10 min at room temperature and subsequently stained with CD3, CD8, and CD100 antibodies for 30 min at 37°C. A minimum of 300,000 total cells were acquired and gated on the CD3⁺ cells. The CD8⁺ pentamer⁺ T cell gate was set by the isotype control staining, and the cutoffs for positive staining were set by staining with an irrelevant pentamer.

Quantitative RT-PCR for HTNV.

RNA from patient plasma was extracted using a PureLink viral RNA/DNA kit (Invitrogen Life Technologies) according to the manufacturer's instructions. A SuperScript III Platinum one-step quantitative reverse transcription-PCR (RT-PCR) system kit (Invitrogen Life Technologies) was used for the real-time RT-PCR assay. The detailed procedure for real-time RT-PCR was performed as previously described (17) using a C1000 thermal cycler (Bio-Rad Laboratories). Analysis was performed with Bio-Rad CFX software. The numbers of viral RNA copies per milliliter were determined twice for every time point, and the mean of the results from the two separate experiments was used.

Statistics.

Statistical analysis was performed using GraphPad Prism 5.0. For parameter comparisons between subject groups, a Mann-Whitney U test was used. For single comparisons of matched groups, two-tailed paired Student t tests were performed. The data in dot plots represent the median, 25th and 75th percentiles, minimum, and maximum. Spearman's test was used for correlations. P values of less than 0.05 were considered significant.

RESULTS

Expansion of CD8^low CD100⁻ T cells in the early phase in HFRS patients.

A total of 68 HFRS patients were confirmed to have HTNV infection via detection of IgM or IgG antibodies specific to HTNV in patient serum specimens. Overall, 104 blood samples were collected at different stages of the disease, and 21 uninfected controls were included in the study. Because hantaviruses have been documented to infect endothelial cells, thus causing viremia that typically clears within the first 2 weeks after the onset of clinical symptom (11, 18), the disease course was divided into two phases for the purpose of this study: early phase (within 13 days after the onset of clinical symptoms; average, day 7) and late phase (more than 14 days after the onset of clinical symptoms; average, day 20).

PBMCs from individuals in the early phase or late phase and from uninfected controls were surface stained for CD100 and analyzed by flow cytometry. In the early-phase HFRS patients, three subsets of CD8⁺ T cells could be defined by the expression of CD8 and CD100: CD8^low CD100⁻, CD8^high CD100⁺, and CD8^low CD100⁺. Interestingly, the population of CD8^low CD100⁻ T cells was almost completely absent in uninfected controls. Furthermore, the CD8^low CD100⁻ subset appeared to decrease in the late phase of the disease and disappeared when the patients fully recovered. An anti-CD8β antibody was also used to stain the PBMCs of HFRS patients, and the CD8 molecule for the CD8^low CD100⁻ subset was confirmed to be CD8αβ, not CD8αα (Fig. 1A).

FIG 1.

Expansion of the CD8^low CD100⁻ subset in the early phase in HFRS patients. Three subsets of CD8 T cells (CD8^high CD100⁺, CD8^low CD100⁺, and CD8^low CD100⁻) were defined in PBMCs from HFRS patients based on the membrane expression of the CD100 and CD8 molecules. (A) Gating strategy and representative plots depicting the different CD8⁺ T cell subpopulations in a representative HFRS patient at different time points after symptom onset and in one representative uninfected control (NC). The distinct population of CD8^low CD100⁻ cells was observed only in HFRS patients and was absent in uninfected controls. (B) Comparison of the distribution of the three CD8⁺ T subsets in HFRS patients between early phase (within 13 days after symptom onset, n = 74) and late phase (more than 14 days after symptom onset, n = 29). (C) Kinetic changes in the CD8^low CD100⁻ subset in HFRS patients during the disease course within the same individuals (n = 16). Statistical analysis was performed using a Mann-Whitney U test (B) or a t test (C). Black lines represent medians, and P values are shown on each graph.

