From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
IMMUNOBIOLOGY
Impaired function of human T-lymphotropic virus type 1 (HTLV-1)–specific
CD8⫹ T cells in HTLV-1–associated neurologic disease
Amir H. Sabouri,1 Koichiro Usuku,2 Daisuke Hayashi,1 Shuji Izumo,3 Yoshiro Ohara,4 Mitsuhiro Osame,1 and Mineki Saito4
1Department
of Neurology and Geriatrics, Kagoshima University Graduate School of Medical and Dental Sciences, Kagoshima; 2Department of Medical
Information Technology and Administration Planning, Kumamoto University Hospital, Kumamoto; 3Center for Chronic Viral Diseases, Kagoshima University
Graduate School of Medical and Dental Sciences, Kagoshima; and 4Department of Microbiology, Kanazawa Medical University, Ishikawa, Japan
Despite abundant activated virus-specific
cytotoxic T lymphocytes (CTLs), patients
with human T-lymphotropic virus type 1
(HTLV-1)–associated myelopathy/tropical
spastic paraparesis (HAM/TSP) showed a
significantly higher frequency of infected
T cells than did healthy virus carriers
(HVCs). Here, we demonstrate that at a
given proviral load, the frequency of CD8ⴙ
T cells that are negative for specific costimulatory molecules was significantly
higher in HAM/TSP than in age-matched
HVCs and uninfected healthy controls
(HCs), whereas the frequency of intracellular perforin-positive CD8ⴙ T cells was
significantly lower in both HAM/TSP and
HVCs than in HCs. An inverse correlation
between HTLV-1 proviral load (PVL) and
percent perforin-positive CD8ⴙ T cells
were observed only in disease-protective
allele HLA-A*02–positive HVCs, but not in
HAM/TSP patients, whether HLA-A*02
positive or negative, nor in HLA-A*02–
negative HVCs. Significantly lower perforin expression was observed in HTLV-1–
specific than in cytomegalovirus-specific
CD8ⴙ T cells. Majority of HTLV-1–specific
CD8ⴙ T cells in HVCs showed a
CD28ⴚCD27ⴙ phenotype, whereas HAM/
TSP showed a CD28ⴚCD27ⴚ phenotype.
HTLV-1–specific CD8ⴙ T cells from HAM/
TSP patients showed significantly lower
degranulation than HVCs by CD107a mobilization assay. These findings suggest
that an impaired function of HTLV-1–
specific CTLs is associated with failing
antiviral control and disease HAM/TSP.
(Blood. 2008;112:2411-2420)
Introduction
Human T-cell lymphotropic virus type 1 (HTLV-1) is a replicationcompetent human retrovirus1,2 associated with adult T-cell leukemia (ATL)3,4 and a slowly progressive neurologic disorder, HTLV-1–
associated myelopathy/tropical spastic paraparesis (HAM/TSP).5,6
The main pathologic features of HAM/TSP are chronic inflammation in the spinal cord, characterized by perivascular lymphocytic
cuffing and parenchymal lymphocytic infiltration including HTLV1–infected CD4⫹ T cells.7 Unlike human immunodeficiency virus
(HIV), HTLV-1 causes no disease in a majority of infected subjects
(healthy asymptomatic virus carriers, HVCs). However, approximately 2% to 3% develop ATL and another 2% to 3% develop a
disabling chronic inflammatory disease involving the central
nervous system (HAM/TSP), eyes, lungs, or skeletal muscles.8 Our
previous studies in HTLV-1 endemic to southern Japan indicated
that the median proviral load (PVL) in peripheral blood mononuclear cells (PBMCs) of patients with HAM/TSP was more than
10 times higher than that in HVCs, and a high PVL was also
associated with an increased risk of progression to disease.9 In the
same population, HLA-A*02 and HLA-Cw*08 genes were independently and significantly associated with a lower PVL and a lower
risk of HAM/TSP.10,11 Namely, possession of the HLA-A*02 allele,
which efficiently presents several epitopes from HTLV-1 proteins
to specific CTLs, was associated with protection against HAM/TSP
as well as a lower PVL.11 These data suggest that class I–restricted
CD8⫹ CTLs play an important part in controlling HTLV-1 PVL and
reducing the risk of disease. Indeed, CD8⫹ T cells in fresh PBMCs
of HTLV-1–infected patients rapidly kill autologous CD4⫹ cells
naturally infected with HTLV-1 by a perforin-dependent mechanism.12,13 However, because HTLV-1–specific CD8⫹ T cells have
the potential to produce proinflammatory cytokines,14 there is a
debate on the role of HTLV-1–specific CD8⫹ T cells; that is,
whether these cells contribute to the inflammatory and demyelinating processes of HAM/TSP, or whether the dominant effect of
such cells in vivo is protective against disease, although these
2 mechanisms are not mutually exclusive.
Antigen-specific CTLs are crucial components of the immune
response against viruses, and they have been shown to have an
important role in limiting viral replication and controlling virusassociated diseases.15 A decisive role of HTLV-1–specific CTLs has
also been suggested by host genetics,10,11 in vitro T-cell function,12,13 and DNA expression microarray analysis.16 However, in
HTLV-1 infection, active and abundant HTLV-1–specific CTLs do
not completely eliminate infected cells, especially in patients with
HAM/TSP. We have therefore hypothesized that the differences in
HTLV-1 PVL and associated differences in the risk of HAM/TSP
should be associated with differences between patients in the
efficiency of CTL-mediated lysis of infected cells. Because both
positive and zero correlations have been found between the
frequency of HTLV-1–specific CD8⫹ T cells and PVL,17-19 the
frequency of HTLV-1–specific CTL itself is not an accurate
indicator of CTL strength or effectiveness. Meanwhile, DNA
expression microarray analysis suggested that a high level of
expression of lymphocyte lysis-related genes, including granzymes, perforin, granulysin, and NKG2D, was associated with
Submitted February 18, 2008; accepted April 24, 2008. Prepublished online as
Blood First Edition paper, May 27, 2008; DOI 10.1182/blood-2008-02-140335.
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 USC section 1734.
An Inside Blood analysis of this article appears at the front of this issue.
The online version of this article contains a data supplement.
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
© 2008 by The American Society of Hematology
2411
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
2412
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
SABOURI et al
Table 1. Characteristics of HTLV-1–infected patients and healthy
uninfected controls
Patient characteristic
HAM/TSP
n ⴝ 72
HVCs
n ⴝ 96
HCs
n ⴝ 20
Age, y, mean ⫾ SE
55.9 ⫾ 1.6
51.1 ⫾ 1.2
55.2 ⫾ 1.9
Sex
Male
27
51
13
Female
45
45
7
36 932 ⫾ 8414
2106 ⫾ 990
N/A
16 348
1024
N/A
793.2 ⫾ 677
245.0 ⫾ 381
N/A
537.0
109.0
N/A
Serum anti–HTLV-I antibody
titer*
Mean ⫾ SE
Median
HTLV-I provirus load in PBMCs†
Mean ⫾ SE
Median
HVCs indicates HTLV-1 carriers; HCs, uninfected healthy controls; N/A, not
applicable.
*Anti–HTLV-1 antibodies were titrated by the particle agglutination method.
