Human virus-specific T cells in peripheral blood and lymph nodes: Phenotype, function and clonal relationships - Chapter 7: Emergence of a CD28⁻granzyme B+CD4+ cytomegalovirus-specific T cell subset after recovery of primary cytomegalovirus

(1)

UvA-DARE is a service provided by the library of the University of Amsterdam (https://dare.uva.nl)

UvA-DARE (Digital Academic Repository)

Human virus-specific T cells in peripheral blood and lymph nodes: Phenotype,

function and clonal relationships

Remmerswaal, E.B.M.

Publication date

2014

Document Version

Final published version

Link to publication

Citation for published version (APA):

Remmerswaal, E. B. M. (2014). Human virus-specific T cells in peripheral blood and lymph

nodes: Phenotype, function and clonal relationships.

General rights

It is not permitted to download or to forward/distribute the text or part of it without the consent of the author(s)

and/or copyright holder(s), other than for strictly personal, individual use, unless the work is under an open

content license (like Creative Commons).

Disclaimer/Complaints regulations

If you believe that digital publication of certain material infringes any of your rights or (privacy) interests, please

let the Library know, stating your reasons. In case of a legitimate complaint, the Library will make the material

inaccessible and/or remove it from the website. Please Ask the Library: https://uba.uva.nl/en/contact, or a letter

to: Library of the University of Amsterdam, Secretariat, Singel 425, 1012 WP Amsterdam, The Netherlands. You

will be contacted as soon as possible.

(2)

(3)

EMERGENCE OF A CD28

⁻

_{GRANZYME B}

+

_CD4

+

CYTOMEGALOVIRUS-SPECIFIC T CELL

SUBSET AFTER RECOVERY OF PRIMARY

CYTOMEGALOVIRUS INFECTION

Ester M. M. van Leeuwen

1,2

_{, Ester B. M. Remmerswaal}

2

_,

Mireille T. M. Vossen

3

_{, Ajda T. Rowshani}

1

_,

Pauline M. E. Wertheim-van Dillen

4

_{, René A. W. van Lier}

2

and Ineke J. M. ten Berge

1

1_{Department of Internal Medicine, Divisions of Nephrology}

and Clinical Immunology and Rheumatology,

2_{Laboratory for Experimental Immunology,} 3_{Emma Children’s Hospital, and}

4_{Department of Virology, Academic Medical Center, Amsterdam, The Netherlands}

(4)

CD28

⁻

CD4

+

T CELLS ARE INDUCED BY HCMV

7 ABSTRACT

Cytotoxic CD28

⁻

CD4+_{T cells form a rare subset in human peripheral blood. The}

presence of CD28

⁻

CD4+_{cells has been associated with chronic viral infections, but}

how these particular cells are generated is unknown. In this study, we show that in primary human CMV (hCMV) infections, CD28

⁻

CD4+_{T cells emerge just after cessation}

of the viral load, indicating that infection with hCMV triggers the formation of CD28

⁻

CD4+_{T cells. In line with this, we found these cells only in hCMV-infected persons.}

CD28

⁻

CD4+_{cells had an Ag-primed phenotype and expressed the cytolytic molecules}

granzyme B and perforin. Importantly, CD28

⁻

CD4+_{cells were to a large extent}

hCMV-specific because proliferation was only induced by hCMV-Ag, but not by recall Ags such as purified protein derivative or tetanus toxoid. CD28

⁻

CD4+_{cells only produced}

IFN-

γ

after stimulation with hCMV-Ag, whereas CD28+_CD4+_{cells also produced IFN-}

γ

in response to varicella-zoster virus and purified protein derivative. Thus, CD28

⁻

CD4+

(5)

CD28

⁻

CD4

+

7 INTRODUCTION

Within the circulating human CD4+_{T cell population, a subset of cytotoxic cells has}

recently been described (1). These cells express the cytolytic molecules perforin and granzyme B (2) and have no expression of the costimulatory molecules CD28 and CD27 or the chemokine receptor CCR7. CD28

⁻

CD4+_{T cells have been reported to}

be expanded in patients with rheumatoid arthritis (RA), especially in those with extra-articular inflammatory lesions and rheumatoid vasculitis (2). CD28

⁻

CD4+_{T cells in RA}

have a limited TCR diversity, suggesting that they recognize only a few Ags (3, 4). Additionally, in patients with unstable angina, high numbers of CD28

⁻

CD4+_IFN-

γ

-producing cells have been described, which were able to effectively lyse endothelial cells and thereby possibly contribute to plaque destabilization (5, 6). Finally, an increased percentage of CD28

⁻

CD4+_{T cells was found in the circulation of elderly}

individuals, but exceptions indicated that other factors besides age may be involved in the generation of these cells (7, 8).

The origin and specificity of CD28

⁻

CD4+_{T cells is unknown. Because the percentage}

of CD28

⁻

CD4+_{cells varied among individuals but was generally higher in HIV-infected}

individuals, Appay et al. (1) suggest that the presence of these cells is related to chronic viral infections. In patients with RA and in the elderly, the expansion of CD28

⁻

CD4+_and

CD28

⁻

CD8+_{T cells has been described to be associated with hCMV infection (9-11).}

HCMV is a widespread member of the

β

-herpesvirus family that persists in the host in a latent state after primary infection. In healthy individuals, virus and host exist in a symbiotic equilibrium, such that infectious disease manifestations are hardly encountered. However, when the immune system is compromised, for example in HIV-infected individuals or in transplant recipients, hCMV infection can lead to a number of disease symptoms (12). Although CD8+_{T cells are believed to be most important in controlling}

hCMV infection, CD4+_{T cells also play a role in the defense. Previously, we showed that}

during primary hCMV infection, virus-specific CD4+_{T cells precede the appearance of}

both specific Abs and virus-specific CD8+_{T cells in renal transplant recipients (13, 14).}

In symptomatic primary hCMV infection, hCMV-specific IFN-

γ

-producing CD4+_{T cells}

were delayed and only appeared after antiviral therapy, suggesting that CD4+_{T cells are}

indispensable in protection against hCMV disease (13).

HCMV infection exerts a profound effect on the CD8+_{T cell pool that persists}

long after primary infection (9, 15-17). In hCMV carriers, increased percentages and absolute numbers of circulating cytolytic CD45RA+_CD27

⁻

_CD8+_{T cells have been}

detected that were not observed after EBV or varicella-zoster virus (VZV) infection nor after vaccination with measles-mumps-rubella (18).

The fact that hCMV infection leaves a fingerprint within the CD8+_{T cell compartment}

together with the observations that CD28

⁻

CD4+_{T cells are predominantly found}

in hCMV-infected individuals prompted us to analyze emergence and specificity of CD28

⁻

CD4+_{T cells in primary hCMV infection and during latency.}

(6)

CD28

⁻

CD4

+

7 MATERIALS AND METHODS

Subjects

Healthy hCMV-seronegative (n = 13) and -seropositive (n = 15) healthy volunteers as well as hCMV-seronegative (n = 7) and -seropositive (n = 26) renal transplant recipients (at least 1 year after transplantation) were included in this study. The renal transplant patients were treated with basic immunosuppressive therapy consisting of prednisolone, mycophenolate mofetil, and cyclosporine. In addition, we longitudinally studied four renal transplant recipients who were hCMV-seronegative before transplantation and who received a kidney from a hCMV-seropositive donor (14). All patients gave written informed consent, and the study was approved by the local medical ethics committee.

