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Journal of Virology, June 2009, p. 5442-5450, Vol. 83, No. 11
0022-538X/09/$08.00+0     doi:10.1128/JVI.00106-09
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Differential Effect of Human Herpesvirus 6A on Cell Division and Apoptosis among Naïve and Central and Effector Memory CD4⁺ and CD8⁺ T-Cell Subsets

Sudhir Gupta,^* Sudhanshu Agrawal, and Sastry Gollapudi

Division of Basic and Clinical Immunology, University of California, Irvine, California

Received 14 January 2009/ Accepted 13 March 2009

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The immune responses of naive and different memory subsets of CD4⁺ and CD8⁺ T cells to human herpesvirus 6 (HHV-6) have not been previously investigated. We show that HHV-6A induces cell division, as measured by 5,6-carboxyfluorescein succinimidyl ester dye and flow cytometry, predominantly in two populations of effector memory CD4⁺ and CD8⁺ T cells (T_EM and T_EMRA); naïve (T_N) and central memory (T_CM) CD4⁺ and CD8⁺ T cells showed almost no cell division. In contrast, HHV-6A induced apoptosis primarily in T_N and T_CM CD4⁺ and CD8⁺ T cells, whereas T_EM and T_EMRA CD4⁺ and CD8⁺ T cells were resistant to HHV-6A-induced apoptosis. HHV-6A-induced apoptosis was associated with activation of caspase-8, caspase-9, and caspase-3, suggesting the involvement of death receptor and mitochondrial signaling pathways. In addition, HHV-6A induced secretion of interleukin-6 (IL-6), tumor necrosis factor alpha (TNF-), IL-8, and gamma interferon by peripheral blood mononuclear cells; TNF- secretion was observed exclusively from CCR7⁺ (T_N plus T_CM) CD4⁺ T cells. These data show that HHV-6 differentially influences the functions of naïve T cells and different subsets of memory CD4⁺ and CD8⁺ T cells, which in part may be due to differential susceptibility to HHV-6A-induced apoptosis.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Human herpesvirus 6 (HHV-6) was first discovered in patients with AIDS and lymphoma (24). In the last 2 decades, marked progress has been made in understanding the molecular biology of this virus, its tropism, and its seroprevalence. HHV-6 consists of two closely related yet distinct subgroups, HHV-6A and HHV-6B (2, 6). Yamanishi et al. (37) identified HHV-6B as an etiological agent of sixth disease or roseola infantum. Based upon serological and PCR techniques, both HHV-6 subgroups have been implicated in a number of disorders, including infectious mononucleosis (Epstein-Barr virus negative), encephalitis, multiple sclerosis, chronic myelopathy, bone marrow transplants, solid organ transplants, leukemia and lymphoma, Kaposi's sarcoma, collagen vascular diseases, and chronic fatigue syndrome (6, 7). HHV-6 grows most efficiently in primary CD4⁺ T cells; however, in vivo, HHV-6 has been found in lymphocytes, monocytes, macrophages, salivary glands, renal endothelial cells, neurons, and oligodendrocytes. Specific immune responses to HHV-6 include, but are not limited to, suppression of a proliferative response to mitogens, regulation of cytokine production, downregulation of T-cell receptor and impairment of T-cell activation, and inhibition of dendritic cell maturation and functions (5, 7, 10, 11, 27, 29, 34). However, there is no information regarding the responses of CD8⁺ T cells to HHV-6, which are the most critical cells in protection against virus infection, and the responses of different memory CD4⁺ and CD8⁺ T cells to HHV-6.