†HTLV-1 tax copy number per 104 PBMCs.
lower PVL both in patients with HAM/TSP and in HVCs.16 Moreover,
differentiation of CD8⫹ T cells and their functional profile, such as
secretion of cytotoxic molecules, cytokines, and degranulation markers,
seems to play a key role in controlling several other chronic viral
infections such as herpes simplex virus (HSV), Epstein-Barr virus
(EBV), cytomegalovirus (CMV), and HIV.20-22 In this study, based on
these observations, we have evaluated the expression of costimulatory
markers, cytotoxic granules (perforin and GzmB), and degranulation
marker CD107a in CD8⫹ T cells from HTLV-1–infected patients
(HAM/TSP and HVCs) and age-matched healthy controls (HCs), and
compared them with HTLV-1 PVL.
Methods
Patients and cells
Peripheral blood was studied from 72 patients with a clinical diagnosis of
HAM/TSP, 96 HVCs, and 20 HCs in total. Characteristics of these patients are
shown in Table 1. Fresh PBMCs were isolated on a Histopaque-1077 (SigmaAldrich, St Louis, MO) density gradient centrifugation, washed twice in RPMI
1640 with 10% heat-inactivated fetal calf serum (FCS), and stored in liquid
nitrogen as stocked lymphocytes until use. All experiments were performed by
using age-matched samples selected from stocked lymphocytes. The diagnosis of
HAM/TSP was made according to the World Health Organization diagnostic
criteria.23 All patients and control subjects were Japanese and resided in
Kagoshima Prefecture, an HTLV-1 endemic region in southern Japan. Samples
from patients with HAM/TSP were studied only if the patients had never received
any immunomodulators (oral steroid or interferon-␣ injection) or more than 10
years had passed since the last immune-modulating therapies. This research was
approved by the institutional review boards of the authors’ institutions and
informed consent was obtained from all patients in accordance with the
Declaration of Helsinki.
Lymphocyte phenotyping and perforin detection by
flow cytometry
After thawing, cells were washed 3 times with phosphate-buffered
saline (PBS) and fixed in PBS containing 2% paraformaldehyde
(Sigma-Aldrich) for 20 minutes and resuspended in PBS at 4°C. Fixed
cells were washed with PBS containing 7% of normal goat serum
(Sigma-Aldrich) and then incubated for 15 minutes at room temperature
with various combinations of fluorescence-conjugated monoclonal
antibodies (mAbs) as follows: phycoerythrin-cyanin 5.1 (PC5)–labeled
anti-CD27, PC5-labeled anti-CD28, PC5-labeled anti-CD8, energycoupled dye (ECD)–labeled anti-CD8 (Beckman Coulter, Fullerton,
CA), phycoerythrin (PE)–labeled anti-CD28, fluorescein isothiocyanate
(FITC)–labeled antiperforin and GzmB (BD PharMingen, San Diego,
CA). Isotype-matched mouse immunoglobulins were used as a control.
The phenotype was determined by flow cytometry (EPICS XL; Beckman Coulter) in the lymphocyte gate based on forward versus side
scatter. For intracellular staining of perforin and GzmB, PBMCs were
first surface stained with the ECD-labeled anti-CD8 mAb for 15 minutes
at room temperature. After cell-surface labeling, cells were washed and
permeabilized with PBS/7% normal goat serum containing 0.2%
saponin (PBS-SAPO; Sigma-Aldrich) for 10 minutes at room temperature. The cells were then washed twice and resuspended for 20 minutes
at room temperature in PBS-SAPO containing FITC-labeled anticytotoxin mAbs (perforin, GzmB; BD PharMingen) or an isotype control
mAb. Finally, the cells were washed twice and analyzed by an EPICS
XL flow cytometer and EXPO32 analysis software (Beckman Coulter)
in the lymphocyte gate based on forward versus side scatter.
Detection of HTLV-1 Tax– and CMV pp65–specific CTLs by HLA
class I tetramer
To evaluate the expression of perforin in antigen-specific CD8⫹ T cells,
CD8⫹ T cells were stained with fluorescent-labeled tetramers of HLAA*02 ⫹ ␤2 microglobulin ⫹ Tax11-19 (LLFGYPVYV) or CMV pp65
(NLVPMVATV) peptides, which were purchased from Medical Biological
Laboratories (Nagoya, Japan). PBMCs were incubated with each tetramer
and ECD-labeled anti-CD8 mAb at 37°C for 25 minutes then washed
3 times in ice-cold PBS, fixed in 2% paraformaldehyde for 20 minutes at
4°C. Fixed PBMCs were then permeabilized with PBS/7% normal goat
serum (NGS) containing 0.2% PBS-SAPO for 10 minutes at room
temperature for intracellular staining of perforin and GzmB. The cells were
washed again and resuspended for 20 minutes at room temperature in
PBS-SAPO containing anticytotoxin mAbs (perforin, GzmB) or an isotype
control mAb. Finally, the cells were washed and analyzed by flow
cytometry. At least 5 ⫻ 104 events were collected in each assay.
Quantification of HTLV-1 proviral load and anti–HTLV-1
antibody titers
To examine the HTLV-1 PVL, we carried out a quantitative polymerase
chain reaction (PCR) method using ABI Prism 7700 (Applied Biosystems,
Foster City, CA) with 100 ng genomic DNA (roughly equivalent to 104
cells) from PBMC samples, as reported previously.9 Based on the standard
curve created by 4 known concentrations of template, the concentrations of
unknown samples were determined. Using ␤-actin as an internal control,
the amount of HTLV-1 proviral DNA was calculated by the following
formula: copy number of HTLV-1 tax per 1 ⫻ 104 PBMC ⫽ [(copy number
of tax) / (copy number of ␤-actin/2)] ⫻ 104. All samples were performed in
triplicate. Serum antibody titers to HTLV-1 were determined by a particle
agglutination method.
CD107a mobilization assay
To evaluate cell-mediated cytotoxicity of HTLV-1–specific CD8⫹ T cells in
patients with HAM/TSP and HVCs, we performed a CD107a mobilization
assay.24,25 This assay enables rapid assessment of cell-mediated cytotoxicity
via sensitive detection of CD107a exposed on the cell surface after antigen
stimulation and subsequent secretion of the lytic granule contents such as
perforin and granzymes. PBMCs (2 ⫻ 106) were cultured for 4 hours with
or without 1 ␮g/mL HTLV-1 Tax peptide in combination with FITC-labeled
anti-CD107a mAb and the secretion inhibitor monensin in RPMI 1640
complete medium with 50 IU/mL IL-2, according to the manufacturer’s
instruction (IMMUNOCYTO CD107a detection kit; MBL, Nagoya, Japan).
After incubation, cell suspensions were washed with PBS and the cells were
further stained with Tax-tetramer–PE and PC5-labeled anti-CD8 mAb
(Beckman Coulter). A minimum of 3 ⫻ 104 CD8⫹ T lymphocytes per
sample were acquired by flow cytometry.