PBMCs

Heparinized peripheral blood samples were collected, and PBMCs were isolated using standard density gradient centrifugation techniques and were subsequently cryopreserved until the day of analysis.

HCMV-PCR, anti-hCMV IgG, and anti-EBV IgG

Quantitative PCR was performed in EDTA whole blood samples as described before (19). To determine hCMV serostatus, anti-hCMV IgG was measured in serum using the AxSYM microparticle enzyme immunoassay (Abbott Laboratories, Abbott Park, IL) according to the manufacturer’s instructions. Measurements were calibrated relative to a standard serum. The EBV serostatus was investigated by determination of IgG to both EBV-viral capsid Ag and Epstein-Barr nuclear Ag by ELISA (Biotest, Dreieich, Germany).

Immunofluorescent staining and flow cytometry

PBMCs were washed in PBS containing 0.01% (w/v) NaN3 and 0.5% (w/v) BSA. A total of 200,000 PBMCs were incubated with fluorescent-labeled conjugated mAbs (concentrations according to manufacturer’s instructions) for 30 min at 4°C. For analysis of expression of surface markers, the following mAbs were used in different combinations: CD4 PerCP, CD4 PerCP Cy5.5, CD4 APC, CD25 PE, CD27 PE, CD28 PE, CD45RA FITC, CD45R0 PE, CD49d PE, CD57 FITC, CD69 FITC, HLA-DR FITC, anti-TCR

γδ

FITC, and streptavidin-APC (all from BD Biosciences, San Jose, CA); CD4 APC, CD45RA biotin, and CCR7 PE, (all from BD Pharmingen, San Diego, CA); CD11a FITC, CD11b FITC, CD11c R-PE, and CD18 FITC (all from DAKO, Glostrup, Denmark); CD27 biotin, CD28 FITC, CD49f FITC, and CD54 unlabeled (all from Sanquin, Amsterdam, The Netherlands); CXCR3 (R&D Sytems, Minneapolis, MN); CD69 APC (Caltag Laboratories, Burlingame, CA); CD49e FITC (Chemicon Europe, Chandlers Ford, U.K.); and TCR

αβ

(7)

CD28

⁻

CD4

+

7

FITC (Instruchemie, Delfzijl, The Netherlands). To stain unlabeled monoclonals, goat anti-mouse Ig R-PE (Southern Biotechnology Associates, Birmingham, AL) was used and for blocking normal mouse serum (Sanquin) was added before the rest of the staining was performed. Stainings for chemoattractant receptor of Th2 cells (CRTh2) PE and CCR5 APC (both from BD Biosciences) were performed on whole blood at room temperature followed by lysis of red cells with lysing solution (BD Biosciences). Cells were washed in PBS containing 0.01% (w/v) NaN₃ and 0.5% (w/v) BSA and were analyzed using a FACSCaliber flow cytometer and CellQuest software (BD Biosciences).

Intracellular granzyme B, perforin, and cytokine staining

For intracellular staining, cells were fixed in FACS lysing solution (BD Biosciences) and permeabilized (BD Biosciences). Cells were then incubated with anti-granzyme B PE (Sanquin) or anti-perforin FITC (BD Biosciences) mAbs, according to manufacturer’s instructions. Flow cytometric analysis was performed thereafter. For cytokine stainings, cells were first stimulated for 6 h at 37°C with hCMV-Ag (inactivated whole virus, 10 μl/ ml; Microbix Biosystems, Toronto, Canada), VZV-Ag (20 μl/ml; Microbix Biosystems), purified protein derivative (PPD; 11.8 μg/ml; Satens Serum Institut, Copenhagen, Denmark), or tetanus toxoid (TT; 17.6 Lf/ml; RIVM, Bilthoven, The Netherlands). As positive controls, cells were stimulated with PMA (1 ng/ml)/ionomycin (1 μg/ml) (Sigma-Aldrich, St. Louis, MO) or Staphylococcus aureus enterotoxin B (SEB; 2 μg/ml; ICN/Fluka, Buchs, Switzerland). All stimulations were performed in a final volume of 1 ml of RPMI 1640 (Life Technologies, Rockville, MD) containing 10% heat-inactivated FCS and 1 μg/ml (final concentration) VLA-4 mAb (CD49d; BD Biosciences). For the final 5 h of culture, brefeldin A (Sigma-Aldrich) was added to the culture in a final concentration of 10 μg/ml. After culture, the same intracellular staining procedure described above was performed using anti-IFN-

γ

FITC, anti-TNF-

α

FITC, anti-IL-2 FITC, and anti-IL-4 PE (all from BD Biosciences).

CFSE labeling

PBMCs were resuspended in PBS at a final concentration of 5–10 × 106_{cells/ml. PBMCs}

were labeled with 0.5 μM CFSE (final concentration; Molecular Probes, Eugene, OR) in PBS for 8–10 min, shaking at 37°C. Cells were washed and subsequently resuspended in IMDM supplemented with 10% human pool serum (BioWhittaker, Verviers, Belgium), antibiotics, and 3.57 × 10−4_{% (v/v) 2-ME (Merck, West Point, PA) (culture medium).}

Proliferation assays

PBMCs from hCMV-seropositive donors were sorted by FACSAria (BD Biosciences) into a CD28

⁻

CD4+_{population, a CD28}+_CD45RA

⁻

_CD4+_{population, and a}

(8)

CD28

⁻

CD4

+

7

and left in culture medium for 4 days. Cells were cultured in the presence of autologous PBMCs that were, before irradiation, cultured for 5 h with hCMV-Ag, PPD, or TT or in just medium. All stimulations were performed in both the absence and presence of IL-2 (50 U/ml; Biotest). Control stimulations consisted of medium only, IL-2 only, autologous irradiated PBMCs only, or autologous irradiated PBMCs with IL-2. As a positive control, cells were stimulated with PHA (Life Technologies) for 3 days.

Amplification of CDR3 regions

cDNA was synthesized from RNA from equal amounts of sorted CD28

⁻

CD4+_and

CD28+_CD4+_{cells. PCR was performed as described previously (20, 21) in single TCRBV}

PCRs with a TCRBC primer labeled with a fluorescent dye (Life Technologies; CAG GCA CAC CAG TGT GGC-FAM). CDR3 size distributions were visualized with the ABI Prism 3100 Genetic Analyzer (Applied Biosystems, Foster City, CA).

Statistical Analysis

The two-tailed Mann-Whitney U test was used for analysis of differences between groups. Statistical significance was indicated by p values < 0.05.

