A role for HVEM, but not lymphotoxin-beta receptor, in LIGHT-induced tumor cell death and chemokine...
Transcript of A role for HVEM, but not lymphotoxin-beta receptor, in LIGHT-induced tumor cell death and chemokine...
A role for HVEM, but not lymphotoxin-b receptor, inLIGHT-induced tumor cell death and chemokineproduction
Christine Pasero1, Bernadette Barbarat1,2, Sylvaine Just-Landi1,2,
Alain Bernard3, Therese Aurran-Schleinitz4, Jerome Rey1,2,4,
Eric Eldering5, Alemsedeg Truneh6, Regis T Costello7 and Daniel Olive1,2
1 INSERM UMR891, Centre de Recherche en Cancerologie de Marseille, Universite de la
Mediterranee, Institut de Cancerologie et d’Immunologie de Marseille, Marseille, France2 Laboratoire d’Immunologie des Tumeurs, Institut Paoli-Calmettes,, Marseille, France3 INSERM UMR576, Hopital de l’Archet, Nice, France4 Departement d’Hematologie, Institut Paoli-Calmettes, Marseille, France5 Department of Experimental Immunology, Academic Medical Center, Amsterdam,
The Netherlands6 Synaptex Inc, Boston, MA, USA7 Departement d’Hematologie, Hopital Nord, Marseille, France
The TNF member LIGHT also known as TL4 or TNFSF14) can play a major role in cancer
control via its two receptors; it induces tumor cell death through lymphotoxin-b receptor
(LT-bR) and ligation to the herpes virus entry mediator (HVEM) amplifies the immune
response. By studying the effect of LIGHT in the transcriptional profile of a lymphoid
malignancy, we found that HVEM, but not LT-bR, stimulation induces a significant increase
in the expression of chemokine genes such as IL-8, and an unexpected upregulation of
apoptotic genes. This had functional consequences, since LIGHT, or HVEM mAb, thus far
known to costimulate T- and B-cell activation, induced chronic lymphocytic leukemia cell
death. Many of the mediators involved were identified here, with an apoptotic pathway as
demonstrated by caspases activation, decrease in mitochondrial membrane potential,
upregulation of the pro-apoptotic protein Bax, but also a role of TRAIL. Moreover, HVEM
induced endogenous TNF-a production and TNF-a enhanced HVEM-mediated cell death.
HVEM function was mainly dependent on LIGHT, since other ligands like HSV-glycoprotein
D and B and T lymphocyte attenuator were essentially ineffective. In conclusion, we
describe a novel, as yet unknown killing effect of LIGHT through HVEM on a lymphoid
malignancy, and combined with induction of chemokine release this may represent an
additional tool to boost cancer immunotherapy.
Key words: Cell death . Chronic lymphocytic leukemia . LIGHT . HVEM . TNF
Supporting Information available online
Correspondence: Professor Daniel Olivee-mail: [email protected]
& 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.eji-journal.eu
DOI 10.1002/eji.200939069 Eur. J. Immunol. 2009. 39: 2502–2514Christine Pasero et al.2502
Introduction
The TNF and TNF receptor (TNFR) superfamilies have diverse and
widespread physiological functions, including apoptosis, proliferation,
differentiation, and immune system regulation. TNFR are schemati-
cally classified into two groups: those with a 80-amino acids death
domain (DD) in the cytoplasmic region of the receptor, which signals
cell death, and those without DD, which have been implicated in
lymphoid cell proliferation and differentiation [1, 2]. LIGHT also
known as TL4 or TNFSF14) is a glycosylated, type II transmembrane
protein of about 29kDa, which can bind lymphotoxin-b receptor
(LT-bR) and the herpes virus entry mediator (HVEM) both of which
belong to the TNFR superfamily. HVEM is a type I transmembrane
protein of 32 to 36kDa, devoid of intracytoplasmic DD. To date, five
ligands have been identified for HVEM. The first one is HSV
glycoprotein D (HSV-gD), a structural component of the HSV
envelope, essential for HSV entry into host cells [3]. Two of the
ligands are members of the TNF superfamily: the trimeric lymphotoxin
a (LTa3) and LIGHT [4]. Recently, it was also found that HVEM
interacts with two immunoglobulin superfamily members B and T
lymphocyte attenuator (BTLA) and CD160 [5–7]. HVEM is expressed
in most immune cells such as monocytes, T and B lymphocytes, and
natural killer cells whereas expression of LT-bR is restricted to stromal
cells or non-lymphoid hematopoietic cells [8, 9]. Functional
differences between the two receptors of LIGHT have also been
observed. LIGHT binding to HVEM confers mainly costimulatory
effects; LIGHT has a potent, CD28-independent, co-stimulatory role in
initial T-cell priming and expansion. Blockade of endogenous LIGHT
efficiently prevents the development of T-cell-mediated graft-versus-
host disease [10–12, 13]. Moreover, LIGHT and CD40L act
synergistically to induce dendritic cell maturation, LIGHT protein
induction at the surface of B lymphocytes, and B-cell proliferation
[14–17]. In contrast, biological consequences of LIGHT/LTbR
signaling include apoptosis of adenocarcinoma when combined with
IFN-g [18, 19], but also organization and maintenance of lymphoid
structures [20, 21], partly because of its ability to induce the
expression of various chemokines and adhesion molecules [22]. Yu et
al. have described an anti-tumor potential for LIGHT in a murine in
vivo model [23]. Forced expression of LIGHT inside the tumor, by
transducing a stable transmembrane form of LIGHT into a mouse
fibrosarcoma, led to eradication of established tumor cells. This in vivo
model showed that LIGHT could be an attractive target for anti-tumor
response. Some other TNF superfamily members devoid of DD, e.g.