Statistical analysis
To test for significant differences among the cell populations between
3 different groups of subjects (HAM/TSP, HVCs, and HCs), one-factor
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
HTLV-1–SPECIFIC CTLs IN CHRONIC INFECTION
2413
Table 2. Ex vivo frequency of CD8ⴙ T cells lacking the costimulatory molecules in peripheral blood mononuclear cells from
HTLV-1–infected patients and uninfected healthy controls
P (Sheffe F)
n
Mean ⴞ SE
HAM/TSP
16
HVCs
20
HAM/TSP
16
31.6 ⫾ 2.3
HVCs
20
22.3 ⫾ 1.7
HCs
20
19.7 ⫾ 1.3
P (ANOVA)
HAM-HCs
HVCs-HCs
805.1 ⫾ 461.0
.001
N/A
N/A
354.5 ⫾ 106.2
(Mann-Whitney)
.003
.004
.83
.003
.003
.91
.015
.012
.20
.009
.007
.23
.004
.003
.28
N/A
N/A
.012
.009
1.0
.006
.007
.78
.001
.001
.16
.007
.013
.11
.19
.32
.35
All (PVL-unmatched)
PVL
%CD8⫹CD27⫺*
%CD8⫹CD28⫺
HAM/TSP
16
31.4 ⫾ 3.1
HVCs
20
23.6 ⫾ 1.5
HCs
20
20.3 ⫾ 1.5
%CD8⫹CD80⫺
HAM/TSP
16
36.4 ⫾ 3.3
HVCs
20
32.3 ⫾ 1.6
HCs
20
26.2 ⫾ 1.7
%CD8⫹CD86⫺
HAM/TSP
16
35.3 ⫾ 3.2
HVCs
20
30.6 ⫾ 1.5
HCs
20
25.1 ⫾ 1.5
%CD8⫹CD152⫺
HAM/TSP
16
35.6 ⫾ 2.5
HVCs
20
30.1 ⫾ 1.9
HCs
20
25.2 ⫾ 1.7
HAM/TSP
8
529.8 ⫾ 79.0
.79
HVCs
9
486.0 ⫾ 104.5
(Mann-Whitney)
HAM/TSP
8
31.8 ⫾ 5.9
HVCs
9
19.8 ⫾ 3.6
20
19.7 ⫾ 1.3
HAM/TSP
8
30.6 ⫾ 3.2
HVCs
9
22.6 ⫾ 2.4
20
20.3 ⫾ 1.5
HAM/TSP
8
39.5 ⫾ 3.8
HVCs
9
32.5 ⫾ 2.1
20
26.2 ⫾ 1.7
HAM/TSP
8
35.3 ⫾ 4.2
HVCs
9
31.8 ⫾ 1.8
20
25.1 ⫾ 1.5
HAM/TSP
8
31.0 ⫾ 4.2
HVCs
9
30.5 ⫾ 2.8
20
25.2 ⫾ 1.7
PVL-matched
PVL
%CD8⫹CD27⫺
HCs
%CD8⫹CD28⫺
HCs
%CD8⫹CD80⫺
HCs
%CD8⫹CD86⫺
HCs
%CD8⫹CD152⫺
HCs
HVCs indicates HTLV-1 carriers; HCs, uninfected healthy controls; and PVL, proviral load.
One-factor ANOVA was done when variance of each group was equal by Levene test. If variance of each group was different, Kruskal-Wallis test was used. For multiple
comparison, we used Sheffe F to analyze the statistical difference.
* % CD8⫹CD27⫺ indicates the frequency of CD8⫹CD27⫺ in PBMCs.
analysis of variance (ANOVA) was done when the variance of each group
was equal by Levene test. If the variance of each group was different, the
Kruskal-Wallis test was used. For multiple comparisons, we used Sheffe F
to analyze statistical difference. The Mann-Whitney U test was used for
comparing the differences in the frequencies of cell populations or PVL between
patients with HAM/TSP and HVCs. The results represent the mean plus or minus
the standard error (SE) where applicable. Correlations between variables were
examined by Spearman rank correlation analysis. All statistical analyses were
performed using SPSS version 13.5 software (SPSS, Chicago, IL). Values of
P less than .05 were considered statistically significant.
Results
The expression of costimulatory molecules on T cells of
HTLV-1–infected patients and uninfected controls
To determine whether the frequency of costimulatory molecules
differed between patients with HAM/TSP and HVCs as well as
HCs, we stained PBMCs with monoclonal antibodies to CD27,
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
2414
SABOURI et al
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
Figure 1. Frequencies and mean fluorescence intensities of perforin- and granzyme B–positive CD8ⴙ T-cell populations in HTLV-1–infected patients.
(A) Representative dot plots of PBMCs stained for cytotoxic function markers and costimulatory molecules. PBMCs from patients with HAM/TSP, asymptomatic healthy virus
carriers (HVCs), and uninfected healthy controls (HCs) were costained for CD8, CD27, or CD28, and markers associated with cytotoxic function (ie, perforin or granzyme B
[GzmB]). Cells in a CD8⫹ lymphocyte gate were analyzed. Representative dot plots from 1 uninfected control and 1 HAM/TSP patient are shown. (B) The frequency and mean
fluorescence intensity (MFI) of the perforin-positive CD8⫹ T-cell population and its subpopulations (CD8⫹CD28⫺ and CD8⫹CD27⫺) in patients with HAM/TSP, HVCs, and HCs.
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
CD28, CD80, CD86, and CD152 (cytotoxic T-cell lymphocyte–
associated antigen-4 [CTLA-4]) in 3 groups of age-matched
patients with HAM/TSP, HVCs, and HCs (Table 2). The percentages of CD8⫹ cells that were negative for costimulatory molecules
were significantly higher in patients with HAM/TSP than in HCs;
however, these differences were not observed between HVCs and
HCs. Because the mean HTLV-1 PVL of the HAM/TSP group was
significantly higher than that of the HVCs (HAM/TSP
805.1 ⫾ 461.0 vs HVCs 354.5 ⫾ 106.2, P ⫽ .001, MannWhitney), we further compared the frequencies of each costimulatory molecule between patients with HAM/TSP and the HVCs with
a similar PVL (HAM/TSP 529.8 ⫾ 79.0 vs HVCs 486.0 ⫾ 104.5,
P ⫽ .79, Mann-Whitney). The mean frequency of each costimulatory molecule was still significantly higher in patients with
HAM/TSP than HCs except for CD8⫹ CD152⫺ (Table 2), and there
was no correlation between the mean frequency of each costimulatory molecule and PVL (data not shown). These data indicate that
the observed differences do not simply reflect differences in PVL
(ie, in the frequency of HTLV-1–infected cells). Furthermore,
although HTLV-1 mainly infects CD4⫹ cells, the percentages of
CD4⫹ cells that were negative for costimulatory molecules were
not significantly different between patients with HAM/TSP and
HVCs or HCs (Table S1, available on the Blood website; see the
Supplemental Materials link at the top of the online article),
indicating that the mean frequency of each costimulatory molecule
on CD4⫹ cells was not influenced by the PVL.