RESULTS

CD28

⁻

CD4

+

_{granzyme B-expressing cells appear}

in peripheral blood after primary hCMV infection

Because the presence of cytotoxic CD28

⁻

CD4+_{T cells in the circulation seems to be}

associated with chronic viral infections (1) and because prior hCMV infection leaves a fingerprint in the CD8+_{T cell pool (9, 15-18), we investigated the involvement of}

CD28

⁻

CD4+_{cells in hCMV infection. Therefore, we longitudinally studied}

hCMV-seronegative recipients of a hCMV-seropositive kidney graft who experienced a primary hCMV infection. As described before, the first sign of specific immunity to hCMV is the appearance of IFN-

γ

-producing CD4+_{T cells in the circulation around the}

peak of the viral load, followed by specific Abs and hCMV-specific CD8+_{T cells (13, 14).}

Interestingly, we found that only after cessation of the viral load an increase was observed in CD28

⁻

CD4+_{T cells expressing the cytolytic molecule granzyme B (figure 1A). The}

percentage of CD28

⁻

CD4+_{granzyme B}+_{T cells continued to increase long after the}

viral load became undetectable (figure 1B). The increase in CD28

⁻

CD4+_{granzyme B}+

T cells during primary hCMV infection was not seen in renal transplant recipients who remained seronegative for hCMV, excluding that either the allograft itself or initiation of immunosuppressive drugs caused the appearance of cytotoxic CD4+_{T cells (data}

not shown). The presence of CD4+_CD28

⁻

_{granzyme B}+_{cells was also not affected by}

(9)

CD28

⁻

CD4

+

7

wk 11 A wk 46 gr anz y m e B CD28 pre Tx wk 18 wk 15 wk 01 B % CD2 8 -CD4 +gr anz y m e B +c e lls %_I F N -γ pr oduc ing C D 4 + c e lls

weeks after transplantation

hC M V l oad hCMV viral load IFNγ+_CD4+_{T cells} CD28⁻CD4+_{T cells}

FIGURE 1. CD28⁻granzyme B+_CD4+_{T cells appear in the peripheral blood after primary hCMV}

infection. A, Dot plots gated on total CD4+_{T cells show the appearance of CD28}⁻_{granzyme B}+_cells

after cessation of the viral load in a primary hCMV infection. The numbers in the indicated quadrants represent the percentage within total CD4+_{T cells. Pre Tx indicates pretransplantation timepoint; wk}

01, 11, 15, 18, and 46 indicate weeks after transplantation. Representative flow cytometric analysis of one of four patients is shown. B, The upper graph shows the kinetics of the viral load (hCMV DNA, copies/ml; light gray) during primary hCMV infection. The lower graph shows the percentages of CD28⁻granzyme B+_{cells (left y-axis;}_●_{) and IFN-}γ_{-producing cells upon hCMV stimulation (right}

(10)

CD28

⁻

CD4

+

7

that hCMV infection is the key factor that causes the large increase in the percentage of cytotoxic CD28

⁻

CD4+_{granzyme B}+_{T cells. CD28}

⁻

_CD4+_{granzyme B}+_{T cells will be}

referred to as CD28

⁻

CD4+_{cells in the following paragraphs.}

The percentage of circulating CD4

+

_CD28

⁻

_{cells is highly}

increased in CMV-seropositive individuals

We next investigated whether the observed change in the composition of the CD4+_T

cell compartment lasted during hCMV latency. When comparing hCMV-seropositive and hCMV-seronegative renal transplant recipients, no clear differences were observed in the distribution of CD4+_{T cell subsets defined by CD27 and CD45RA, CCR7}

and CD45RA, or CD45R0 and CD45RA (data not shown). However, only in hCMV-seropositive patients a clearly distinguishable population of CD28

⁻

CD27

⁻

CD4+_cells

was found (figure 2). A population of cells lacking only CD27 expression was found in both groups (figure 2). The population of CD4+_{T cells expressing granzyme B was}

also larger in hCMV-seropositive individuals, supporting the association between lack of CD28-expression and the expression of cytolytic molecules (Figure 2).

Also, in cohorts of both renal transplant recipients and healthy individuals the percentage of CD28

⁻

CD27

⁻

cells within CD4+_{T cells was significantly higher in}

hCMV-seropositive individuals (figure 3A). Whereas the percentage of CD28

⁻

CD27

⁻

CD4+_{T cells}

in hCMV-seronegative individuals was always below 0.5%, these percentages ranged in hCMV-seropositive healthy individuals from 0.7% to 6.2% and in renal transplant recipients

granzyme B CD27 hCMV -hCMV + CD2 8 CD4

FIGURE 2. During latency, CD4+_{T cells in hCMV-seropositive individuals are CD28}⁻_CD27⁻_{and express}

granzyme B. Representative flow cytometric analysis is shown of one of five hCMV-seronegative and five hCMV-seropositive renal transplant recipients, respectively (at least 1 year after transplantation). Left plots are gated on total CD4+_{T cells; right plots are gated on total lymphocytes. The numbers}

represent the quadrant percentages of cells within CD4+_{T cells. “hCMV −” indicates a}

(11)

CD28

⁻

CD4

+

7

from 0.9% to 61.4% (figure 3A). The presence of CD28

⁻

T cells has been associated with age (7). In our study, however, the mean ages of the seronegative and hCMV-seropositive groups did not differ (32.8 vs 32.5 for the healthy individuals and 47.6 vs 48.7 for the renal transplant patients, respectively; NS). Despite a large variation in percentages of CD28

⁻

CD27

⁻

CD4+_{T cells in hCMV-seropositive renal transplant recipients, the median}

was significantly higher than in seropositive healthy individuals (p = 0.0006; Figure 3A). This is not due to the difference in age because no relation was found between the percentage of CD28

⁻

CD27

⁻

CD4+_{cells and age (data not shown).}

To investigate a possible effect of other persistent viruses such as EBV on the presence of CD28

⁻

CD27

⁻

CD4+_{cells, four groups were discerned in the healthy}

individuals tested, based on EBV and hCMV serostatus. As shown in figure 3B, the percentage of CD28

⁻

CD27

⁻

CD4+_{T cells was significantly higher in peripheral blood}

of hCMV-seropositive individuals, independent of their EBV serostatus. This indicates that, as described for CD8+_{T cells (18), hCMV, and not other viruses, causes an increase}

in the percentage of circulating cytotoxic CD4+_{T cells.}

CD4

+

_CD28

⁻

_{T cells have the phenotype of cytotoxic}

Ag-experienced cells but are not recently activated

To gain more insight into the role of CD28

⁻

CD4+_{T cells in hCMV infection, we further}

analyzed the phenotype of these cells using different surface markers to classify T cells. As shown in figure 4A, CD28

⁻

CD4+_{T cells may be classified as effector memory type cells:}

they did not express the costimulatory receptor CD27 but uniformly expressed CD57, CD45R0, and not CD45RA. CD28

⁻

CD4+_{cells expressed the cytolytic molecules granzyme}

B and perforin (figure 4A), suggesting cytotoxic potential (1). Furthermore, CD28

⁻

CD4+_T

cells expressed LFA-1

α

- and

β

-chain (CD11a and CD18), macrophage adhesion molecule 1 (CD11b), ICAM-1 (CD54), and VLA 4–6 (CD49d–f) and did not express CD11c (data not shown). All CD4+_CD28