Memory and effector T cells play an important role in virus eradication and/or by secretion of various cytokines in the suppression of viral replication. Recent work has suggested that following virus infection or antigen stimulation, T cells undergo a series of proliferative and differentiation steps ultimately culminating in the acquisition and maintenance of memory for a particular antigen/pathogen (21, 32, 33, 36, 40). Naïve T cells (T_N) following exposure to a virus/antigen undergo clonal expansion, followed by clearance of the virus/antigen. This phase is followed by a phase of contraction during which virus-specific T cells undergo apoptosis, and then a number of virus-specific T cells stabilize and remain as memory T cells. One of the hallmarks of memory T cells is their capacity to undergo antigen-independent homeostatic turnover and thus maintain a stable pool of antigen-specific memory T cells (12, 28). Memory T cells display differential expression of adhesion molecules (CD62L) and chemokine receptors (CCR-7), which allow them to extravasate into lymphoid and nonlymphoid tissues and respond to microbes at peripheral tissue sites (3, 8, 14, 22, 25, 35). CD62L interacts with peripheral lymph node addressin on high endothelial venules (3), whereas CCR7 binds the chemokines CCL19 and CCL21 that are present on the luminal surface of endothelial cells in the lymph nodes (8). Therefore, CCR7⁺ and CD62^high T cells are found in lymph nodes whereas CCR7^– and CD62L^low T cells are found at extranodal sites such as the liver and lung. Based upon these adhesion molecules and chemokine receptors, memory CD4⁺ and CD8⁺ T cells have been divided into "central" memory T cells (T_CM) that are found in lymphoid organs and "effector" memory T cells (T_EM) that are found in peripheral nonlymphoid tissues and mucosal sites (21-25). T_EM T cells, especially CD8⁺ T cells, are further subdivided into two subpopulations based upon the presence or absence of CD45RA antigen and termed T_EM (CD45RA^–) and T_EMRA (CD45RA⁺) (12, 14, 25, 32). T_EMRA CD4⁺ T cells represent a very small population of effector memory CD4⁺ T cells (1 to 2%), whereas T_EMRA CD8⁺ T cells constitute a significant population of effector memory CD8⁺ T cells (10 to 12%). T_N and T_CM T cells proliferate significantly more than T_EM and T_EMRA T cells (28, 33, 40).

The purpose of the present study was to investigate the response of T_N cells and various subsets of memory CD4 and CD8⁺ T cells to HHV-6A. We demonstrate that HHV-6A induces cell division in T_EM and T_EMRA CD4⁺ and CD8⁺ T cells, which are resistant to apoptosis, whereas T_N and T_CM CD4⁺ and CD8 T cells display increased sensitivity to HHV-6A-induced apoptosis and minimal or no cell division. Furthermore, HHV-6A induces tumor necrosis factor alpha (TNF-) production exclusively by CCR7⁺ (T_N plus T_CM) CD4⁺ T cells.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Subjects. Peripheral blood mononuclear cells (MNCs) were isolated from the blood of healthy young subjects by Ficoll-Hypaque density gradient centrifugation. The protocol used was approved by the Human Subject Committee of the Institutional Review Board of the University of California, Irvine. HHV-6A lysates were provided by Advanced Biotechnologies, Inc., Columbia, MD. HHV-6A strain GS was grown in HSB-2 cells free of mycoplasma (>80% of the cells were infected with HHV-6A, as shown by testing for the presence of core protein with gp116/64/54 monoclonal antibodies [MAbs]) and purified by differential centrifugation, and a virus pallet was made through high-speed centrifugation through a sucrose gradient. Purified virus was lysed with 0.25% Triton X-100. Supernatants were collected, filtered, and pelleted as described above (1). Supernatants from uninfected HSB-2 cells served as controls. Pellets were resuspended in buffer and examined by electron microscopy for virions, and the count was 10¹¹ to 10¹² virus particles. Virus stocks were tested and found to be negative for bacteria and fungi. The only cytokine we tested was TNF- (since it induced apoptosis), and virus lysates were negative. A 1:1,000 dilution of a stock solution of HHV-6A and a control (determined after performing a concentration-dependent response) was used.

Reagents and chemicals. MAbs directly conjugated with fluorescein isothiocyanate (FITC), phycoerythrin (PE), allophycocyanin (APC), and peridinin chlorophyll protein (PerCP) were used. FITC-labeled anti-CD46, PerCP-labeled anti-CD4 and anti-CD8, APC-labeled anti-CD45, and isotype control antibodies were purchased from BD Biosciences, San Diego, CA. PE- and FITC-labeled anti-CCR7 antibodies were obtained from R&D Systems, Minneapolis, MN, and PE-labeled anti-DR5 antibodies were obtained from eBioscience, San Diego, CA. PE-labeled anti-TNF receptor 1 (anti-TNFR1) (CD120a) and anti-TNFR2 (CD120b) and isotype control antibodies were from CalTag Laboratories, Burlingame, CA. FITC-labeled anti-Bcl-2 and isotype control antibodies were purchased from Dako Corporation, Carpentaria, CA. PE-labeled anti-CD95L antibodies were from MBL International, Woburn, MA. Peptide substrates that recognize cleaved caspase-8 (FAM-LETD-FMK), cleaved caspase-9 (FAM-LETD-FMK), and cleaved caspase-3 (FAM-DEVD-FMK) were obtained from Cell Technology, Inc., Mountain View, CA. 5,6-Carboxyfluorescein diacetate succinimidyl ester (CFSASE) was obtained from Molecular Probes, Eugene, OR. Anti-CD3 and anti-CCR7⁺ magnetic beads were purchased from Stem Cell Technology, Vancouver, British Columbia, Canada.