Expression of perforin and granzyme B in CD8ⴙCD28ⴚ and
CD8ⴙCD27ⴚ T-cell populations of patients with HAM/TSP and
healthy virus carriers
We next focused on the CD8⫹CD28⫺ T cells and CD8⫹CD27⫺
T cells, because these subsets include CTLs and because abnormal
expansion of these subsets was also described in chronic CMV,26
EBV,27 and HIV20 infection. As shown in Table 2, both percent
CD8⫹CD28⫺ and percent CD8⫹CD27⫺ T-cell populations in
PBMCs were significantly higher in patients with HAM/TSP than
in HVCs and HCs, whereas there were no significant differences of
percent CD8⫹CD28⫺ and percent CD8⫹CD27⫺ T-cell populations
between HVCs and HCs. Interestingly, as typical examples
show in Figure 1A, the percentages of perforin⫹CD8⫹,
perforin⫹CD8⫹CD28⫺, and perforin⫹CD8⫹CD27⫺ T cells were
significantly lower in HTLV-1–infected patients (patients with
HAM/TSP and HVCs) than in HCs, irrespective of PVL (Figure
1B,D). The differences between HVCs and patients with HAM/
TSP were smaller than the differences between HCs and HVCs or
patients with HAM/TSP alone, indicating that a selective decrease
of perforin expression in CD8⫹ T cells was associated with
persistent HTLV-1 infection. The mean fluorescence intensity
(MFI) of the perforin⫹CD8⫹, perforin⫹CD8⫹CD28⫺, and
perforin⫹CD8⫹CD27⫺ T-cell populations were significantly lower
in patients with HAM/TSP than in HCs, irrespective of PVL
(Figure 1B,D). In contrast to the perforin expression, both the
percentages and MFI of the GzmB⫹CD8⫹, GzmB ⫹CD8⫹CD28⫺,
HTLV-1–SPECIFIC CTLs IN CHRONIC INFECTION
2415
and GzmB ⫹CD8⫹CD27⫺ T-cell populations were not significantly
different among the 3 subject groups (patients with HAM/TSP,
HVCs, and HCs) when HTLV-1–infected patients were compared
with a similar proviral load (Figure 1C,E).
There was a marginal negative correlation between percent
perforin⫹CD8⫹ T cells and HTLV-1 PVL in all HTLV-1–infected
patients (patients with HAM/TSP ⫹ HVCs; P ⫽ .046, r ⫽ ⫺0.29
by Spearman rank correlation analysis) but not in patients with
HAM/TSP or HVCs alone (Figure 2 left panels). Meanwhile, a
significant negative correlation between percent perforin⫹CD8⫹
T cells and HTLV-1 PVL was observed in all HLA-A*02–positive
HTLV-1–infected patients (patients with HAM/TSP ⫹ HVCs;
P ⫽ .003, r ⫽ ⫺0.58) and HVCs alone (P ⫽ .024, r ⫽ ⫺0.64), but
not in patients with HAM/TSP (P ⫽ .15, r ⫽ ⫺0.35; Figure 2 right
panels). No correlation was observed between expression of GrzB
and HTLV-1 PVL in patients with HAM/TSP, HVCs, or both
groups combined, irrespective of HLA status (data not shown).
Lower perforin expression in HTLV-1–specific CD8ⴙ T cells
than in CMV-specific CD8ⴙ T cells within HTLV-1–infected
patients
Perforin is essential for killing targets of CTLs by means of the
granule exocytosis pathway, and its expression is more restricted
than that of serine proteases. As we observed significant differences
in perforin expression in CD8⫹ T cells lacking the costimulatory
molecules in PBMCs between HTLV-1–infected patients and
healthy controls, and a significant negative correlation between
percent perforin⫹CD8⫹ T cells and HTLV-1 PVL in HLA-A*02–
positive HTLV-1–infected patients, we next focused on the perforin
expression in HTLV-1–specific CD8⫹ T cells and CMV-specific
CD8⫹ T cells in the same group of HTLV-1–infected patients
(Figure 3A). Although there was no significant difference in
percent perforin expression in virus-specific (both HTLV-1 Tax–
specific and CMV-specific) CD8⫹ T cells between patients with
HAM/TSP and HVCs, the percentage perforin expression in
HTLV-1 Tax–specific CD8⫹ T cells was significantly lower than
CMV-specific CD8⫹ T cells in HTLV-1–infected patients (both
patients with HAM/TSP and HVCs) (Figure 3B). The expression of
perforin in CMV-specific CD8⫹ T cells was always higher than the
expression of perforin in Tax-specific CD8⫹ T cells in the same
patients in both patients with HAM/TSP and HVCs.
HTLV-1 Tax–specific CD8ⴙ T cells in patients with HAM/TSP and
HVCs appear to be at different expression patterns of
costimulatory molecule
Because it has been reported that the cell-surface phenotype of
virus-specific cells was affected by the stage of infection (ie, acute
or chronic) as well as by viral specificity,20 we further investigated
whether the cell-surface phenotype of HTLV-1–specific CD8⫹ cells
differed between patients with HAM/TSP and HVCs. According to
a simple classification of 3 functional subsets based on CD28 and
CD27 expression (ie, early [CD28⫹CD27⫹], intermediate
(C) Frequency and MFI of the GzmB-positive CD8⫹ T-cell population and its subpopulations (CD8⫹CD28⫺ and CD8⫹CD27⫺) in patients with HAM/TSP, HVCs, and HCs.
(D) Frequency and MFI of the perforin-positive CD8⫹ T-cell population and its subpopulations (CD8⫹CD28⫺ and CD8⫹CD27⫺) in proviral load (PVL)–matched patients with
HAM/TSP, HVCs, and HCs. (E) The frequency and MFI of the GzmB-positive CD8⫹ T-cell population and its subpopulations (CD8⫹CD28⫺ and CD8⫹CD27⫺) in PVL-matched
patients with HAM/TSP, HVCs, and HCs. The frequency of perforin- or GzmB-positive cells is shown as a percentage within each cell subset (CD8⫹, CD8⫹CD28⫺,
CD8⫹CD27⫺). Values represent the means plus or minus the standard error (SE). To test for significant differences among the cell populations between 3 different groups of
subjects (HAM/TSP, HVCs, and HCs), one-factor ANOVA was done when the variance of each group was equal by Levene test. If the variance of each group was different, the
Kruskal-Wallis test was used. For multiple comparisons, we used Sheffe F to analyze statistical difference. Values of P ⬍ .05 were considered statistically significant.
***P ⬍ .001; **P ⬍ .01; *P ⬍ .05.
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
2416
SABOURI et al
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
Figure 2. Perforin-positive CD8ⴙ T cells and HTLV-1
proviral load in HTLV-1 infection. (Left panels) There
was a marginal negative correlation between the percentages of perforin-positive CD8⫹ T cells and HTLV-1 proviral load (PVL) in whole HTLV-1–infected patients (HAM/
TSP ⫹ HVCs; P ⫽ .046, r ⫽ ⫺0.29 by Spearman rank
correlation analysis), but not in patients with HAM/TSP or
in HVCs alone. (Right panels) There was a significant
negative correlation between the percentages of perforinpositive CD8⫹ T cells and HTLV-1 PVL in whole HLAA*02–positive HTLV-1–infected patients (HAM/
TSP ⫹ HVCs; P ⫽ .003, r ⫽ ⫺0.58) and in HVCs alone
(P ⫽ .024, r ⫽ ⫺0.64), but not in patients with HAM/TSP
(P ⫽ .15, r ⫽ ⫺0.35). The x axis denotes percentage of
perforin-positive CD8⫹ T cells, and the y axis denotes
HTLV-1 PVL (copy number of HTLV-1 tax per 104
PBMCs ⫽ [(copy number of tax)/(copy number of ␤-actin/
2)] ⫻ 104). Data were analyzed by Spearman rank correlation. The solid line represents a least-squares regression line.