⁻

_{T cells were TCR}

αβ

_{-positive and did not express TCR}

γδ

_(data

not shown). CD4+_CD28

⁻

_{T cells appeared not to be recently activated because CD69,}

CD25, CD38, and HLA-DR were not expressed (Figure 4B). Concerning the expression of chemokine receptors, CD28

⁻

CD4+_{T cells did not express CCR7, which again shows that}

these cells have a memory phenotype (Figure 4C). CRTh2 was not expressed, whereas most cells were CCR5+_{, indicating a Th1 phenotype. Remarkably, the expression of the}

inducible chemokine receptor CXCR3 on CD28

⁻

CD4+_{T cells was highly variable (0–100%)}

among different donors, as shown for three donors in Figure 4C. The data from the phenotypic analysis of CD28

⁻

CD4+_{T cells are summarized in Table I.}

To investigate the clonality of CD28

⁻

CD4+_{cells, the TCR V}

β

_{repertoire of sorted}

CD28

⁻

CD4+_{T cells was determined and compared with that of CD28}+_CD4+_{T cells. The}

CD28

⁻

CD4+_{T cell subset showed a skewed distribution of V}

β

_{subfamilies compared}

with CD28+_CD4+_{T cells (data not shown), indicating that CD28}

⁻

_CD4+_{T cells had}

(12)

CD28

⁻

CD4

+

7

B 0 2 4 6 8 hCMV / EBV status % CD2 8 -CD2 7 -w it h in CD4 +

- / -

_{- / +}

_{+ / -}

_{+ / +}

P=0.0016 P=0.0002 0 5 10 10 20 30 40 50 60 70 hCMV + RTx P<0.0001 P=0.0006 P<0.0001 % CD2 8 -CD2 7 -w it h in CD4 + healthy hCMV - hCMV - hCMV + A

FIGURE 3. Percentages of CD28⁻CD27⁻CD4+_{T cells are significantly higher in hCMV-seropositive}

individuals. Percentages of CD28⁻CD27⁻ cells within total CD4+_{T cells are shown. A, “Healthy”}

indicates healthy individuals, and “RTx” indicates renal transplant recipients at least 1 year after transplantation. “hCMV −” indicates hCMV-seronegative individuals (n = 13 healthy and n = 7 RTx individuals), and “hCMV +” indicates hCMV-seropositive individuals (n = 15 healthy and n = 26 RTx individuals). B, Healthy individuals were divided into four groups according to hCMV and EBV serostatus. “−” indicates seronegative and “+” indicates seropositive for hCMV and/or EBV. n = 7 hCMV −/EBV −, n = 6 hCMV −/EBV +, n = 2 hCMV +/EBV −, and n = 13 hCMV +/EBV +.

CD28

⁻

CD4

+

_{T cells proliferate and produce cytokines}

upon hCMV stimulation

To test whether CD28

⁻

CD4+_{T cells are hCMV-specific, CD4}+_{T cells from a}

(13)

CD28

⁻

CD4

+

7

populations and were stimulated with hCMV Ag, PPD, or TT in the absence or presence of IL-2. Stimulation with IL-2 alone or with irradiated autologous PBMCs alone did not induce proliferation of the sorted cell populations (figure 5). After hCMV Ag stimulation, CD28

⁻

CD4+_{cells proliferated, and this was enhanced by addition of IL-2 (figure 5).}

Upon each division, CD28 was up-regulated on CD28

⁻

CD4+_{cells. Stimulation with}

PPD and TT did not induce proliferation of CD28

⁻

CD4+_{cells, not even when IL-2 was}

added. CD28+_CD45RA

⁻

_{cells from the same donor did proliferate upon stimulation with}

hCMV and PPD. When IL-2 was added, CD28+_CD45RA

⁻

_{cells proliferated in response}

to irradiated autologous PBMCs plus IL-2 without Ag (26), but this was enhanced when hCMV, PPD, or TT was added. CD28+_CD45RA+_{naive cells did not proliferate under}

any stimulatory condition (figure 5). As described before (27), CD28

⁻

CD4+_{, but not}

CD28+_CD4+_{cells, proliferated poorly upon PHA stimulation (data not shown).}

HCMV specificity was corroborated by Ag-induced

cytokine production analysis.

After stimulation of PBMCs from a hCMV-seropositive donor with hCMV Ag, only CD28

⁻

CD4+_{T cells produced IFN-}

γ

_{, whereas upon PMA/ionomycin or SEB stimulation, both}

CD28

⁻

and CD28+_CD4+_{T cells produced this cytokine (figure 6A and data not shown).}

The percentage of cytokine-producing cells upon PMA/ionomycin stimulation within CD28+_CD4+_{cells was lower than within CD28}

⁻

_CD4+_{cells, which can be explained by}

the presence of naive CD4+_{cells in this fraction, which are not so efficient in producing}

cytokines (28). To test whether cytokine production was in general restricted to CD28

⁻

CD4+_{cells or whether this was specific for hCMV, we performed the assay with}

PBMCs from a hCMV-seropositive donor who had recently been in contact with a

Table 1: Expression of phenotypic markers on CD4+_CD28⁻_cells

marker Expressed on CD4+_CD28⁻_cells _marker Expressed on CD4+_CD28⁻_cells CD27 - CD11a + CCR7 - CD11b + CD57 + CD11c -CD45R0 + CD18 + CD45RA - CD54 + granzyme B + CD25 -perforin + CD38 -CD49d + CD69 -CD49e + HLA-DR -CD49f + CXCR3 +/

(14)

-CD28

⁻

CD4

+

7 B

CD6 9 HL A -DR CD2 5 CD28 CD3 8 A CD5 7 CD28 CD2 7 CD4 5 R0 C D 45R A per for in gr anz y m e B CCR7

C

CX CR3 CD28 CCR5 CRT h 2 CX CR3 CX CR3

FIGURE 4. CD28⁻CD4+_{T cells have a primed but not recently activated phenotype. All dot plots}

are gated on total CD4+_{T cells. A, Phenotype of CD28}⁻_CD4+_{and CD28}+_CD4+_{cells in relation}

to subset markers CD27, granzyme B, perforin, CD57, CD45R0, and CD45RA. B, Phenotype of CD28⁻CD4+_{and CD28}+_CD4+_{cells in relation to activation markers CD69, CD25, CD38, and}

HLA-DR. C, Phenotype of CD28⁻CD4+_{and CD28}+_CD4+_{cells in relation to chemokine receptors}

CCR7, CRTh2, CCR5, and CXCR3. Representative flowcytometric analysis from four donors is shown; CXCR3 staining is shown from three different donors selected from a total of eight donors.