Immunophenotyping. Peripheral blood MNCs incubated with control lysates or with HHV-6A for 5 days were stained with PerCP-conjugated anti-CD4 or anti-CD8, APC-conjugated anti-CD45RA, FITC-conjugated anti-CCR7, and PE-conjugated anti-TNFR1, anti-TNFR2, anti-DR5, or anti-CD95L antibodies. After staining, cells were washed extensively with phosphate-buffered saline (PBS) and analyzed. Flow cytometry was performed with a FACScalibur (Becton Dickinson, San Jose, CA) equipped with an argon ion laser emitting at 488 nm (for FITC, PE, and PerCP excitation) and a spatially separate diode laser emitting at 631 nm (for APC excitation). Forward and side scatters were used to gate and exclude cellular debris, and FL3 channels were used to gate CD4⁺ or CD8⁺ T cells. During analysis, an electronic gate for CD4⁺ or CD8⁺ T cells was used and the expression of CD45RA and CCR7 was determined. This allowed us to obtain the percentages of cells uniquely identifying CD45RA⁺ CCR7⁺ (T_N) CD45RA^– CCR7⁺ (T_CM), CD45RA^– CCR7^– (T_EM), and CD45RA^– CCR7⁺ (T_EMRA) CD4⁺ and CD8⁺ T-cell subsets. An electronic gate was placed on the above-defined CD4⁺ or CD8⁺ T-cell subsets, and TNFR1, TNFR2, DR5, and CD95L expression was analyzed by using red fluorescence (FL2) on the x axis against the cell number on the y axis. Ten thousand cells were acquired and analyzed with CellQuest software. Expression of CD46 on various subsets of CD4 and CD8 T cells was analyzed by multicolor flow cytometry.

Determination of Bcl-2 expression. Bcl-2 is an intracellular protein. Briefly, cells (2 x 10⁶/ml) were cultured with or without HHV-6A for 5 days at 37°C. After incubation, cells were stained with PerCP-labeled anti-CD4 or anti-CD8, APC-labeled anti-CD45RA, and PE-labeled anti-CCR7 antibodies. Stained cells were fixed with 2% paraformaldehyde, permeabilized with Perm-2 buffer (BD Biosciences, San Diego, CA), and stained with FITC-labeled anti-Bcl-2 or isotype control antibodies. Bcl-2 staining in T-cell subsets was analyzed by FACScalibur as described above.

Isolation of CCR7⁺ and CCR7^– T-cell subsets. T cells were isolated by negative selection with EasySep CD3 enrichment cocktail and magnetic nanoparticles (Stem Cell Technologies, Vancouver, British Columbia, Canada). Briefly, unwanted cells were specifically labeled with bispecific tetrameric antibody complexes that recognize unwanted cells and dextran. Dextran-coated magnetic nanoparticles were added, and magnetically labeled cells were then separated from unlabeled target cells (CD3⁺) with a magnet. Purified T cells were further separated into CCR7⁺ and CCR7^– T cells with a PE selection kit (Stem Cell Technologies). Briefly, T cells were labeled with PE-conjugated anti-CCCR7 antibody. The labeled cells were incubated with bispecific tetrameric antibody complexes that recognize PE-labeled cells and dextran. After a 15-min incubation at room temperature, dextran-coated magnetic nanoparticles were added and magnetically labeled cells (CCR7⁺) were separated from unlabeled cells (CCR7^–) with a magnet.