[CD28⫺CD27⫹], and late [CD28⫺CD27⫺] differentiated T cells;
Figure 4A),20 more than 60% of HTLV-1 Tax–specific CD8⫹ T
cells from patients with HAM/TSP showed late differentiated
phenotype, whereas more than 60% of HTLV-1 Tax–specific CD8⫹
T cells from HVCs showed intermediate differentiated phenotype
(Table 3 and Figure 4B,C). These phenotypes did not change even
when we compared the frequencies between patients with HAM/
TSP and the HVCs with a similar PVL (P ⫽ .11, Mann-Whitney U
test, Table 3). As previously reported, more than 80% of CMVtetramer–specific CD8⫹ T cells both from patients with HAM/TSP
and HVCs showed a late differentiated phenotype (Table 3 and
Figure 4B,C).20 As shown in Figure 4D, there was a negative
correlation between percent Tax-specific intermediate differentiated T cells and HTLV-1 PVL in the whole cohort (n ⫽ 31;
P ⫽ .008, r ⫽ ⫺0.47 by Spearman rank correlation analysis)
whereas there was a positive correlation between percent Taxspecific late differentiated T cells and HTLV-1 PVL in the whole
cohort (n ⫽ 31; P ⫽ .005, r ⫽ 0.49 by Spearman rank correlation
analysis). There was no correlation between percent Tax-specific
early differentiated T cells and HTLV-1 PVL in the whole cohort
(n ⫽ 31; P ⫽ .55, r ⫽ ⫺0.1 by Spearman rank correlation analysis).
CD107a mobilization assays
To determine the functional reactivity of HTLV-1 Tax-specific
CD8⫹ T cells, we performed a CD107a mobilization assay (Figure
5). PBMCs derived from HLA-A*02–positive HTLV-1–infected
patients (patients with HAM/TSP and HVCs) were stained with
HLA-A*02/ Tax11-19 tetramer after incubation with Tax11-19
peptide and anti-CD107a monoclonal antibody for 4 hours. Significant anti-CD107a staining was observed on the Tax-tetramer–
positive population after coculture with Tax11-19 peptide, whereas
only background staining was seen without peptide (Figure 5A).
We compared the frequency of expression of the degranulation
marker CD107a in Tax-tetramer–positive cells, which allows
measurement of cytolytic cell activation, between 10 patients with
HAM/TSP and 14 HVCs. The percentage of CD107a⫹ HLA-A*02/
Tax11-19 tetramer–positive cells within the HLA-A*02/Tax11-19
tetramer–positive gate (percent Tet⫹ CD107a⫹/ Tet⫹) was significantly lower in patients with HAM/TSP than in HVCs (Figure 5B).
The same results were obtained in a comparison between 5 patients
with HAM/TSP and 6 HVCs with similar PVL (patients with
HAM/TSP 307.2 ⫾ 125.3 vs HVCs 270.0 ⫾ 92.2, P ⫽ .57, MannWhitney), indicating that the observed difference did not simply
reflect the higher PVL (ie, the higher frequency of HTLV-1–
infected T cells) in patients with HAM/TSP (Figure 5C).
Discussion
To characterize the HTLV-1–specific CD8⫹ T cells in infected
patients, we first examined the expression of costimulatory molecules on CD4⫹ and CD8⫹ T cells, which have a significant impact
on the cytokine profile and proliferation response. Interestingly, the
percentages of CD8⫹ T cells that were negative for costimulatory
molecules CD27, CD28, CD152 (CTLA-4), CD80 (B7-1), and
CD86 (B7-2) were significantly higher in patients with HAM/TSP
compared with HVCs or HCs, whereas this discrepancy was not
found within the CD4⫹ T-cell population. Namely, there was a
selective decrease of costimulatory molecule expression on CD8⫹
T cells in patients with HAM/TSP. It is noteworthy that the same
differences were observed in a comparison between patients with
HAM/TSP and HVCs with a similar PVL, whereas there were no
significant differences between the patients with HAM/TSP and
HVCs in either the frequency of CD4⫹ cells negative for costimulatory molecules or the PVL in PBMCs. These findings suggest that
the expression of costimulatory molecules was influenced by the
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
HTLV-1–SPECIFIC CTLs IN CHRONIC INFECTION
2417
Figure 3. Perforin expression in HTLV-1 Tax– and
CMV pp65–specific CD8ⴙ T cells in HTLV-1 infection.
(A) Left panel: Dot plots of HLA-A*02/Tax11-19 tetramer-PE (x axis) versus CD8-ECD fluorescence within
the lymphocyte gate based on forward versus side
scatter. Right panel: Histogram of perforin expression
(x axis, arbitrary units, log scale) versus cell number
(y axis) in gated virus-specific (tetramer-positive) cells.
Representative dot plot from one patient with HAM/TSP is
shown. (B) The perforin expression in HTLV-1 Tax– and
CMV pp65–specific CD8⫹ T cells in HTLV-1–infected
patients. Statistical analysis was done as described in the
legend for Figure 1. ***P ⬍ .001
disease status rather than the PVL. The emergence of high
frequencies of CD8⫹ T cells negative for costimulatory molecules
might have been caused by a greater degree of continuous or
repeated antigenic stimulation in patients with HAM/TSP than in
HVCs at a similar PVL; such repeated antigenic stimulation might
lead immune cells into a loss of antiviral CD8⫹ T-cell function, as
suggested in HIV infection. In fact, it has been reported that the
number of naive T cells was low in HTLV-1–infected patients
compared with uninfected controls, whereas the number of memory
T lymphocytes was greater in HTLV-1–infected patients.28 This
finding supports the rapid turnover of T cells by antigenic
stimulation in HTLV-1–infected patients, which has recently been
directly demonstrated by metabolic labeling of lymphocytes in
vivo in human HTLV-1–infected subjects.29 We next focused on the
CD8⫹CD28⫺ T cells and CD8⫹CD27⫺ T cells, because these
subsets include CTLs and abnormal expansion of these subsets was
also described in chronic CMV,26 EBV,27 and HIV20 infection.
Especially, increased expression of CD8⫹CD28⫺ T cells was
associated with disease activity and CD8⫹ T-cell dysfunction in
HIV infection.30,31 As expected, the percentages of CD8⫹ cells that
were negative for CD28 or CD27 were significantly increased in
patients with HAM/TSP by comparison with HVCs or HCs,
irrespective of PVL.
Because impaired CTL-mediated lysis is a general feature of
chronic HIV, CMV, and EBV infections,32 especially when the
antigen load is high,33 it is important to examine the phenotypic and
functional property of CD8⫹ T cells in HTLV-1 infection. Our data
showed that the frequency of perforin and GzmB⫹CD8⫹,
CD8⫹CD28⫺, and CD8⫹CD27⫺ T cells in PBMCs was significantly lower in HTLV-1–infected patients (patients with HAM/TSP
and HVCs) than in HCs. Interestingly, although both the percentage
and MFI of perforin⫹CD8⫹, perforin⫹CD8⫹CD28⫺, and
perforin⫹CD8⫹CD27⫺ T-cell populations were significantly lower
in HTLV-1–infected patients (patients with HAM/TSP and HVCs)
than in HCs, irrespective of PVL, the MFIs of GzmB⫹CD8⫹,
GzmB ⫹CD8⫹CD28⫺, and GzmB⫹CD8⫹CD27⫺ T-cell popula-
tions were not significantly different among patients with HAM/
TSP, HVCs, and HCs compared with similar PVLs. Such discordant expression patterns between perforin and granzymes are a
common feature of circulating virus-specific CD8⫹ T cells in
chronic HIV, CMV, and EBV infection.32,34-37 Because perforin
staining often lessens after T-cell activation, presumably because of
the release of preformed perforin,34 decreased perforin expression
in HTLV-1 infection, especially patients with HAM/TSP, might
arise from aberrant perforin secretion or from frequent discharge of
perforin caused by constant antigenic stimulation.38 In this case, the
higher MFI of GzmB compared with perforin in CD8⫹ T cells of
patients with HAM/TSP might be a result of the lack of granzyme
release caused by lower perforin levels. Interestingly, there was a
significant negative correlation between percent perforin⫹CD8⫹
T cells and HTLV-1 PVL in HTLV-1–infected HLA-A*02–positive
HVCs, but not in HLA-A*02–negative HVCs and patients with
HAM/TSP who were either HLA-A*02 positive or negative. As the
possession of HLA-A*02 was associated with a significant reduction in both HTLV-1 PVL and the risk of HAM/TSP,11 these
findings suggest that the HLA-A*02–restricted CTLs are particularly efficient at killing HTLV-1–infected cells.39 The absence of
correlation between perforin-expressing CD8⫹ T cells and HTLV-1
PVL in patients with HAM/TSP also supports the hypothesis of
CD8⫹ T-cell inefficiency in the diseased situation.