To determine whether the three CD8 cell subsets exhibited dynamic changes over the course of HFRS, all 104 samples were analyzed for CD100 expression. To account for the typical expansion in total CD8⁺ T cell counts that occurs during HTNV infection, the percentage of each subset of CD8⁺ T cells was assessed. The results showed that CD8^low CD100⁻ subsets accounted for a median of 14.3% (interquartile range [IQR], 7.6% to 22.2%) of total CD8⁺ T cells in the early phase and decreased to 2.7% (1.2% to 5.6%) of CD8 T cells in late phase (P < 0.0001). CD8^high CD100⁺ cells accounted for the majority of total CD8⁺ cells and exhibited a small but insignificant increase in frequency in the late phase (73.4% [60.3% to 79.7%]) compared with the early phase (66.7% [50.5% to 76.8%]), whereas the CD8^low CD100⁺ subset represented 8.2% (5.2% to 15.9%) of the total CD8⁺ cells in the early phase and increased to 17.8% (13.9% to 24.4%) in the late phase (P < 0.0001) (Fig. 1B). In addition, we performed a longitudinal analysis on 16 HFRS patients over the disease course, and there was a trend toward a lower frequency of the CD8^low CD100⁻ subset in the late phase compared with the early phase (paired t test, P < 0.0001) (Fig. 1C).

CD8^low CD100⁻ T cells are associated with high activation and express high levels of cytolytic effector molecules.

PBMCs from early-phase HTNV-infected patients (HFRS) and uninfected controls (NC) were assessed. The expression levels of Ki67, CD38, and HLA-DR on CD8⁺ T cells were analyzed to evaluate the responding CD8⁺ cells. Specifically, CD38 expression on the CD8^low CD100⁻ subset (median, 99.3%; IQR, 99.1% to 99.6%) was significantly higher than that on the other two subsets at the early phase of HTNV infection (85.3% [73.9% to 88.4%]) for CD8^high CD100⁺ and 88.5% [87.7% to 91.7%] for CD8^low CD100⁺), whereas the CD8^low CD100⁻ subset was not present in healthy individuals. CD38 expression on the CD8^high CD100⁺ subset and CD8^low CD100⁺ subset was 21.6% (10.6% to 31.1%) and 65.6% (64.4% to 71.8%), respectively (Fig. 2A). HLA-DR expression on the CD8^low CD100⁻ subset (72.9% [75.5% to 96.3]) was also significantly higher than that on the other two subsets in the early HTNV-infected patients (49.1% [41.1% to 58.8%] for CD8^high CD100⁺ and 30.8% [28.4% to 33.2%] for CD8^low CD100⁺). In healthy individuals, HLA-DR expression was 24.0% (19.7% to 38.7%) for CD8^high CD100⁺ and was 34.1% (19% to 41.4%) for CD8^low CD100⁺ (Fig. 2B). The percentage of Ki67-positive cells among the CD8^low CD100⁻ subset (23.7% [16.7% to 26.4%]) was also higher than those among the other two subsets in HFRS patients, whereas there were almost undetectable levels of Ki67-positive cells in both the CD8^high CD100⁺ and CD8^high CD100⁺ subsets from healthy individuals (Fig. 2C). The expression of the effector molecules granzyme B and perforin was also assessed. In HFRS patients, CD8^low CD100⁻ cells comprised a higher proportion of cells expressing perforin (95.8% [94.5% to 97.4%]) and granzyme B (94.6% [92.5% to 95.9%]) than the CD8^high CD100⁺ or CD8^low CD100⁺ T cell subsets. In healthy individuals, expression of granzyme B (41.9% [30.1% to 50.9%]) and perforin (32.8% [28.2% to 38.7%]) on CD8^high CD100⁺ cells was much lower than that on CD8^low CD100⁺ T cells (granzyme B, 85.7% [79.7% to 88.0%]; perforin, 80.9% [77.2%, 88.1%]) (Fig. 2D to F). These results showed that the majority of CD8⁺ T cells in healthy individuals, which were CD8^high CD100⁺ cells, expressed low levels of activation markers and effector molecules, and the CD8^low CD100⁺ cells in HFRS patients expressed relatively high levels of CD38 and effector molecules compared to CD8^high CD100⁺ cells. In the early-phase HTNV-infected patients, the CD8^low CD100⁻ subset exhibited distinct activation markers and high expression of effector molecules among the three CD8 subsets, suggesting that the CD8^low CD100⁻ T cells are responding CD8⁺ T cells.

FIG 2.