Cell division. Cell division was measured by labeling with CFSE dye, which is a nonfluorescent, highly membrane-permeable diacetate taken up readily by cells. Once it is inside the cell, intracellular esterases cleave the diacetate groups and the resultant fluorescent moiety, 5,6-carboxyfluorescein succinimidyl ester (CFSE), is retained within the cell. CFSE, which is partitioned in DNA, is divided between daughter cells in each successive division, and the mean fluorescence intensity of the cells decreases with each successive division. Hence, discrete populations can be detected on the basis of decreasing fluorescence intensity, identifying cells that have undergone up to 10 successive divisions. In these studies, MNCs (10⁷/ml) were incubated at 37°C for 15 min with 10 µl of 500 µM CFSE diluted in PBS. In order to stop the reaction, 100 µl of fetal calf serum (FCS) was added to per ml of cells; cells were washed twice in RPMI-FCS medium and then resuspended in RPMI-FCS medium. Labeled cells were cultured in the presence of HHV-6A for 5 days and harvested by centrifugation. The cells were stained with PerCP-anti-CD4 or -anti-CD8, APC-anti-CD45RA, and PE-anti-CCR7 and run on a FACScalibur (Becton Dickinson, San Jose, CA). T_N (CD4⁺/CD8⁺ CD45RA⁺ CCR7⁺), T_CM (CD4⁺/CD8⁺ CD45RA^– CCR7⁺), T_EM (CD4⁺/CD8⁺ CD45RA^– CCR7^–), and T_EMRA (CD4⁺/CD8⁺ CD45RA⁺ CCR7^–) T-cell subsets were gated, and cell division was assessed by the FlowJo software program.

Measurement of cytokines. Cytokine secretion was measured in MNCs and purified CCR7⁺ (T_N plus T_CM) and CCR7^– (T_EM plus T_EMRA) T-cell subsets by enzyme-linked immunosorbent assay (ELISA). Cells were activated by HHV-6A for 2 to 5 days. Supernatant were collected and stored at –20°C until assayed for detection of cytokines by ELISA (with ELISA kits from BD Pharmingen, San Jose, CA). The cytokines interleukin-8 (IL-8), IL-6, TNF-, and gamma interferon (IFN-) were assayed in accordance with BD Pharmingen protocols.

Measurement of apoptosis. Apoptosis was measured by terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling assay. MNCs (1 x 10⁶/ml) were activated with HHV-6A for 5 days. At the end of the incubation period, the cells were washed with PBS and stained with PerCP-conjugated anti-CD4 or -CD8, APC-conjugated anti-CD45RA (BD Biosciences, San Jose, CA), and PE-conjugated anti-CCR7 (R&D Systems, Minneapolis, MN). Cells were washed and fixed in 2% paraformaldehyde for 30 min at room temperature. Cells were permeabilized with sodium citrate buffer containing Triton X-100 for 2 min on ice, washed, and incubated with FITC-dUTP in the presence of TdT enzyme solution for 1 h at 37°C. Following incubation, cells were washed two times with PBS and analyzed with a FACScalibur. Ten thousand cells were acquired, and data were analyzed with CellQuest software.

Caspase activation. The activation of caspase-8, caspase-9, and caspase-3 in four distinct subsets of T cells was analyzed by FACScalibur with carboxyfluorescein-labeled cell-permeable peptide substrates that recognize cleaved caspase-8 (FAM-LETD-FMK), caspase-9 ((FAM-LETD-FMK), and caspase-3 (FAM-DEVD-FMK) according to the protocol provided by the manufacturer. Briefly, 10 µl of appropriately diluted caspase-8, caspase-9, or caspase-3 substrate was added to MNCs that had been cultured with or without HHV-6A and incubated for 1 h at 37°C. Cells were then stained with PerCP-conjugated anti-CD4 or -CD8, APC-conjugated anti-CD45RA, and PE-conjugated anti-CCR-7 MAbs. Cells were washed with wash buffer and analyzed by FACScalibur.