A large number of phenotypic markers have been used to define
different populations of antigen-specific CD4⫹ and CD8⫹ T cells,
and these markers have been proposed to identify functionally
distinct T-cell populations and different stages of T-cell function.
A previous study indicated that HIV-specific CD8⫹ T cells appear
to have a different cell-surface phenotype from CMV-specific
CD8⫹ T cells,20 and the lack of perforin in HIV-specific populations
was correlated with decreased cytotoxic activity compared with
CMV-specific populations.34 In this study, we have also shown that
the cell-surface phenotypes of virus-specific CD8⫹ T cells are
distinct among HVCs and patients with HAM/TSP, irrespective of
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
2418
SABOURI et al
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
Figure 4. CD27 and CD28 coexpression on virus-specific CD8ⴙ T cells in HTLV-1 infection. Flow cytometric analysis of CD27 and CD28 coexpression gated on
HLA-A*02/Tax11-19 tetramer–positive cells. (A) The model of antigen-specific CD8⫹ T-cell differentiation based on the expression of CD27 and CD28. In the CD8⫹ T-cell
population, early differentiated cells differentiate into late-differentiated cells, following a stage of intermediate cells that have down-regulated CD28, but not yet CD27. (B) Dot
plots of HLA-A*02/Tax11-19 tetramer-PE (x axis) versus CD8-ECD fluorescence within the lymphocyte gate based on forward versus side scatter. (C) CD28/CD27 expression
on total CD8⫹ T-cell– and HTLV-1– or CMV-specific T cells detected by tetramers were shown. The HTLV-1 Tax–specific CD8 T cells were enriched in intermediatedifferentiated stage in HVCs and in late-differentiated stage in patients with HAM/TSP, whereas there was no difference in distribution of CMV pp65–specific CD8 T cells
between patients with HAM/TSP and HVCs. Representative dot plots from 1 HVC and 1 patient with HAM/TSP for HTLV-1–specific cells, and 1 HAM/TSP patient for
CMV-specific cells are shown. (D) Correlation between PVL (y axis; copy number of HTLV-1 tax per 104 PBMCs) and “early,” “intermediate,” or “late” phenotype of HTLV-1
Tax–specific cells (x axis; percentage of Tax-tetramer–positive cells in each subset) in PBMCs from HTLV-1–infected patients. Data were analyzed by Spearman rank
correlation.
PVL, indicating distinct characteristics of the anti–HTLV-1 immune response in infected patients with different disease status.
The observed increase in the frequencies of CD8⫹, CD8⫹CD28⫺,
and CD8⫹CD27⫺ T-cell population in PBMCs in HTLV-1 infection
without correlation with perforin or granzyme expression in the
corresponding cells suggests that the variation in perforin and
GzmB staining among different subjects with HTLV-1 infection
results from different levels of antigenic stimulation among these
hosts. In addition, the costimulatory molecule expression, perforin
content, and GzmB content might represent the state of activation
of these cells when viral antigen expression reaches equilibrium
with the CTL population. It is highly likely that this point of
equilibrium will differ not only between different viral infections
because of different kinetics of viral expression, but also between
patients with a different “efficiency” of the HTLV-1–specific CTL
response.
Finally, we performed a CD107a mobilization assay to compare
the functional reactivity of HTLV-1–specific CD8⫹ T cells between
patients with HAM/TSP and HVCs, because the CD107a is much
closer to the actual effector function than the equilibrium content of
perforin and GzmB. After cultivation with Tax11-19 peptide,
significantly higher anti-CD107a staining was observed in Taxtetramer–positive CD8⫹ T cells in HVCs than in patients with
HAM/TSP. This was also the case when we compared patients with
HAM/TSP and HVCs with a similar PVL, indicating that the
observed differences were not attributable to the difference in PVL
between patients with HAM/TSP and HVCs. Rather, the data
suggest that the CTLs in PBMCs from patients with HAM/TSP
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
HTLV-1–SPECIFIC CTLs IN CHRONIC INFECTION
2419
Table 3. Percentages of intermediate- and late-differentiated CD8ⴙ T cells based on the expression of CD28 and CD27 in HTLV-1 Tax– and
CMV pp65–specific CD8ⴙ T cells of patients with HAM/TSP and HVCs
Tax-specific
HAM/TSP
CMV-specific
Intermediate
Late
Intermediate
Late
38.1 ⫾ 5.2 (n ⫽ 16)
61.0 ⫾ 5.3 (n⫽16)
17.7 ⫾ 5.8 (n ⫽ 12)
81.9 ⫾ 5.8 (n ⫽ 12)
HVCs
66.3 ⫾ 3.8 (n ⫽ 17)
32.7 ⫾ 3.6 (n⫽17)
10.3 ⫾ 4.0 (n ⫽ 8)
89.0 ⫾ 2.4 (n ⫽ 8)
All
52.6 ⫾ 4.0 (n ⫽ 33)
46.4 ⫾ 4.0 (n⫽33)
14.8 ⫾ 3.6 (n ⫽ 20)
84.7 ⫾ 3.6 (n ⫽ 20)
P*
⬍ .001
⬍ .001
.88
.88
Tax-specific
Intermediate
Late
PVL
P*
PVL-matched, HAM/TSP
39.2 ⫾ 7.6 (n ⫽ 9)
56.6 ⫾ 7.9 (n ⫽ 9)
470.3 ⫾ 75.0
.11
PVL-matched, HVCs
68.2 ⫾ 4.9 (n ⫽ 10)
31.4 ⫾ 4.9 (n ⫽ 10)
303.5 ⫾ 70.0
PVL-matched, all
54.4 ⫾ 5.5 (n ⫽ 19)
44.8 ⫾ 4.9 (n ⫽ 19)
382.5 ⫾ 50.4
.004
.006
P*
Values represent the means plus minus SE.
*P values between HAM-HVCs were calculated by Mann-Whitney U test. PVL: HTLV-1 tax copy number per 104 PBMCs.
have discharged their granules much more recently because of the
greater antigenic stimulus, and so have fewer lytic granules and
less CD107a to discharge in the short-term in vitro assay. Because
there is evidence that the concentration of antigen required to
stimulate CD8⫹ T cells to produce cytokines is greater than the
concentration required to induce CTL killing of target cells,40
higher HTLV-1 PVL as well as higher antigen load in patients with
HAM/TSP may deteriorate the function of CD8⫹ T cells. In this
case, the abundance of antigen in patients with HAM/TSP might
exceed the threshold required to stimulate the CD8⫹ T cells to
produce inflammatory cytokines such as interferon-␥ (IFN-␥) and
tumor necrosis factor-␣ (TNF-␣); therefore, the effect of CTLs is
not protective but harmful for infected patients.