CD8^low CD100⁻ T cells are highly activated and express cytolytic molecules after HTNV infection. (A and B) PBMCs from early HTNV-infected patients (HFRS) and uninfected controls (NC) were assessed for CD38 (A) and HLA-DR (B) surface staining. (C) Cell proliferation was analyzed by Ki67 staining. (D and E). The cytotoxic effector molecules (D) granzyme B and (E) perforin were detected by intracellular staining. Numbers represent the positive expression rate (%) of each molecule in the three CD8 T cell subsets. (F) Representative dot plots of granzyme B and perforin expression in the three CD8⁺ T cell subsets. The CD8^low CD100⁻ subset expressed the highest levels of all the detected markers among the three CD8 T cell subsets (n = 6 for all groups). Box plots show median, 25th and 75th percentiles, minimum, and maximum. Statistical analysis was performed using a paired t test. P values are shown on each graph.

Phenotype and functional characteristics of CD8^low CD100⁻ T cells.

We performed a detailed phenotypic and functional characterization of CD8 subsets from HFRS patients in the early phase and from uninfected controls. The surface expression of representative phenotypic markers was analyzed to characterize the differentiation and memory status of the CD8 cell subsets, especially the CD8^low CD100⁻ subset. When the surface expression of CCR7 and CD45RA was analyzed simultaneously, we found that CD8^low CD100⁻ cells consisted primarily of CCR7⁻ CD45RA⁻ (mean, 70.6%) and CCR7⁺ CD45RA⁻ (mean, 25.3%) cells in the HFRS patients. The other two CD8 T cell subsets demonstrated heterogeneous levels of CCR7 and CD45RA expression. The highest proportions (mean, 75.9%) of CD8^high CD100⁺ cells were CCR7⁻ CD45RA⁻ cells, and the highest proportions (mean, 54.8%) of CD8^high CD100⁺ cells were CCR7⁻ CD45RA⁺ cells. However, the majority (mean, 76.1%) of CD8^high CD100⁺ cells were CCR7⁺ CD45RA⁺ cells, and the highest proportions (mean, 74.2%) of CD8^low CD100⁺ cells were CCR7⁻ CD45RA⁺ cells in healthy controls (Fig. 3A and B). We next characterized CD8⁺ T cells for surface expression of CD127, which is necessary for the transduction of interleukin7 (IL-7) signaling and essential for the survival of memory T cells. The results indicated that CD8^low CD100⁻ T cells expressed significantly higher levels of CD127 (median fluorescence intensity [MFI], 28.9; IQR, 21.0 to 41.1) than CD8^high CD100⁺ cells (8.8 [7.3 to 10.4]) or CD8^low CD100⁺ cells (10.3 [9.3 to 11.1]) in HFRS patients, whereas CD127 expression on CD8^high CD100⁺ cells and CD8^low CD100⁺ cells was 11.9 (7.6 to 15.6) and 5.3 (4.5 to 14.3), respectively, in healthy controls (Fig. 3C and D). Because the CD28 and CD27 markers are also involved in CD8⁺ T cell differentiation, we assessed CD28 and CD27 expression in the CD8⁺ T cell subsets. In HFRS patients, the lowest levels of CD28 were expressed on CD8^low CD100⁻ T cells (MFI, 9.2 [6.9 to12.6]) compared with CD8^high CD100⁺ T cells (22.2 [20.6 to 53.8]) or CD8^low CD100⁺ T cells (16.2 [15.1 to 20.4]), while CD28 expression on CD8^high CD100⁺ cells (29.2 [20.5 to 32.4]) was much higher than that on CD8^high CD100⁺ cells (4.4 [3.8 to 4.5]) in healthy controls (Fig. 3E and F). The CD27 expression levels of CD8^low CD100⁻ T cells in HFRS patients (MFI, 18.6 [16.2 to 27.6]) were intermediate and lower than that of CD8^high CD100⁺ T cells (109.5 [94 to 123.8]) but higher than that of CD8^low CD100⁺ cells (7.2 [6.2 to 12.2]). In healthy controls, CD27 expression on CD8^high CD100⁺ cells was 17.4 (5.4 to 52.1) and 4.7 (4.4 to 22.0) on CD8^low CD100⁺ cells (Fig. 3G and H). CD62L is necessary for memory cell homing to secondary lymphoid organs. We found that the CD8^low CD100⁻ T cell subsets in HFRS patients presented a CD62L⁻ phenotype. CD8^high CD100⁺ cells in HFRS patients and healthy controls expressed high levels of CD62L, and a small proportion of CD8^high CD100⁺ cells expressed CD62L (Fig. 3I and J). To summarize, CD8^low CD100⁻ T cells have a distinct expression profile of effector CD8⁺ T cells (CCR7^+/− CD45RA⁻ CD127^high CD27^int CD28^low CD62L⁻). Interestingly, CCR7 and CD127, markers closely associated with memory T cells, were found mainly in the CD8^low CD100⁻ subset, which implied that a portion of cells in the CD8^low CD100⁻ subset may be involved in the development and maintenance of CD8 T cell memory.