OX40, 4-1BB, and CD27, have also been shown to efficiently induce
anti-tumor responses by their properties of costimulation [24–26].
Hematopoietic malignancies are life-threatening afflictions.
Despite recent progress in their treatment with the use of mAb,
such as rituximab and alemtuzumab [27, 28], failure to obtain
complete remission and the high potential for relapse impair
patient prognosis. It is therefore important to continue to search
for new therapeutic approaches.
The anti-tumor role of LIGHT in solid tumors prompted us to
investigate its role in hematopoietic malignancies. Given the
strong expression of HVEM in chronic lymphocytic leukemia
(CLL) cells and our previous experience on the effects of LIGHT in
both normal [17] and mantle-cell lymphoma [29] B lymphocytes,
we focused our attention on the analysis of the effect of LIGHT
stimulation on B-CLL cells. Here, we studied the effect of LIGHT
and HVEM first on the transcriptional regulation of a number of
genes coding for chemokines, chemokine receptors, cytokines,
and molecules involved in adhesion, migration, or apoptosis.
Then, we examined LIGHT and HVEM effects in vitro in B-CLL
cells, in order to determine if they could be effective candidates
for anti-tumor immunotherapy approaches.
Results
Expression of HVEM and LT-bR on hematopoieticmalignancies
We and others had previously shown that HVEM was expressed in
lymphoid malignancies, particularly on leukemias of B origin,
with high intensity in all the CLL and all the mantle cell
lymphomas (MCL) tested, and often observed in acute lympho-
blastic leukemias [29, 30]. As shown in Fig. 1A, we confirmed
that HVEM was strongly expressed in all CLL and MCL samples
tested, whereas HVEM expression was weak in leukemias of
myeloid origin (acute myeloid leukemias, AML). In sharp
contrast, LT-bR was expressed in AML, but absent in MCL and
infrequent or low in CLL. The expression of LT-bR had already
been described in the K562 cell line, derived from a chronic
myelogenous patient, and in HL60 cell line, derived from an AML
patient [9], but not yet in primary AML cells. As seen in Fig. 1B,
HVEM was expressed at uniformly high levels in the CLL tested,
and LT-bR was either expressed weakly or not expressed.
Figure 1. Expression of HVEM and LT-bR on hematopoietic malignan-cies. (A) Surface expression of HVEM and LT-bR on CLL, MCL, and AMLwas monitored by flow cytometry. Represented are one of nine CLLpatients, one of two MCL patients, and one of two AML patients. Theopen histograms represent the fluorescence of the cells stained withisotype-matched control mAb of irrelevant specificity. The filledhistograms represent staining with specific HVEM-FITC or LT-bR-PEconjugated mAb. (B) The bars represent the ratio of MFI of stainedantibody for HVEM or LT-bR compared with isotype-matched controlperformed on nine B-CLL patients. The error bars indicate the SEM.
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HVEM stimulation induces over-expression of 12chemokine and apoptotic genes
B-CLL was therefore a study model with a predominant
expression of HVEM and a weak or absent LT-bR expression.