Figure 5. CD107a expression in patients with HAM/
TSP and in HVCs after coculture with immunodominant Tax peptide. To determine the functional reactivity
of HTLV-1 Tax–specific CD8⫹ T cells, we performed a
CD107a mobilization assay. PBMCs derived from HLAA*02–positive HTLV-1–infected patients (patients with
HAM/TSP and HVCs) were stained with anti–CD8-PC5
and HLA-A*02/Tax11-19 tetramer after incubation with
Tax11-19 peptide and anti-CD107a monoclonal antibody
for 4 hours. (A) Left panel: Dot plots of HLA-A*02/
Tax11-19 tetramer-PE (x axis) versus CD8-PC5 (y axis)
fluorescence within the lymphocyte gate based on forward versus side scatter. Significant anti-CD107a staining was observed on the Tax-tetramer–positive population after coculture with Tax11-19 peptide (left panel),
whereas only background staining was seen without
peptide (center panel). (B) Comparison of degranulation
marker CD107a expression in Tax-tetramer–positive cells
between 10 patients with HAM/TSP and 14 HVCs. The
percentage of CD107a⫹ Tax11-19 tetramer–positive cells/
Tax11-19 tetramer–positive cells was significantly decreased in patients with HAM/TSP compared with HVCs
(P ⫽ .003, Mann Whitney). (C) The CD107a staining was
still significantly lower in patients with HAM/TSP than in
HVCs compared between PVL-matched groups
(P ⫽ .006, Mann-Whitney). Horizontal bars represent the
median value of percent CD107a⫹ Tax11-19 tetramer–
positive cells/Tax11-19 tetramer–positive cells.
In conclusion, our present data indicate that CTL efficiency
appears to be lower in patients with HAM/TSP than in HVCs. The
different costimulatory molecule expression, perforin content, and
GzmB content between patients with HAM/TSP and HVCs may
reflect the different state of activation of the cells when viral
antigen expression reaches equilibrium with the CTL population
within infected patients. This point of equilibrium may differ not
only between different viral infections (ie, different kinetics of viral
expression), but also between HTLV-1–infected patients with a
different “efficiency” of CTL response. Future studies investigating
the nature of the defects in these T cells may provide opportunities
to overcome such inefficient functions, and may hold the potential
for reducing disease or eradicating persisting infection.
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
2420
BLOOD, 15 SEPTEMBER 2008 䡠 VOLUME 112, NUMBER 6
SABOURI et al
Acknowledgments
Authorship
We thank the staff and blood donors of Kagoshima University
Hospital. The authors thank Ms Fumiko Inoue and Yoko Nishino of
Kagoshima University and Ms Sumie Saito of Kanazawa Medical
University for their excellent technical assistance.
This work was supported by the Ministry of Health, Labor and
Welfare, Japan (Neuroimmunological Disease Research Committee Grant; S.I., M.O., and Y.O.); the Japan Society for the
Promotion of Science (JSPS; grant 17590886; M.S.); the Japan
Brain Foundation (M.S.); the Takeda Science Foundation (M.S.);
Kanazawa Medical University (grants S2006-1, H2007-11, and
C2007-5; M.S.). A.H.S. was supported by a Postdoctoral Fellowship for Foreign Researchers from JSPS.
Contribution: A.H.S. designed and performed the experiments,
analyzed the data, and wrote the paper; K.U. provided input into the
data analyses and interpretation; M.O. and D.H. provided samples,
clinical data, and advice; S.I., M.O., and Y.O. contributed to
obtaining funding and gave advice; M.S. designed and supervised
the research, performed experiments, and wrote the paper; and all
authors checked the final version of the manuscript.
Conflict-of-interest disclosure: The authors declare no competing financial interests.
Correspondence: Mineki Saito, Department of Microbiology,
Kanazawa Medical University, 1-1 Daigaku, Uchinada-machi,
Ishikawa 920-0293, Japan; e-mail: [email protected].
References
1. Poiesz BJ, Ruscetti FW, Gazdar AF, Bunn PA,
Minna JD, Gallo RC. Detection and isolation of
type C retrovirus particles from fresh and cultured
lymphocytes of a patient with cutaneous T-cell
lymphoma. Proc Natl Acad Sci U S A. 1980;77:
7415-7419.
2. Yoshida M, Miyoshi I, Hinuma Y. Isolation and
characterization of retrovirus from cell lines of
human adult T-cell leukemia and its implication in
the disease. Proc Natl Acad Sci U S A. 1982;79:
2031-2035.
3. Hinuma Y, Nagata K, Hanaoka M, et al. Adult Tcell leukemia: antigen in an ATL cell line and detection of antibodies to the antigen in human
sera. Proc Natl Acad Sci U S A. 1981;78:64766480.
4. Yoshida M, Seiki M, Yamaguchi K, Takatsuki K.
Monoclonal integration of human T-cell leukemia
provirus in all primary tumors of adult T-cell leukemia suggests causative role of human T-cell leukemia virus in the disease. Proc Natl Acad Sci
U S A. 1984;81:2534-2537.
5. Gessain A, Barin F, Vernant JC, et al. Antibodies
to human T-lymphotropic virus type-I in patients
with tropical spastic paraparesis. Lancet. 1985;2:
407-410.
6. Osame M, Usuku K, Izumo S, et al. HTLV-I associated myelopathy, a new clinical entity. Lancet.
1986;1:1031-1032.
7. Izumo S, Umehara F, Osame M. HTLV-Iassociated myelopathy. Neuropathology. 2000;
20(suppl):S65-68.
8. Uchiyama T. Human T cell leukemia virus type I
(HTLV-I) and human diseases. Annu Rev Immunol. 1997;15:15-37.
9. Nagai M, Usuku K, Matsumoto W, et al. Analysis
of HTLV-I proviral load in 202 HAM/TSP patients
and 243 asymptomatic HTLV-I carriers: high proviral load strongly predisposes to HAM/TSP.
J Neurovirol. 1998;4:586-593.
10. Jeffery KJ, Siddiqui AA, Bunce M, et al. The influence of HLA class I alleles and heterozygosity on
the outcome of human T cell lymphotropic virus
type I infection. J Immunol. 2000;165:7278-7284.
11. Jeffery KJ, Usuku K, Hall SE, et al. HLA alleles
determine human T-lymphotropic virus-I (HTLV-I)
proviral load and the risk of HTLV-I-associated
myelopathy. Proc Natl Acad Sci U S A. 1999;96:
3848-3853.
12. Hanon E, Hall S, Taylor GP, et al. Abundant tax
protein expression in CD4⫹ T cells infected with
human T-cell lymphotropic virus type I (HTLV-I) is
prevented by cytotoxic T lymphocytes. Blood.
2000;95:1386-1392.
13. Hanon E, Stinchcombe JC, Saito M, et al. Fratricide among CD8(⫹) T lymphocytes naturally infected with human T cell lymphotropic virus type I.
Immunity. 2000;13:657-664.