(15)

CD28

⁻

CD4

+

7

CD28- CD28+ CD45RA-CD28+ CD45RA+ CFSE CD2 8 With IL-2 <1 <1 <1 1 1 <1 <1 <1 2 <1 46 63 70 58 76 med irr PBMC hCMV Ag PPD TT CD28- CD28+ CD45RA-CD28+ CD45RA+ CD2 8 CFSE Without IL-2 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 38 26 58

FIGURE 5. CD28⁻CD4+_{cells proliferate upon hCMV Ag stimulation but do not proliferate upon}

stimulation by PPD or TT. “CD28⁻” are sorted CD28⁻CD4+_{cells, “CD28}+_CD45RA⁻_{” are sorted}

CD28+_CD45RA⁻_CD4+_{cells, and “CD28}+_CD45RA+_{” are sorted CD28}+_CD45RA+_CD4+_{naive cells. Dot}

plots show CFSE profiles of the sorted cell populations after 4 days of stimulation with medium or irradiated autologous PBMCs unloaded or loaded with hCMV Ag, PPD, or TT. The left panel shows stimulations without IL-2 and the right panel shows them with IL-2. For clarity, only 33% of dots are shown from the two CD28+_{populations; numbers indicate percentage of divided cells. Flowcytometric}

analysis is displayed from one hCMV-seropositive donor who was known to show a proliferative response upon PPD stimulation. Experiments in which proliferation of CD28⁻CD4+_{cells upon hCMV}

Ag stimulation was tested have been performed in four different individuals and gave similar results.

child experiencing varicella (chickenpox), in which case it is possible to measure VZV-specific CD4+_{T cells by IFN-}

γ

_{production (29). Only CD28}+_CD4+_{T cells produced}

IFN-

γ

upon VZV stimulation, in contrast with the hCMV Ag-induced IFN-

γ

production by CD28

⁻

CD4+_{cells (figure 6B). In cells from a hCMV-seropositive donor who reacted}

to PPD and TT, IFN-

γ

production after stimulation with hCMV was mostly restricted to CD28

⁻

cells, whereas only CD28+_{cells produced this cytokine upon PPD stimulation}

(figure 6C). Stimulation with TT did not result in any IFN-

γ

production (data not shown). These data indicate that CD28

⁻

CD4+_{are especially hCMV-specific, whereas}

(16)

CD28

⁻

CD4

+

7

IFN-CD6 9 A C B CD28+ CD28⁻

medium hCMV Ag PMA / iono

0.03 80.01 5.58 0.01 21.19 0.07 CD28+ CD28⁻

medium hCMV Ag PPD PMA / iono

0.04 57.18 9.95 0.01 13.83 0.48 0.03 0.15 CD28+ CD28⁻

medium hCMV Ag PMA / iono

0.03 42.57 0.91 0.00 19.71 0.11 0.02 0.14 VZV Ag

γ

FIGURE 6. CD28⁻CD4+_{cells produce IFN-}γ_{upon hCMV Ag stimulation, but not upon stimulation by}

VZV or PPD. All dot plots are gated on CD28+_CD4+_{cells (CD28+) or CD28}⁻_CD4+_{cells (CD28−). Dot}

plots show IFN-γ production vs CD69 expression of CD28+_{or CD28}⁻_CD4+_{T cells after 6-h stimulation}

with medium, hCMV Ag, VZV Ag, or PPD in the presence of VLA-4 or with PMA/ionomycin as a positive control. The number of events shown from the CD28+_CD4+_{cells is adapted to the number}

of events from the CD28⁻CD4+_{cells (except for VZV and PPD). Numbers indicate percentages of}

CD69+_IFN-γ+_{cells (upper right quadrant) within CD28}+_CD4+_{or CD28}⁻_CD4+_{cells. A, Cells from a}

hCMV-seropositive donor. B, Cells from a hCMV-seropositive donor after recent contact with a VZV-infected child. C, Cells from a hCMV-seropositive donor who reacted to PPD stimulation.

(17)

CD28

⁻

CD4

+

7

Concerning the production of other cytokines by CD4+_CD28

⁻

_{cells, we found that}

TNF-

α

, like IFN-

γ

, was produced after stimulation with hCMV, but not with VZV or PPD. IL-4 was not produced at all, whereas only low amounts of IL-2 were produced by CD4+_CD28

⁻

_{cells after stimulation with PMA/ionomycin or SEB (data not shown).}

DISCUSSION

In this study, we show that the percentage of CD28

⁻

CD27

⁻

granzyme B-expressing CD4+_{T cells in the circulation largely increases after primary hCMV infection. Previously,}

we have demonstrated the emergence of hCMV-specific, IFN-

γ

-producing CD4+_{T cells}

shortly after first appearance of hCMV DNA in peripheral blood. These cells were in cell cycle and showed the features of recently activated cells (14). In contrast, cytotoxic CD28

⁻

CD4+_{T cells appeared in the circulation only after cessation of viral replication}

and were detectable in much higher frequencies in hCMV-seropositive individuals during latency. The very low percentages (<0.5%) of CD28

⁻

CD4+_{cells in hCMV-seronegative}

individuals might be the result of sterile hCMV infections or they could be induced by infections with other pathogens. The percentages of cytotoxic CD28

⁻

CD4+_{T cells were}

higher in hCMV-seropositive renal transplant recipients than in healthy individuals, which corresponds with the higher percentages of hCMV-specific, effector-type T cells during immunosuppression (18, 30-32). Within the group of hCMV-seropositive renal transplant recipients, a dichotomy is observed. We related the percentages of CD28

⁻

CD4+_{cells to different parameters like prior primary hCMV infection after}

transplantation, number of rejections, age, and the development of chronic rejection, but none of these seemed to explain the division in the two groups. This contrasts with a recent publication that states that patients with chronic kidney graft rejection have higher percentages of CD28

⁻

CD4+_{cells (33). We could demonstrate hCMV specificity}

of a considerable portion of CD28

⁻

CD4+_{cells because both proliferation and cytokine}

production were induced by hCMV stimulation and not by stimulation with other Ags such as PPD or VZV. This is in line with recent data for other Ags because in patients with chronic beryllium disease, CD28

⁻

CD4+_{cells from peripheral blood did not produce}

IFN-

γ

when stimulated with beryllium (BeSO4), whereas CD28+_CD4+_{cells did (34).}

The increase in CD28

⁻

CD4+_{cells in the peripheral blood compartment is only after}

the viral load became undetectable. This might be explained by entry of CD28

⁻

CD4+

cells into the circulation from the infected tissues only once the acute infection is over. Apart from this redistribution effect, differentiation of the cells also can cause the appearance of CD28

⁻

CD4+_{cells. As described for CD8}+_{T cells in primary hCMV}

infection, cells change their phenotype during differentiation. T cells lose the expression of CD28 (and CD27 and CCR7), but this is a slow process that even continues long after the antigenic load has become undetectable (13). Thus, the period between start of the infection and appearance of CD28

⁻

CD4+_{cells may reflect the time needed to}

(18)

CD28

⁻

CD4

+

7

The regulation of CD28 and CD27 expression on T cells is not completely understood. Naive cells express both costimulatory molecules, and during differentiation, expression can be lost simultaneously with acquisition of effector functions (28). However, it seems that the order of changes in phenotype is not similar for CD8+_{and CD4}+_{T cells.}