Therefore, to explore the impact of LIGHT on this lymphoid
malignancy, we stimulated B-CLL cells with an anti-HVEM mAb
for 24 h. After RNA preparation, we performed real-time
quantitative PCR (QRT-PCR) global analysis of the expression
of 322 genes coding for chemokines, chemokine receptors,
cytokines, and molecules involved in essential mechanisms of
the cells such as adhesion, migration, or apoptosis. As seen in
Fig. 2 and in the Supporting Information tables, among these 322
genes tested, our results showed that 21 genes were not
expressed at all in the CLL patient samples tested. The 301
expressed genes were either barely transcribed for 80 of them or
expressed to normal or high levels for 221 others. Among the 322
genes tested, only 12 were significantly over-expressed following
HVEM stimulation, suggesting a restricted transcriptional
response mediated via HVEM in B-CLL cells (Fig. 2A). HVEM
stimulation significantly upregulated the expression of two genes
coding for chemokines: IL-8, IFN-inducible protein 10 (IP10),
and one chemokine receptor, Chemokine (C-C motif) receptor 4,
po0.01. Surprisingly, the only nine other genes significantly
induced after HVEM stimulation all correspond to genes coding
for apoptotic proteins (Fig. 2A), most of them pro-apoptotic: Bcl-
XS, Bid, FasL (po0.01), BNIP3, CARD11, Cytochrome c, p53
(po0.05), and other genes that belong to the anti-apoptotic
subgroup: BIRC4 and IEX-1 (po0.05). Fig. 2B represents the
RNA fold induction for the 12 over-expressed genes following
HVEM stimulation, depicted as the mean7SEM for seven
different patients (n 5 11 for Fas-L). IL-8, IP10, and FasL were
the most increased genes, with six-, nine-, and fivefold increase,
respectively. Except for BIRC4 for which the ratios were more
Figure 2. Genes significantly induced after HVEM stimulation in B-CLL cells among the 322 genes tested by QRT-PCR RNA was quantified by QRT-PCR(as described in the Materials and methods) in B-CLL cells treated or untreated with anti-HVEM mAb for 24 h. RNA expression was normalized usingb-actin as an endogenous control, and DCt value was calculated with DCt 5 Ct target–Ct endogenous control. (A) RNA induction levels of the 12 over-expressed genes following HVEM stimulation. Significant difference between the DCt for the HVEM mAb condition and the control condition wastested with the Wilcoxon signed-rank test, and p values of each gene significantly induced are presented in this table, with ��po0.01 and �po0.05,(n represents number of patients; ns : non-significant; nt: not tested). Protein induction levels are shown for IL-8 and Rantes. (B) Data were thenconverted to a fold change ratio as described in the Materials and methods. Ratios between 1 and 1.5 correspond to weakly over-expressed genes andratios over 1.5 correspond to strongly over-expressed genes. Data represent the mean7SEM for seven different patients (except for Fas-L n 5 11). (C)Rantes and IL-8 production were determined by Cytometric Bead Array or ELISA (as described in the Materials and methods) in supernatants of B-CLLcells treated or untreated with anti-HVEM mAb for 24 h. The left panel depicts the mean Rantes secretion observed for nine different patients7SEM(��po0.01, Wilcoxon test), and the right panel depicts the mean IL-8 secretion for 12 different patients7SEM (���po0.001, Wilcoxon test).
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variable, the over-expression observed for the others genes were
due to a homogeneous shift in expression.
We measured the chemokine production in stimulated B-CLL
cells by Cytometric Bead Array or ELISA (Fig. 2C). We observed
that HVEM induced a significant increase in the IL-8 secretion
compared with control condition (18657700 pg/mL versus
3377120 pg/mL, respectively; with a 6.1 fold increase mean) in
12 different patients (po0.001). HVEM stimulation also
increased significantly the regulated on activation, normal T-cell
expressed and secreted (Rantes) production, with 66736 pg/mL
in the control condition and 102751 pg/mL in the treated
condition (1.6 fold increase) in nine patients (po0.01). No
significant increase in the release of IP10 was observed.
LIGHT and HVEM mAb induces death of B-CLL cells
We sought to determine if this apoptotic transcriptional profile
was correlated with a function in vitro. First, B-CLL cells were
cocultured with LIGHT-expressing cells or control cells for 24 h;
then the percentage of apoptotic cells was measured by Annexin
V/PI double staining. Apoptotic cells include Annexin V1/ PI�
(early apoptotic) and Annexin V1/PI1(late apoptotic/necrotic)
populations. We found that stimulation by LIGHT effectively
induced cell death of 41% of the B-CLL cells compared with 8%
with the control condition (spontaneous cell death) on this
representative experiment (Fig. 3A). These results were reproduced
in 15 B-CLL samples. Then, we stimulated leukemic cells with HVEM
mAb for 24 h. We had previously shown that this antibody blocks the
interaction between HVEM and its ligand LIGHT [14]. Figure 3B
shows that both LIGHT and anti-HVEM mAb were able to induce
CLL cell death, with 4273 and 5273% dead cells, respectively,
compared with 2272% in the control condition, corresponding to
highly statistically significant (po0.001) 1.9- and 2.4-fold increase.