FIG 3.

CD8^low CD100⁻ T cells express phenotypic and functional markers of effector T cells. (A and B) Coexpression of CCR7 and CD45RA was determined and summarized for each CD8 T cell subset in the early phase in HFRS patients and uninfected controls. (C to J) Expression of CD127 (C and D), CD28 (E and F), CD27 (G and H), and CD62L (I and J) among the three CD8 T cell subsets from a representative HFRS patient in the early phase and an uninfected control (n = 4 for all groups). Box plots show median, 25th and 75th percentiles, minimum, and maximum. Statistical analysis was performed using a paired t test. P values are shown on each graph.

CD8^low CD100⁻ T cells secrete the highest levels of IFN-γ and TNF-α when stimulated with HTNV NP-specific peptide pools.

To assess the responses of the three CD8 subsets, especially CD8^low CD100⁻ cells, to virus-derived peptides, the cytokine secretion properties of CD8 cells stimulated with HTNV-NP derived 9-mer peptide pools or without peptide stimulation were determined by intracellular cytokine staining. After stimulation, CD8^low CD100⁻ cells produced large amounts of TNF-α and IFN-γ, whereas the other two subsets of CD8⁺ T cells produced small amounts of TNF-α and IFN-γ. To quantitate how many cells responding were CD8^low CD100⁻ cells, the IFN-γ-producing cells from among the total number of CD8⁺ T cells and from each cell subset were plotted first (Fig. 4A). Then, the results of 3 independent experiments exhibiting the proportions of each CD8⁺ T subset among total IFN-γ-positive CD8⁺ T cells were summarized (Fig. 4C). TNF-α-producing CD8⁺ T cells were analyzed by the same method (Fig. 4B and D). The results showed that the CD8^low CD100⁻ subset accounted for an average of 75.1% of total IFN-γ-producing CD8 T cells and 81.1% of total TNF-α-positive CD8⁺ T cells.

FIG 4.

CD8^low CD100⁻ T cells secrete the highest levels of IFN-γ and TNF-α among the three CD8 T cell subsets. PBMCs from early-phase HFRS patients were stimulated with peptide pools of HTNV NP for 6 h and subsequently examined by intracellular staining and flow cytometry. (A and B) Representative plots of IFN-γ (A)- and TNF-α (B)-producing cells from the total number of CD8⁺ T cells and from each CD8⁺ T cell subset in response to HTNV NP 9-mer peptide pools or without peptide stimulation. (C and D) Summary of 3 independent experiments indicating the proportion of each CD8 subset among total IFN-γ-positive (C) and TNF-α-positive (D) CD8 T cells after HTNV-specific peptide pool stimulation. Statistical analysis was performed using a paired t test.

Higher frequencies of CD100⁻ cells among antigen-specific CD8⁺ T cells in milder disease and an inverse correlation between the CD8^low CD100⁻ T cell percentage and viral load.

We next assessed how many antigen-specific CD8⁺ T cells were derived from CD8^low CD100⁻ cells, HTNV-specific CD8⁺ T cell responses were evaluated by pentamer staining. Given that all CD100⁻ cells express low levels of the CD8 molecule in CD8⁺ T cells from HFRS patients, the frequency of CD100⁻ cells among the total epitope-specific CD8⁺ T cells could be considered the proportion of CD8^low CD100⁻ cells among HTNV-specific CD8⁺ T cells. A total of 18 samples from HLA-A2-restricted patients at the early phase of HFRS were investigated. The results showed that the frequency of CD100⁻ cells among the total epitope-specific CD8⁺ T cells varied greatly among different patients (from 18.0% to 88.4%). We then wanted to explore whether this difference is related to disease severity. According to the diagnostic criteria described in Materials and Methods, the HFRS patients were classified as mild, moderate, severe, or critical clinical types. In the present study, to ensure the sample size for statistical analyses, we combined the patients according to disease severity into mild/moderate and severe/critical groups for comparison. We compared the frequencies of CD100⁻ cells among epitope-specific CD8⁺ T cells between patients in the mild/moderate group and patients in the severe/critical group at the early-phase time point in the HLA-A2-positive patients by pentamer staining. The statistical analyses showed that the frequency of CD100⁻ cells among the HLA-A2-restricted epitope-specific CD8⁺ T cells was higher in the mild/moderate group of patients than that in the severe/critical group (Mann-Whitney U test, P = 0.044) (Fig. 5A and B).