Differentiating CD8+_{T cells first lose expression of CD28 and only in a later phase}

they lose CD27, thus CD27

⁻

CD8+_{cells always have lost CD28 expression (13). For}

CD4+_{T cells it seems to be the opposite: all CD28}

⁻

_{cells are CD27}

⁻

_{but not vice versa,}

indicating that during differentiation CD4+_{T cells first lose expression of CD27 (this}

paper and Ref. (35)). CD28

⁻

CD4+_{cells are not commonly seen and, as we show in this}

study, infection with hCMV is the major factor causing this differentiation step of CD4+

T cells. In contrast with reports describing that the CD28

⁻

phenotype is stable and that CD28 expression cannot be restored (8, 36), CD28 was clearly up-regulated on CD28

⁻

CD4+_{T cells that proliferated after hCMV stimulation. This is in line with previous}

data showing re-expression of CD28 in CD28

⁻

T cell clones after anti-CD3 stimulation (37). In addition, it was recently shown that CD4+_CD28

⁻

_{T cells can become CD28}+

after stimulation with anti-CD3 in combination with IL-12 (38). CD28-B7 interactions provide important costimulatory signals for T cell activation. CD28

⁻

T cells cannot be stimulated anymore via this pathway, which could mean that these cells can function independently of costimulation. However, Park et al. (39) showed that CD28

⁻

CD4+_cells

proliferate better in the presence of accessory cells, suggesting that CD28

⁻

T cells are not necessarily costimulation-independent but could receive signals via molecules other than CD28. Recently, 4-1BB ligand has been shown to costimulate CD28

⁻

T cells (40), and we found that cytokine production by CD28

⁻

CD4+_{cells can be enhanced by}

adding an Ab against VLA-4 (CD49d; data not shown). In a paper by Suni et al. (41), CD4+_CD8dim_{T cells were described to be enriched for hCMV-specific cells. Indeed, we}

found that a small percentage (0–20%) of CD4+_CD28

⁻

_{T cells expressed low levels of}

CD8, whereas this was hardly seen in CD28+_CD4+_{T cells. Thus, CD4}+_CD8dim_{T cells}

probably represent a subpopulation of CD28

⁻

CD4+_{T cells.}

What exact function cytotoxic CD28

⁻

CD4+_{T cells have in controlling hCMV}

infection is not clear yet. One of the immune-evasion strategies of hCMV is to reduce MHC class I expression and thereby impede CD8+_{T cell immune surveillance (42, 43).}

Therefore, the role of CD4+_{T cells and their recognition of MHC class II may be critical}

for activating the immune system and sustaining the balance between virus and host immunity during latency. This is in agreement with the need for hCMV-specific CD4+

T cells to protect against hCMV disease (13). Another possible function of cytotoxic CD28

⁻

CD4+_{T cells might be a role in a negative feedback loop, namely eliminating}

APCs to dampen the immune response as described for cytotoxic CD8+_{T cells (44).}

Although CD28

⁻

CD4+_{T cells proliferate and secrete IFN-}

γ

_{only upon hCMV}

stimulation, not all CD28

⁻

CD4+_{T cells responded in these in vitro assays, which may}

seem contradictory in relation to the finding that CD28

⁻

CD4+_{cells only appear after}

(19)

CD28

⁻

CD4

+

7

CD4+_{T cells were generated in vivo upon infection with hCMV, and it could well}

be that the peptides presented during infection and where they are presented are not exactly the same as during the experimental hCMV stimulation. Second, the in vitro stimulation is performed with an inactivated laboratory hCMV strain, which may induce a different immune response than would the primary infection with one of the different natural hCMV strains. Finally, it is well known that hCMV is able to encode a range of gene products and uses several mechanisms to manipulate the host immune system (42, 43). Part of the CD28

⁻

CD4+_{T cells therefore could be inhibited in their}

response to hCMV or, alternatively, could be directed not against the virus itself but against molecules induced by hCMV infection.

Enhanced numbers of CD28

⁻

CD4+_{cells have been found in patients with RA and}

with cardiovascular diseases (2, 3, 5, 6). This raises the question of whether hCMV infection plays a role in these diseases. Also, in RA high frequencies of CD28

⁻

T cells were associated with hCMV seropositivity (9). Furthermore, hCMV DNA has been detected in synovial tissue and fluid of arthritis patients, and subpopulations of CD8+

synovial fluid mononuclear cells showed hCMV-specificity (45, 46). Thinking of the association among T cells, RA, and hCMV, it could be that part of the CD28

⁻

CD4+

T cells in patients with RA responds specifically to hCMV and possibly cross-reacts with other Ags. HCMV infection has also been associated with the development of cardiovascular diseases, as described in several papers (47-50). Interestingly, it was recently described that the increased levels in CD28

⁻

CD8+_{T cells in patients with}

coronary artery disease were mainly determined by hCMV seropositivity (51). Although data are not conclusive yet, numerous studies also suggest an influence of hCMV infection in triggering chronic rejection of different organ grafts in humans as well as in experimental animal models (reviewed in Ref. (52)).

So, how are hCMV and effector-type T cells involved in the tissue damage occurring in RA, cardiovascular diseases, and graft rejection? HCMV establishes latency in various cell types, including myeloid lineage cells but also endothelial cells. Inflammatory processes caused by different stimuli may cause reactivation of hCMV in the endothelial cells, which could directly lead to vascular pathology. Apart from the direct effects from hCMV, the tissue damage in the different diseases can also result from immunopathology. Once there is a reactivation of hCMV and virus is produced in endothelial cells, this will attract cytotoxic CD4+_{and CD8}+_{T cells, which are induced}

in high numbers by hCMV infection. The migration of T cells to the site of infection could be mediated by the inducible chemokine receptor CXCR3, which we found to be expressed on variable percentages (up to 100%) of CD28

⁻

CD4+_{T cells (figure 4C).}

In addition, it has been described that upon hCMV infection, expression of MHC class II molecules on endothelial cells is induced, which means that CD4+_{T cells also can}

recognize the presented Ags (53). At the site of infection, the T cells present will produce inflammatory cytokines and will have high levels of granzyme B and perforin, which can cause tissue damage to the endothelial cells possibly resulting in an increase

(20)

CD28

⁻

CD4

+

7

in atherosclerotic, vasculitic, or extra-articular rheumatoid lesions. Altogether, this provides a link between the presence of CD28

⁻

CD4+_{cells in RA and cardiovascular}

diseases and the data from this paper showing that CD28

⁻

CD4+_{cells emerge as a}

consequence of hCMV infection.

ACkNOwLEDGMENTS

We thank the patients and the healthy volunteers for their blood donations, Berend Hooibrink (Department of Cell Biology and Histology, Academic Medical Center, Amsterdam, The Netherlands) for sorting the different cell populations, technicians from the Department of Clinical Virology for performing hCMV PCRs and hCMV and EBV serology, and Drs. Eric Eldering and Martijn A. Nolte (both from Laboratory for Experimental Immunology, Academic Medical Center, Amsterdam, The Netherlands) for critical reading of the manuscript.

(21)

CD28

⁻

CD4

+

7

1. Appay V, Zaunders JJ, Papagno L, Sutton J, Jaramillo A, Waters A, Easterbrook P, Grey P, Smith D, McMichael AJ, et al. Characterization of CD4(+) CTLs ex vivo.