Statistical analysis. All data were analyzed with the paired t test. Cumulative data are expressed as means ± standard deviations. P values of <0.05 were considered significant.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
HHV-6A induces proliferation selectively in effector memory CD4⁺ and CD8⁺ T cells. We have analyzed the in vitro ability of HHV-6A to induce cell division in T_N and various memory subsets of CD4⁺ and CD8⁺ T cells by using CFSE (a DNA dye) and flow cytometry. MNCs loaded with CFSE dye were incubated in the presence or absence of HHV-6A for 5 days, and cell division was analyzed by multicolor flow cytometry. Our data show that HHV-6A induces cell division in effector memory (T_EM and T_EMRA) CD4⁺ (Fig. 1A) and CD8⁺ (Fig. 1B) T cells but not in T_N and T_CM CD4⁺ and CD8⁺ T cells (Fig. 1).

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FIG. 1. HHV-6-induced cell division in TN cells and various memory subsets of CD4+ (A) and CD8+ (B) T cells. MNCs were loaded with CFSE and then stimulated with HHV-6A for 5 days, and cell division (partitioning of CFSE dye) was measured by multicolor flow cytometry with PerCP-conjugated anti-CD4 or anti-CD8 MAbs together with PE-conjugated anti-CCR7 and APC-conjugated anti-CD45RA MAbs or isotype control antibodies. Cell division was restricted to effector memory (TEM and TEMRA) subsets.

HHV-6A induces apoptosis in naïve and central memory CD4⁺ and CD8⁺ T cells. Next, we explored the possibility that a lack of cell division in T_N and T_CM CD4⁺ and CD8⁺ T cells is due to selective or preferential HHV-6A-induced cell death in T_N and T_CM CD4⁺ and CD8⁺ T cells. Therefore, HHV-6A-induced apoptosis was measured in T_N and different memory subsets of CD4⁺ and CD8⁺ T cells with a terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling assay and multicolor flow cytometry. Our data show that HHV-6A induced apoptosis selectively in T_N and T_CM CD4⁺ (Fig. 2A) and CD8⁺ (Fig. 2B) T cells, which is associated with caspase-8, caspase-9, and caspase-3 activation, suggesting an involvement of both death receptor and mitochondrial pathways of apoptosis. In contrast, T_EM and T_EMRA CD4⁺ (Fig. 2) and CD8⁺ (Fig. 3) T cells were relatively resistant to HHV-6A-induced apoptosis and did not display activation of caspase-8, caspase-9, or caspase-3. Therefore, failure of HHV-6A to induce cell division in T_N and T_CM CD4⁺ and CD8⁺ T cells appears to be due to selective induction of apoptosis.

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FIG. 2. HHV-6A-induced apoptosis in TN cells and various memory subsets of CD4+ (A) and CD8+ (B) T cells. MNCs were activated with HHV-6A for 5 days, and apoptosis and caspase activation were measured by multicolor flow cytometry. Apoptosis and caspase-8, caspase-9, and caspase-3 activation were observed predominantly in the TN and TCM subsets.

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FIG. 3. Expression of death receptors on TN cells and various memory subsets of CD4+ and CD8+ T cells following HHV-6A activation. MNCs were stimulated with controls and HHV-6A for 5 days. Cells were washed and stained for various death receptors with specific antibodies and isotype control antibodies. No induction of DR5 and CD95L or upregulation of TNFRs was observed.

Susceptibility of T_N and T_CM CD4⁺ and CD8⁺ T cells to apoptosis is not associated with upregulation of death receptors. There is a controversy regarding mechanisms of apoptosis associated with HHV-6 (19, 20, 38). Since we observed activation of both caspase-8 (death receptor pathway) and caspase-9 (mitochondrial pathway), we examined the expression of TNFR1, TNFR2, DR5, and CD95L on various subsets of CD4⁺ and CD8⁺ T cells following activation with HHV-6A. MNCs were activated in the presence or absence of HHV-6A for 5 days and then examined for the expression of various death receptors by multicolor flow cytometry. Data from three separate experiments with cells from three subjects are shown in Fig. 3. There was no induction of expression of DR5 or CD95L in any of the subpopulations. Furthermore, no significant (P > 0.1) upregulation of TNFR1 or TNFR2 was observed.