14. Kubota R, Kawanishi T, Matsubara H, Manns A,
Jacobson S. Demonstration of human T lymphotropic virus type I (HTLV-I) tax-specific CD8⫹ lymphocytes directly in peripheral blood of HTLV-I–
associated myelopathy/tropical spastic paraparesis patients by intracellular cytokine detection. J Immunol. 1998;161:482-488.
28.
15. Klenerman P, Hill A. T cells and viral persistence:
lessons from diverse infections. Nat Immunol.
2005;6:873-879.
29.
16. Vine AM, Heaps AG, Kaftantzi L, et al. The role of
CTLs in persistent viral infection: cytolytic gene
expression in CD8⫹ lymphocytes distinguishes
between individuals with a high or low proviral
load of human T cell lymphotropic virus type 1.
J Immunol. 2004;173:5121-5129.
17. Kubota R, Kawanishi T, Matsubara H, Manns A,
Jacobson S. HTLV-I specific IFN-gamma⫹ CD8⫹
lymphocytes correlate with the proviral load in
peripheral blood of infected individuals. J Neuroimmunol. 2000;102:208-215.
18. Wodarz D, Hall SE, Usuku K, et al. Cytotoxic Tcell abundance and virus load in human immunodeficiency virus type 1 and human T-cell leukaemia virus type 1. Proc Biol Sci. 2001;268:12151221.
19. Goon PK, Biancardi A, Fast N, et al. Human T cell
lymphotropic virus (HTLV) type-1–specific CD8⫹
T cells: frequency and immunodominance hierarchy. J Infect Dis. 2004;189:2294-2298.
30.
31.
32.
33.
20. Appay V, Dunbar PR, Callan M, et al. Memory
CD8⫹ T cells vary in differentiation phenotype in
different persistent virus infections. Nat Med.
2002;8:379-385.
34.
21. Harari A, Vallelian F, Meylan PR, Pantaleo G.
Functional heterogeneity of memory CD4 T cell
responses in different conditions of antigen exposure and persistence. J Immunol. 2005;174:
1037-1045.
35.
22. Betts MR, Nason MC, West SM, et al. HIV nonprogressors preferentially maintain highly functional HIV-specific CD8⫹ T cells. Blood. 2006;
107:4781-4789.
23. Osame M. Review of WHO Kagoshima meeting
and diagnostic guidelines for HAM/TSP. New
York, NY: Raven Press; 1990.
24. Rubio V, Stuge TB, Singh N, et al. Ex vivo identification, isolation and analysis of tumor-cytolytic T
cells. Nat Med. 2003;9:1377-1382.
25. Burkett MW, Shafer-Weaver KA, Strobl S,
Baseler M, Malyguine A. A novel flow cytometric
assay for evaluating cell-mediated cytotoxicity.
J Immunother (1997). 2005;28:396-402.
26. Gillespie GM, Wills MR, Appay V, et al. Functional
heterogeneity and high frequencies of cytomegalovirus-specific CD8(⫹) T lymphocytes in healthy
seropositive donors. J Virol. 2000;74:8140-8150.
27. Callan MF, Tan L, Annels N, et al. Direct visualiza-
36.
37.
38.
39.
40.
tion of antigen-specific CD8⫹ T cells during the
primary immune response to Epstein-Barr virus in
vivo. J Exp Med. 1998;187:1395-1402.
Yasunaga J, Sakai T, Nosaka K, et al. Impaired
production of naive T lymphocytes in human Tcell leukemia virus type I–infected individuals: its
implications in the immunodeficient state. Blood.
2001;97:3177-3183.
Asquith B, Zhang Y, Mosley AJ, et al. In vivo T
lymphocyte dynamics in humans and the impact
of human T-lymphotropic virus 1 infection. Proc
Natl Acad Sci U S A. 2007;104:8035-8040.
Choremi-Papadopoulou H, Panagiotou N,
Samouilidou E, et al. CD28 costimulation and
CD28 expression in T lymphocyte subsets in
HIV-1 infection with and without progression to
AIDS. Clin Exp Immunol. 2000;119:499-506.
Vingerhoets JH, Vanham GL, Kestens LL, et al.
Increased cytolytic T lymphocyte activity and decreased B7 responsiveness are associated with
CD28 down-regulation on CD8⫹ T cells from HIVinfected subjects. Clin Exp Immunol. 1995;100:
425-433.
Zhang D, Shankar P, Xu Z, et al. Most antiviral
CD8 T cells during chronic viral infection do not
express high levels of perforin and are not directly
cytotoxic. Blood. 2003;101:226-235.
Wherry EJ, Ahmed R. Memory CD8 T-cell differentiation during viral infection. J Virol. 2004;78:
5535-5545.
Appay V, Nixon DF, Donahoe SM, et al. HIVspecific CD8(⫹) T cells produce antiviral cytokines but are impaired in cytolytic function. J Exp
Med. 2000;192:63-75.
Haridas V, McCloskey TW, Pahwa R, Pahwa S.
Discordant expression of perforin and granzyme
A in total and HIV-specific CD8 T lymphocytes of
HIV infected children and adolescents. AIDS.
2003;17:2313-2322.
Chen G, Shankar P, Lange C, et al. CD8 T cells
specific for human immunodeficiency virus, Epstein-Barr virus, and cytomegalovirus lack molecules for homing to lymphoid sites of infection.
Blood. 2001;98:156-164.
Sandberg JK, Fast NM, Nixon DF. Functional heterogeneity of cytokines and cytolytic effector molecules in human CD8⫹ T lymphocytes. J Immunol. 2001;167:181-187.
Klingstrom J, Gudmundsdotter L, Zuber B, et al.
Elevated levels of serum perforin in chronic HIV-1
and acute SIV/SHIV infection. AIDS. 2006;20:
125-127.
Bangham CR. The immune control and cell-tocell spread of human T-lymphotropic virus type 1.
J Gen Virol. 2003;84:3177-3189.
Valitutti S, Muller S, Dessing M, Lanzavecchia A.
Different responses are elicited in cytotoxic T lymphocytes by different levels of T cell receptor occupancy. J Exp Med. 1996;183:1917-1921.
From www.bloodjournal.org by guest on June 15, 2017. For personal use only.
2008 112: 2411-2420
doi:10.1182/blood-2008-02-140335 originally published
online May 23, 2008
Impaired function of human T-lymphotropic virus type 1 (HTLV-1)−
specific CD8+ T cells in HTLV-1−associated neurologic disease
Amir H. Sabouri, Koichiro Usuku, Daisuke Hayashi, Shuji Izumo, Yoshiro Ohara, Mitsuhiro Osame
and Mineki Saito
Updated information and services can be found at:
http://www.bloodjournal.org/content/112/6/2411.full.html
Articles on similar topics can be found in the following Blood collections
Immunobiology (5489 articles)
Information about reproducing this article in parts or in its entirety may be found online at:
http://www.bloodjournal.org/site/misc/rights.xhtml#repub_requests
Information about ordering reprints may be found online at:
http://www.bloodjournal.org/site/misc/rights.xhtml#reprints
Information about subscriptions and ASH membership may be found online at:
http://www.bloodjournal.org/site/subscriptions/index.xhtml
Blood (print ISSN 0006-4971, online ISSN 1528-0020), is published weekly by the American Society
of Hematology, 2021 L St, NW, Suite 900, Washington DC 20036.
Copyright 2011 by The American Society of Hematology; all rights reserved.