Journal of immunology (Baltimore, Md : 1950). 2002;168(11):5954-8.

2. Martens PB, Goronzy JJ, Schaid D, and Weyand CM. Expansion of unusual CD4+ T cells in severe rheumatoid arthritis. Arthritis

and rheumatism. 1997;40(6):1106-14.

3. Schmidt D, Goronzy JJ, and Weyand CM. CD4+ CD7- CD28- T cells are expanded in rheumatoid arthritis and are characterized by autoreactivity. The Journal of clinical

investigation. 1996;97(9):2027-37.

4. Schmidt D, Martens PB, Weyand CM, and Goronzy JJ. The repertoire of CD4+ CD28- T cells in rheumatoid arthritis.

Molecular medicine (Cambridge, Mass).

1996;2(5):608-18.

5. Liuzzo G, Kopecky SL, Frye RL, O’Fallon WM, Maseri A, Goronzy JJ, and Weyand CM. Perturbation of the T-cell repertoire in patients with unstable angina. Circulation. 1999;100(21):2135-9.

6. Nakajima T, Schulte S, Warrington KJ, Kopecky SL, Frye RL, Goronzy JJ, and Weyand CM. T-cell-mediated lysis of endothelial cells in acute coronary syndromes. Circulation. 2002;105(5):570-5. 7. Posnett DN, Sinha R, Kabak S, and Russo

C. Clonal populations of T cells in normal elderly humans: the T cell equivalent to “benign monoclonal gammapathy”.

The Journal of experimental medicine.

1994;179(2):609-18.

8. Vallejo AN, Nestel AR, Schirmer M, Weyand CM, and Goronzy JJ. Aging-related deficiency of CD28 expression in CD4+ T cells is associated with the loss of gene-specific nuclear factor binding activity. The Journal of biological

chemistry. 1998;273(14):8119-29.

9. Hooper M, Kallas EG, Coffin D, Campbell D, Evans TG, and Looney RJ. Cytomegalovirus seropositivity is associated with the expansion of CD4+CD28- and CD8+CD28- T cells in rheumatoid arthritis. The Journal of

rheumatology. 1999;26(7):1452-7.

10. Khan N, Shariff N, Cobbold M, Bruton R, Ainsworth JA, Sinclair AJ, Nayak L, and Moss PA. Cytomegalovirus seropositivity drives

the CD8 T cell repertoire toward greater clonality in healthy elderly individuals.

11. Looney RJ, Falsey A, Campbell D, Torres A, Kolassa J, Brower C, McCann R, Menegus M, McCormick K, Frampton M, et al. Role of cytomegalovirus in the T cell changes seen in elderly individuals. Clinical immunology

(Orlando, Fla). 1999;90(2):213-9.

12. Emery VC. Investigation of CMV disease in immunocompromised patients. Journal of

clinical pathology. 2001;54(2):84-8.

13. Gamadia LE, Remmerswaal EB, Weel JF, Bemelman FJ, van Lier RA, and ten Berge IJ. Primary immune responses to human CMV: a critical role for IFN-gamma-producing CD4+ T cells in protection against CMV disease. Blood. 2003;101(7):2686-92. 14. Rentenaar RJ, Gamadia LE, van der Hoek

N, van Diepen FN, Boom R, Weel JF, Wertheim-van Dillen PM, van Lier RA, and ten Berge IJ. Development of virus-specific CD4(+) T cells during primary cytomegalovirus infection. The Journal of

clinical investigation. 2000;105(4):541-8.

15. Evans TG, Kallas EG, Luque AE, Menegus M, McNair C, and Looney RJ. Expansion of the CD57 subset of CD8 T cells in HIV-1 infection is related to CMV serostatus. AIDS

(London, England). 1999;13(9):1139-41.

16. Gratama JW, Naipal AM, Oosterveer MA, Stijnen T, Kluin-Nelemans HC, Ginsel LA, den Ottolander GJ, Hekker AC, D’Amaro J, van der Giessen M, et al. Effects of herpes virus carrier status on peripheral T lymphocyte subsets. Blood. 1987;70(2):516-23.

17. Wang EC, Moss PA, Frodsham P, Lehner PJ, Bell JI, and Borysiewicz LK. CD8highCD57+ T lymphocytes in normal, healthy individuals are oligoclonal and respond to human cytomegalovirus.

18. Kuijpers TW, Vossen MT, Gent MR, Davin JC, Roos MT, Wertheim-van Dillen PM, Weel JF, Baars PA, and van Lier RA. Frequencies of circulating cytolytic, CD45RA+CD27-, CD8+ T lymphocytes depend on infection with CMV. Journal

of immunology (Baltimore, Md : 1950).

(22)

CD28

⁻

CD4

+

7

19. Boom R, Sol C, Weel J, Gerrits Y, de Boer M, and Wertheim-van Dillen P. A highly sensitive assay for detection and quantitation of human cytomegalovirus DNA in serum and plasma by PCR and electrochemiluminescence. Journal of

clinical microbiology. 1999;37(5):1489-97.

20. Maslanka K, Piatek T, Gorski J, Yassai M, and Gorski J. Molecular analysis of T cell repertoires. Spectratypes generated by multiplex polymerase chain reaction and evaluated by radioactivity or fluorescence.

Human immunology. 1995;44(1):28-34.

21. Schelonka RL, Raaphorst FM, Infante D, Kraig E, Teale JM, and Infante AJ. T cell receptor repertoire diversity and clonal expansion in human neonates. Pediatric

research. 1998;43(3):396-402.

22. Hirokawa M, Horiuchi T, Kawabata Y, Kitabayashi A, Saitoh H, Ichikawa Y, Matsutani T, Yoshioka T, Tsuruta Y, Suzuki R, et al. Oligoclonal expansion of CD4(+) CD28(-) T lymphocytes in recipients of allogeneic hematopoietic cell grafts and identification of the same T cell clones within both CD4(+)CD28(+) and CD4(+) CD28(-) T cell subsets. Bone marrow

transplantation. 2001;27(10):1095-100.

23. Liuzzo G, Goronzy JJ, Yang H, Kopecky SL, Holmes DR, Frye RL, and Weyand CM. Monoclonal T-cell proliferation and plaque instability in acute coronary syndromes.

Circulation. 2000;101(25):2883-8.

24. Serrano D, Monteiro J, Allen SL, Kolitz J, Schulman P, Lichtman SM, Buchbinder A, Vinciguerra VP, Chiorazzi N, and Gregersen PK. Clonal expansion within the CD4+CD57+ and CD8+CD57+ T cell subsets in chronic lymphocytic leukemia.

25. Wagner U, Pierer M, Kaltenhauser S, Wilke B, Seidel W, Arnold S, and Hantzschel H. Clonally expanded CD4+CD28null T cells in rheumatoid arthritis use distinct combinations of T cell receptor BV and BJ elements. European journal of

immunology. 2003;33(1):79-84.

26. Roosnek EE, Brouwer MC, Kipp JB, and Aarden LA. Monocyte-dependent induction of proliferation of human peripheral T cells by recombinant interleukin 2. European journal of

immunology. 1986;16(1):35-40.