Susceptibility to apoptosis is not associated with Bcl-2 expression. Bcl-2 is a prototypical antiapoptotic protein that plays an important role in the regulation of apoptosis via the mitochondrial pathway (9, 31). Therefore, we examined the expression of Bcl-2 on various subsets following HHV-6A activation. MNCs were activated with HHV-6A for 5 days, and Bcl-2 expression was observed on various subsets with a specific Bcl-2 antibody and isotype control antibodies by multicolor flow cytometry. Since the majority of cells express Bcl-2, the change in Bcl-2 density, as measured by mean fluorescence channel numbers, was analyzed. Data in Fig. 4 show that HHV-6A downregulated Bcl-2; however, downregulation of Bcl-2 was observed to similar extents in both apoptosis-sensitive T_N and T_CM and apoptosis-resistant T_EM and T_EMRA CD4⁺ and CD8⁺ T-cell subsets, suggesting that differential sensitivity/resistance to HHV-6A-induced apoptosis is independent of Bcl-2.

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FIG. 4. Expression of Bcl-2 on TN cells and various memory subsets of CD4+ and CD8+ T cells following HHV-6A activation. MNCs were cultured with controls and HHV-6A for 5 days and then washed, stained with anti-Bcl-2 MAb and isotype control antibodies, and analyzed by multicolor flow cytometry. No correlation was observed between apoptosis and the expression of Bcl-2. Data are expressed as means ± standard deviations from three separate experiments.

Susceptibility to apoptosis is not associated with changes in CD46. Since CD46 is a cellular receptor for HHV-6 (23, 26) and HHV-6A induces fusion from outside through CD46 (32), we determined whether the differential sensitivity of T_N cells and different subsets of memory CD4⁺ and CD8⁺ T cells to HHV-6A-induced apoptosis was due to the differential expression of CD46. CD46 expression was measured by multicolor flow cytometry with a cocktail of MAbs against CD46 and cell surface antigens that define T_N cells and various subsets of memory CD4⁺ and CD8⁺T cells. Figure 5 shows that the proportions of T_N cells and various subsets of memory CD4⁺ and CD8⁺T cells expressing CD46, as well as the density of CD46 (as measured by mean fluorescence channel numbers), were similar among four subsets of CD4⁺ and CD8⁺ T cells. Therefore, selective induction of cell division or induction of apoptosis in naïve T cells and among various memory subsets is independent of CD46 expression.

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FIG. 5. Expression of CD46 on TN cells and various memory subsets of CD4+ and CD8+ T cells following HHV-6A activation. MNCs were stimulated with controls and HHV-6A for 5 days. Cells were washed and stained with anti-CD46 MAbs and isotype control antibodies. Ten thousand cells were acquired and analyzed by multicolor flow cytometry. No correlation between apoptosis and CD46 expression was observed. MFC#, mean fluorescence channel number.

HHV-6A induces cytokine production. We examined the induction of cytokine production by HHV-6A. Peripheral blood MNCs were stimulated with control or HHV-6A for 48 h, and supernatants were examined for cytokine production by ELISA. Data in Fig. 6 show that HHV-6A induces IL-6, TNF-, IL-8, and IFN- secretion; however, no secretion of IL-4, IL-5, or IL-10 was observed (data not shown).

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FIG. 6. HHV-6A-induced cytokine production. MNCs were cultured with controls (Cont) and HHV-6A for 2 to 5 days, and culture supernatants were collected and analyzed by ELISA. HHV-6A induced significant amounts of IL-6, IL-8, TNF-, and IFN-.

Next, we determined whether all or some of these cytokines are produced by a selective memory subset. First, T cells were purified, which was followed by purification of CCR7⁺ (T_N plus T_CM) and CCR7^– (T_EM plus T_EMRA) cells. TNF- was almost exclusively produced by CCR7⁺ T cells (T_N plus T_CM), whereas other cytokines were produced by both CCR7⁺ and CCR7^– T cells (Fig. 7). Since CCR7⁺ T cells are sensitive to HHV-6A-induced apoptosis, it is likely that cytokines are produced early in the response to HHV-6A by T_N and T_CM CD4⁺ T cells prior to undergoing apoptosis.