FIG 5.

Higher frequencies of CD100⁻ cells among antigen-specific CD8⁺ cells in milder disease and an inverse correlation between CD8^low CD100⁻ T cell percentage and viral load. (A) Representative dot plot of CD100 expression on CD8⁺ T cells costained with pentamer folded with HLA-A2-restricted HTNV NP-derived peptide (FVVPILLKA) in the early phase from one representative HFRS patient. Pen⁺, pentamer-positive cells. Right, histogram of CD100 expression by pentamer-specific CD8 T cells. (B). Frequency of CD100⁻ cells in HTNV-specific CD8⁺ T cells from the early phase of different disease severities (n = 6 for mild/moderate, n = 12 for severe/critical). Statistical analysis was performed using a Mann-Whitney U test. P values are shown on each graph. The results showed that the CD100⁻ cell frequency of HTNV-specific CD8⁺ T cells is higher in the mild/moderate group than in the severe/critical group. (C) Correlations between the frequency of CD8^low CD100⁻ cells and plasma viral load in the early phase of HFRS. There was a significant inverse correlation between the frequency of the CD8^low CD100⁻ subset and plasma viral load (n = 72). Correlation statistics were analyzed using the Spearman correlation. The r and P values are shown on the graph.

To analyze the correlation between viral load and the frequency of CD8^low CD100⁻ T cells among CD8 cells in HFRS patients, all blood samples collected in the early phase were used for viral load analysis. In total, 78 samples were tested, and viral RNA could be detected in 72 of them. In general, viral load decreased with the disease course. The viral load within 7 days after symptom onset was higher than that at 8 to 13 days after symptom onset. However, the number of CD8^low CD100⁻ T cells contrasted with the decreased trend in viral load. Next, the relationship between viral load in the early phase and the frequency of CD8^low CD100⁻ T cells among CD8⁺ T cells was analyzed. The results indicated that the frequency of CD8^low CD100⁻ cells was inversely associated with viral load levels (Spearman's test, P < 0.0001 and r = −0.42) (Fig. 5C).

Expression of inhibitory receptors on CD8^low CD100⁻ T cells.

Given that the CD8^low CD100⁻ T subset appeared to decrease in the late phase of the disease and disappeared in the convalescent stage, we wanted to explore the mechanisms that may regulate the kinetic changes of this subset. The expression of inhibitory receptors may regulate the effector response to pathogens and ensure T cell tolerance under steady-state conditions. In this regard, we studied the expression of PD-1 and Fas on CD8⁺ T cells of HFRS patients. The results showed that on CD8^low CD100⁻ T cells, PD-1 was consistently highly expressed throughout, although expression somewhat decreased over the course of the disease. PD-1 expression on the CD8^high CD100⁺ subset was heterogeneous in the early phase and became low in the late phase. PD-1 expression on the CD8^low CD100⁺ subset was quite low throughout the full disease course (Fig. 6A and B). In contrast, Fas expression on the CD8^low CD100⁻ subset was much lower (median, 45.9%) than that on the other two CD8⁺ T subsets (81.3% on CD8^high CD100⁺ and 72.8% on CD8^low CD100⁺) (Fig. 6C and D). These results provide the possibility that the apparent decrease in CD8^low CD100⁻ T cell number in the late phase of the disease may be regulated by the PD-1 pathway.

FIG 6.

CD8^low CD100⁻ T cells express high levels of PD-1 but not Fas during the full disease course. (A) Expression of PD-1 among the three CD8 T cell subsets from one representative HFRS patient in the early phase and late phase. (B) Comparison of frequencies of PD-1 expression among the three CD8 T cell subsets from HFRS patients between the early phase and late phase (n = 13). Statistical analysis was performed using a t test. (C) Expression of Fas among the three CD8 T cell subsets from one representative HFRS patient in the early phase. (D). Frequency of Fas expression among the three CD8 T cell subsets from HFRS patients in the early phase (n = 6). Statistical analysis was performed using a Mann-Whitney U test. Black lines represent medians. P values are shown on each graph.