27. Morishita Y, Martin PJ, Bean MA, Yamada H, and Hansen JA. Antigen-specific functions of a CD4+ subset of human T lymphocytes with granular morphology.

28. Hamann D, Baars PA, Rep MH, Hooibrink B, Kerkhof-Garde SR, Klein MR, and van Lier RA. Phenotypic and functional separation of memory and effector human CD8+ T cells. The Journal of experimental

medicine. 1997;186(9):1407-18.

29. Vossen MT, Gent MR, Weel JF, de Jong MD, van Lier RA, and Kuijpers TW. Development of virus-specific CD4+ T cells on reexposure to Varicella-Zoster virus. The Journal of infectious diseases. 2004;190(1):72-82.

30. Gamadia LE, Rentenaar RJ, Baars PA, Remmerswaal EB, Surachno S, Weel JF, Toebes M, Schumacher TN, ten Berge IJ, and van Lier RA. Differentiation of cytomegalovirus-specific CD8(+) T cells in healthy and immunosuppressed virus carriers. Blood. 2001;98(3):754-61. 31. Mackus WJ, Frakking FN, Grummels A,

Gamadia LE, De Bree GJ, Hamann D, Van Lier RA, and Van Oers MH. Expansion of CMV-specific CD8+CD45RA+CD27- T cells in B-cell chronic lymphocytic leukemia. Blood. 2003;102(3):1057-63. 32. Waldrop SL, Pitcher CJ, Peterson DM,

Maino VC, and Picker LJ. Determination of antigen-specific memory/effector CD4+ T cell frequencies by flow cytometry: evidence for a novel, antigen-specific homeostatic mechanism in HIV-associated immunodeficiency. The Journal of clinical

investigation. 1997;99(7):1739-50.

33. Pawlik A, Florczak M, Masiuk M, Dutkiewicz G, Machalinski B, Rozanski J, Domanski L, and Gawronska-Szklarz B. The expansion of CD4+CD28- T cells in patients with chronic kidney graft rejection. Transplantation

proceedings. 2003;35(8):2902-4.

34. Fontenot AP, Gharavi L, Bennett SR, Canavera SJ, Newman LS, and Kotzin BL. CD28 costimulation independence of target organ versus circulating memory antigen-specific CD4+ T cells. The Journal of clinical

investigation. 2003;112(5):776-84.

35. Amyes E, Hatton C, Montamat-Sicotte D, Gudgeon N, Rickinson AB, McMichael

(23)

CD28

⁻

CD4

+

7

AJ, and Callan MF. Characterization of the CD4+ T cell response to Epstein-Barr virus during primary and persistent infection.

The Journal of experimental medicine.

2003;198(6):903-11.

36. Vallejo AN, Weyand CM, and Goronzy JJ. Functional disruption of the CD28 gene transcriptional initiator in senescent T cells. The Journal of biological chemistry. 2001;276(4):2565-70.

37. Wierenga EA, Snoek M, Jansen HM, Bos JD, van Lier RA, and Kapsenberg ML. Human atopen-specific types 1 and 2 T helper cell clones. Journal of immunology (Baltimore,

Md : 1950). 1991;147(9):2942-9.

38. Warrington KJ, Vallejo AN, Weyand CM, and Goronzy JJ. CD28 loss in senescent CD4+ T cells: reversal by interleukin-12 stimulation. Blood. 2003;101(9):3543-9. 39. Park W, Weyand CM, Schmidt D, and

Goronzy JJ. Co-stimulatory pathways controlling activation and peripheral tolerance of human CD4+CD28- T cells. European journal of immunology. 1997;27(5):1082-90.

40. Bukczynski J, Wen T, and Watts TH. Costimulation of human CD28- T cells by 4-1BB ligand. European journal of

immunology. 2003;33(2):446-54.

41. Suni MA, Ghanekar SA, Houck DW, Maecker HT, Wormsley SB, Picker LJ, Moss RB, and Maino VC. CD4(+)CD8(dim) T lymphocytes exhibit enhanced cytokine expression, proliferation and cytotoxic activity in response to HCMV and HIV-1 antigens. European journal of

immunology. 2001;31(8):2512-20.

42. Mocarski ES, Jr. Immunomodulation by cytomegaloviruses: manipulative strategies beyond evasion. Trends in

microbiology. 2002;10(7):332-9.

43. Reddehase MJ. Antigens and immunoevasins: opponents in cytomegalovirus immune surveillance. Nature reviews Immunology. 2002;2(11):831-44.

44. Wong P, and Pamer EG. Feedback regulation of pathogen-specific T cell priming. Immunity. 2003;18(4):499-511. 45. Stahl HD, Hubner B, Seidl B, Liebert UG,

van der Heijden IM, Wilbrink B, Kraan MC, Emmrich F, and Tak PP. Detection of multiple viral DNA species in synovial

tissue and fluid of patients with early arthritis. Annals of the rheumatic diseases. 2000;59(5):342-6.

46. Tan LC, Mowat AG, Fazou C, Rostron T, Roskell H, Dunbar PR, Tournay C, Romagne F, Peyrat MA, Houssaint E, et al. Specificity of T cells in synovial fluid: high frequencies of CD8(+) T cells that are specific for certain viral epitopes. Arthritis

research. 2000;2(2):154-64.

47. Adam E, Melnick JL, Probtsfield JL, Petrie BL, Burek J, Bailey KR, McCollum CH, and DeBakey ME. High levels of cytomegalovirus antibody in patients requiring vascular surgery for atherosclerosis. Lancet. 1987;2(8554):291-3.

48. Melnick JL, Hu C, Burek J, Adam E, and DeBakey ME. Cytomegalovirus DNA in arterial walls of patients with atherosclerosis. Journal of medical

virology. 1994;42(2):170-4.

49. Nieto FJ, Adam E, Sorlie P, Farzadegan H, Melnick JL, Comstock GW, and Szklo M. Cohort study of cytomegalovirus infection as a risk factor for carotid intimal-medial thickening, a measure of subclinical atherosclerosis. Circulation. 1996;94(5):922-7.

50. Streblow DN, Kreklywich C, Yin Q, De La Melena VT, Corless CL, Smith PA, Brakebill C, Cook JW, Vink C, Bruggeman CA, et al. Cytomegalovirus-mediated upregulation of chemokine expression correlates with the acceleration of chronic rejection in rat heart transplants. Journal of virology. 2003;77(3):2182-94.

51. Jonasson L, Tompa A, and Wikby A. Expansion of peripheral CD8+ T cells in patients with coronary artery disease: relation to cytomegalovirus infection. Journal of internal medicine. 2003;254(5):472-8.

52. Cainelli F, and Vento S. Infections and solid organ transplant rejection: a cause-and-effect relationship? The Lancet infectious

diseases. 2002;2(9):539-49.

53. Waldman WJ, Knight DA, Adams PW, Orosz CG, and Sedmak DD. In vitro induction of endothelial HLA class II antigen expression by cytomegalovirus-activated CD4+ T cells. Transplantation. 1993;56(6):1504-12.