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FIG. 7. HHV-6A-induced cytokine production by CCR7+ and CCR7– T cells. Purified T cells were further separated into CCR7+ (TN plus TCM) and CCR7– (TEM plus TEMRA) T cells. These subpopulations were stimulated with controls and HHV-6A, and supernatants were analyzed for cytokine production by ELISA. IL-6, IL-8, and IFN- were produced by both subsets. However, TNF- was predominantly produced by CCR7+ T cells.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
HHV-6 is a member of the Betaherpetovirinae subfamily. Two major subgroups with distinct genetic, immunological, and virologic characteristics have been named HHV-6A and HHV-6B (2, 6). HHV-6B is associated with a number of human diseases; however, pathological consequences of HHV-6A are less well defined. Since HHV-6 has preferential tropism for CD4⁺ T cells, studies have been limited to CD4⁺ T cells and those too have not been extensively studied.

Recently, memory CD4⁺ and CD8⁺ T-cell subsets have been further subdivided into central and effector memory cells, which display differential homing characteristics and differential proliferative capacities and effector functions (12, 14, 22, 25, 28, 30, 32, 33, 35, 40). These subpopulations are identified by a set of cell surface antigens, which allows their functional characterization (14, 25, 30). In this report, we show that HHV-6A induces cell division primarily in T_EM and T_EMRA CD4⁺ and CD8⁺ T cells, which otherwise display characteristics of immunosenescence, including slow replicative capacity. HHV-6 infection is associated with immunosuppression, and HHV-6-infected patients display decreased T-cell proliferation (10, 27, 34). This is consistent with our observation of a lack of proliferation of T_N and T_CM cells, which constitute major subsets of CD4⁺ and CD8⁺ T cells, which have major proliferative potential (28), and proliferation of T_EM and T_EMRA cells, which constitute only a minor subpopulation of CD4⁺ and CD8⁺ T cells. Next, we investigated the mechanisms of the differential effect of HHV-6A on cell division among T_N cells and various subsets of memory CD4⁺ and CD8⁺T cells.

HHV-6A has been reported to induce apoptosis both in vivo and in vitro in CD4⁺ T cells (19, 20, 38). We have reported that T_N and T_CM CD4⁺ and CD8⁺ cells are susceptible to death receptor (13, 15, 16, 18) and mitochondrial signaling (13, 17) for apoptosis, whereas T_EM and T_EMRA CD4⁺ and CD8⁺ T cells are resistant to apoptosis. In the present study, we show that HHV-6A also induces apoptosis in T_N and T_CM CD4⁺ and CD8⁺ T cells and that T_EM and T_EMRA CD4⁺ and CD8⁺ T cells are resistant to apoptosis, which could explain the lack of HHV-6A-induced cell division in T_N and T_CM CD4⁺ and CD8⁺ cells. Ichimi et al. (19) reported that in cord blood lymphocytes (which contain predominantly T_N cells), HHV-6A-induces apoptosis by an unknown mechanism which is independent of death receptors. In contrast, Inoue et al. (20) reported that both TNF- and anti-CD95 and anti-CD95L antibodies augment HHV-6A-induced apoptosis, therefore suggesting an involvement of death receptors in HHV-6A-induced apoptosis. The difference between these reports may be due to the different cell populations and different culture conditions used. These investigators did not study naïve T cells and various memory subsets, or CD8⁺ T cells, and no activation of caspases was studied. In our study, susceptibility to HHV-6A-induced apoptosis in T_N and T_CM CD4⁺ and CD8⁺ T cells was associated with activation of caspase-8, caspase-9, and caspase-3. Caspase-3 is a common effector of both death receptor and the mitochondrial signaling pathways. Caspase-8 is activated by the death receptor signaling pathway, whereas caspase-9 is activated during mitochondrial signaling for apoptosis (13). However, in certain cell types where the caspase-8 level is low, caspase-9 may be activated by death receptor signaling as an apoptosis-amplifying mechanism (39). Therefore, HHV-6A-induced apoptosis could be via death receptor signaling or via both death receptor signaling and the mitochondrial pathways of apoptosis.

To further investigate apoptotic signaling pathways, we examined the effect of HHV-6A on the expression of several death receptors and the expression of antiapoptotic Bcl-2. We did not observe any significant or differential effect of HHV-6A on the expression of TNFR1, TNFR2, DR5, or CD95L among various subsets of CD4⁺ and CD8⁺ T cells. However, an involvement of downstream adapter molecules or regulatory molecules in death receptor signaling pathways cannot be excluded. HHV-6A downregulated the expression of Bcl-2; however, downregulation was similar among all four subsets of CD4⁺ and CD8⁺ T cells, suggesting that Bcl-2 is unlikely to be involved in the differential sensitivity of T_N and T_CM cell subsets to HHV-6A-induced apoptosis. It is possible; however, other antiapoptotic molecules like Bcl-x_L or proapoptotic molecules like Bax, Bak, or Bid (31) are involved.