DISCUSSION

The role of CD8 T cells in the pathogenesis of HFRS and the immune response to hantavirus infection has been the subject of much debate. In our previous study, HTNV RNA could be detected in the plasma of HFRS patients only at 3 to 12 days after the onset of the disease, and the viral load gradually decreased to an undetectable level with the course of the disease, which is in accordance with the self-limiting properties of HFRS. There is a statistically significant association between plasma HTNV RNA load in the early stage and disease severity in patients, but there is no relationship between HTNV RNA load and the peak value of the titers of HTNV NP-specific IgM and IgG (17). Our previous studies also showed that the magnitude of the cellular immune response and the frequency of the epitope-specific pentamer-positive CD8⁺ T cell response were much higher in patients with mild/moderate HFRS than in those with severe/critical disease at the acute stage of the disease. These results indicated that both HTNV clearance and the CD8⁺ T cellular immune response contribute to a better outcome for HFRS disease processes. In this study, we found the expansion of a CD8^low CD100⁻ subset in the early phase in HFRS patients, which was not present in PBMCs from normal controls. Importantly, the frequency of the CD8^low CD100⁻ T cell subset was inversely associated with plasma HTNV viral load level, and these cells produced high levels of IFN-γ and TNF-α in response to HTNV NP peptide pool stimulation. Taken together, these data indicated that the expansion of the CD8^low CD100⁻ T cells may be related to HTNV clearance and therefore might play a protective role in the antiviral response during HTNV infection. The protective function of the CD8^low CD100⁻ T cell subset during HFRS may be useful for understanding the mechanisms of the CD8 cell-mediated antiviral response to HTNV infection.

The following additional important questions arose from this study: how are CD8^low CD100⁻ T cells generated, and what is the function of these cells in the antiviral cellular immune response? Previous studies have shown that membrane-associated CD100 expression on T cells decreases with certain types of stimulation. Phorbol myristate acetate (PMA) stimulation or CD100 monoclonal antibody cross-linking downregulates CD100 expression on PBMCs in vitro (1). CD100 was reported to be associated with the protein tyrosine phosphatase CD45. This association is enhanced during T cell activation and downregulates CD100 molecules at the cell surface (19). To further assess which type of stimulation resulted in diminished CD100 expression on CD8⁺ T cells after HIV infection, Eriksson et al. (8) assessed the effects of various stimulating conditions on CD100 expression using both HIV-1-infected and uninfected donors. The results showed that HIV infection for 7 days, proinflammatory (IL-1β, IL-6, IL-12 IL-18, IFN-γ, and TNF-α) and homeostatic (IL-7 and IL-15) cytokines, and direct T cell receptor (TCR) stimulation using anti-CD3 and anti-CD28 antibodies did not change the frequency or geometric MFI of CD100 expression, whereas virus-specific peptide stimulation induced a rapid loss of CD100 expression on responding T cells (8). CD100 is also essential for the interaction between T cells and APCs. CD100 expressed on T cells can interact with DCs to promote their activation and maturation, which in turn enhances T cell activation (7). In this study, CD8^low CD100⁻ T cells highly expressed activation markers, and furthermore, when stimulated by HTNV NP-specific peptide pools, most responding CD8⁺ T cells (IFN-γ⁺ and/or TNF-α⁺) were CD8^low CD100⁻ cells. Therefore, it is tempting to assume that the downregulation of CD100 expression on CD8⁺ T cells may result from the interaction between CD8⁺ T cells and APCs under stimulation with virus-specific peptides. Studies in murine models have indicated that the CD100-CD72 interaction is required for effective APC presentation of peptides to antigen-specific T cells (20). As a ligand in T cell activation, CD100 significantly enhances the costimulatory activity of dendritic cells and the priming of antigen-specific T cells (21). In the present study, the CD100⁻ frequency of the HTNV-specific CD8⁺ T cells was higher in milder HFRS cases than in more severe cases at the early phase of the disease, which may imply that CD100⁻ CD8^low cells are more effective than CD100⁺ CD8⁺ T cells in the anti-HTNV immune response among the virus-specific CD8⁺ T cells due to the activation of the CD100 signal. As a consequence, CD8^low CD100⁻ cells maybe rich in activated HTNV-specific T cells and can function as a protective CD8⁺ T cell subset during HTNV infection.