Since CD46 serves as a receptor for HHV-6 (23, 26), we explored the possibility that the differential expression of CD46 among T_N cells and various memory subsets of CD4⁺ and CD8⁺ T cells could explain their differential proliferative responses and sensitivities to apoptosis. However, we did not find any difference in the proportions of T_N cells and various memory subsets of CD4 and CD8⁺ T cells expressing CD46 or in the density of CD46. HHV-6 primarily infects CD4⁺ T cells; however, both CD4⁺ and CD8⁺ T cells express CD46, which suggests that infection with HHV-6 requires another coreceptor, which may be lacking in CD8⁺ T cells. HHV-6 has been shown to induce CD4 transcript in CD8⁺ T cells (23). Some CD4⁺ T cells have also been reported to be resistant to HHV-6 infection (23). Our data also suggest that HHV-6 binding to CD46 is sufficient to induce a proliferative response in T_EM and T_EMRA CD4⁺ and CD8⁺ T cells and apoptosis in T_N and T_CM CD4⁺ and CD8⁺ T cells. Furthermore, HHV-6A infection does not appear to be responsible for apoptosis since less that 1% of memory CD4⁺ T cells were infected, as observed with a MAb against HHV-6 core protein (data not shown), and CD8⁺ T cells are not infected with HHV-6A. Why HHV-6A induces apoptosis in T_N and T_CM CD4⁺ and CD8⁺ T cells and proliferation in T_EM and T_EMRA CD4⁺ and CD8⁺ T cells remains unclear.

HHV-6 regulation of cytokine secretion has been studied in HHV-6 infection in vivo (5) and in vitro (10, 27). In vitro, HHV-6 has been shown to suppress IL-2 secretion by MNCs (6) and IL-12 secretion by dendritic cells (27). In contrast, Flamand et al. (11) reported HHV-6-induced induction of IL-1β and TNF-, but not IL-6, in MNCs. In the present study, however, we observed HHV-6A induction of IL-1β, TNF-, IL-6, IFN-, and IL-8 secretion by MNCs. Furthermore, with the exception of TNF-, all of these cytokines are produced by T_N cells and all of the memory subsets of T cells, which are produced primarily by CCR7⁺ T cells (containing T_N and T_CM cells). It also appears that cytokines are produced before (within the first 48 h) T_N and T_CM CD4⁺ cells undergo apoptosis (5 days). Our observations of cytokine production by MNCs are in agreement with those of Arena et al. (4), who reported that HHV-6 induces the production of IL-1β, TNF-, IL-6, and IFN-.

In summary, HHV-6A induces cell division almost exclusively in T_EM and T_EMRA CD4⁺ and CD8⁺ T cells; in contrast, HHV-6A induces apoptosis selectively in T_N and T_CM CD4⁺ and CD8⁺ T cells. This suggests that apoptosis may play a role in poor cell division or lack thereof in T_N and T_CM CD4⁺ and CD8⁺ T cells in response to HHV-6A. It also appears that the secretion of TNF- by T_N plus T_CM CD4⁺ and CD8⁺ T cells and secretion of IL-6, IL-8, and IFN- by T_N cells and all memory subsets occur prior to induction of apoptosis. Thus, HHV-6A differentially regulates the immune responses of naïve T cells and various memory subsets of CD4⁺ and CD8⁺ T cells.

 
This work was in part supported by a grant from the HHV-6 Foundation, Santa Barbara, CA.

 
* Corresponding author. Mailing address: Medical Sciences I, C-240, University of California, Irvine, CA 92697. Phone: (949) 824-5818. Fax: (949) 824-4362. E-mail: sgupta{at}uci.edu

Published ahead of print on 18 March 2009.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

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Journal of Virology, June 2009, p. 5442-5450, Vol. 83, No. 11
0022-538X/09/$08.00+0     doi:10.1128/JVI.00106-09
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

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