In response to viral infection, CD8⁺ T cells undergo a dramatic, highly orchestrated activation and differentiation process. An initial encounter with an appropriately activated antigen-presenting cell leads to an exponential increase in antigen-specific CD8⁺ T cell numbers and the gain of effector functions. After viral clearance, only 5 to 10% of virus-specific effector T cells survive and gradually acquire memory properties, while most effector cells are subject to activation-induced cell death (22–25). In this study, CD8^low CD100⁻ T cells highly expressed cytolytic effector molecules (perforin and granzyme B) and exhibited the phenotypes of CCR7^+/− CD45RA⁻, CD127^high CD27^int CD28^low, and CD62L⁻, which were consistent with the major features of effector CD8⁺ T cells. Some studies have shown that after virus infection or intracellular antigen stimulation, effector CD8 T cell subpopulations display the CD8^low phenotype, which was consistent with our findings (26–28). CD8 effector T cells can mediate antiviral protection via perforin-dependent cytotoxicity and/or Fas ligand (CD95L)-Fas (CD95) interaction pathways. Our results showed abundant intracellular perforin and granzyme B but not surface FasL expression (data not shown) in CD8^low CD100⁻ cells, suggesting that the protection against HTNV infection by CD8^low CD100⁻ cells may be perforin/granzyme B dependent. Moreover, CD8^low CD100⁻ cells produce large amounts of IFN-γ when stimulated by HTNV-specific peptides, which could increase the expression of major histocompatibility complex (MHC) class I and other molecules involved in viral peptide loading. This increase can facilitate the recognition of HTNV-infected target cells by CTL. In addition, IFN-γ could also upregulate Fas expression on target cells and renders them susceptible to Fas ligand-mediated apoptosis (29). In the late phase of the disease, HTNV viral load cannot be detected in the plasma of HFRS patients; meanwhile, the frequency of CD8^low CD100⁻ T cells undergoes a dramatic decrease. The dynamic changes in the CD8^low CD100⁻ T cell subset are in accordance with the theoretical pattern of the effector CD8 T cell antiviral response. Lindgren et al. (30) found that there are high numbers of Ki67⁺ CD38⁺ HLA-DR⁺ effector CD8 T cells among the PBMCs of patients during the early phase after Puumala virus infection, and this subset subsequently contracted in parallel with a decrease in viral load, which is also consistent with our results. In our study, PD-1 was found to be consistently and highly expressed on the CD8^low CD100⁻ T cell subset during the disease course, which may represent the possibility that the apparent decrease in the CD8^low CD100⁻ T cell number in the late phase of the disease may be regulated by the PD-1 pathway. Memory CD8 T cells are an important component of protective immunity against viral infections. Previous studies have described high frequencies of hantavirus-specific memory CD8 T cells in PBMCs of individuals who recovered from HTNV infection (31, 32). These findings, together with the absence of any evidence for hantavirus persistence or symptomatic reinfection in humans, have suggested a role for CD8 T cells in the generation of protective long-lasting immunity. However, the differentiation process of virus-specific memory CD8 T cells is far from clearly understood. It is generally accepted that throughout the course of infection, there is considerable heterogeneity in the populations of effector CD8⁺ T cells, which can be phenotypically separated into different populations, including short-lived effector cells (SLECs) and memory precursor effector cells (MPECs) (33, 34). SLECs are short-lived, play an active role in virus clearance, and undergo apoptosis once the pathogen is eliminated. In contrast, MPECs survive and give rise to self-renewing memory cells. Some phenotype markers are considered to be more highly expressed on MPECs than on SLECs. Interestingly, the phenotype markers closely related to memory precursor effector T cells were found mainly in the CD8^low CD100⁻ T cell subset, which indicated that a portion of the CD8^low CD100⁻ T cells might be involved in the development and maintenance of CD8 T cell memory.

In summary, for the first time, we report a novel subset of CD8^low CD100⁻ T cells among PBMCs from HFRS patients that expanded in the early phase of disease and contracted in the late phase. The proportion of the CD8^low CD100⁻ cells among CD8⁺ T cells from HFRS patients was negatively correlated with the HTNV viral load, and the frequency of CD8^low CD100⁻ among the HLA-A2-restricted epitope-specific CD8⁺ T cells was higher in milder HFRS cases than in more severe cases. These results may be useful for our understanding of CD8⁺ T cell-mediated protective immunity as well as the T cell contraction participating in immune homeostasis during HTNV infection in humans. Whether CD8^low CD100⁻ T cells exist in other infectious diseases must be further investigated.