Of note, in these experiments, LIGHT or anti-HVEM mAb-induced
CLL cell death did not require priming of the cells with IFN-g (data
not shown).
Anti-HVEM and cross-linked anti-CD20 mAb were compared
for their effectiveness in inducing CLL cell death (Fig. 3B). The
HVEM mAb-induced CLL cell death was comparable with that of
the pan-B cell mAb rituximab, with 5273 and 46.276% of dead
cells, respectively.
In the recent past, several ligands were identified for HVEM
including LIGHT, but also HSV-gD and BTLA. To compare the
efficacy of the different HVEM ligands to the efficacy of HVEM
mAb, B-CLL cells were cocultured with LIGHT-, HSV-gD, BTLA-
transfected cells, or control cells for 24 h; then the percentage
of dead cells was measured by Annexin V/PI double staining.
Figure 3. LIGHT and HVEM mAb induce death of B-CLL cells (A) B-CLL cells were incubated with irradiated (50 Gy) CD32-transfected (control) orLIGHT-transfected L-cells at a ratio of one transfectant for ten leukemic cells. After 24 h of incubation, the cells were analyzed by flow cytometrywith Annexin V/PI double staining as described in the Materials and methods. The dead cells include Annexin V1/PI� (early apoptotic) and AnnexinV1/PI1 (late apoptotic/necrotic) cells. The figure shows one representative experiment out of 15 performed. (B) B-CLL cells were incubated witheither irradiated (50 Gy) CD32-transfected (control) or LIGHT-transfected L-cells or were treated with either 30mg/mL of anti-HVEM mAb or with10 mg/mL therapeutic anti-CD20 (Rituximab) mAb cross-linked with goat anti-human antisera, and analyzed by Annexin V/PI staining. The barsdepict the mean percentage of dead cells observed7SEM, with n 5 25 for control and HVEM mAb, n 5 15 for LIGHT-transfected cells, and n 5 5 forRituximab �po0.05; ���po0.001 compared with control condition. (C) Effects of HVEM ligand, as compared with HVEM mAb, stimulation. B-CLLcells were cocultured with irradiated CD32� (control), CD40L�, LIGHT�, CD40L/LIGHT�, gD, or BTLA-transfected L-cells for 24 h or treated withanti-HVEM mAb (30 mg/mL), and analyzed by Annexin V/PI staining. The bars depict the mean percentage of dead cells observed7SEM, with n 5 6per treatment. Statistical significance in each condition was calculated versus the control condition with a Wilcoxon test as indicated in theMaterials and methods section (�po0.05; ns, not significant).
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CD40L-transfected cells were used as a ‘‘non-ligand’’ negative
control. As seen in Fig. 3C, LIGHT was almost as efficient as HVEM
mAb (35.274% of dead cells, po0.05 and 4274%, po0.05,
respectively). HSV-gD had a weak effect (27.473%, po0.05),
while BTLA had no significant effect (24.476%) compared with
control condition (17.773%). As previously reported, CD40L did
not induce B-CLL cell death (22.672%). Finally, we also tested
CD40L/LIGHT costimulation because we had previously shown
that these two molecules act in cooperation for B-lymphocyte
proliferation [17]. Here, CD40L/LIGHT costimulation gave a
similar effect to LIGHT alone; thus neither does CD40L appear to
cooperate with LIGHT to induce cell death nor hinders it.
HVEM-mediated cell death involves caspases 3, 8, and9 activation
We observed (Fig. 4A) that stimulation of B-CLL cells by anti-
HVEM mAb results in statistically significant caspase 3 activation
(16.572% positive cells, HVEM mAb condition versus 5.271%,
control condition, po0.01). This observation was confirmed by
Western blotting analysis, with an increase of the 19 and 17 kDa
active forms (Fig. 4B). Figure 4A also shows that caspase 3
activation induced by HVEM mAb is completely abrogated by pre-
treatment with the pan-caspase inhibitor Z-VAD-FMK (471%
positive cells, left panel). In contrast, pre-treatment with Z-VAD-
FMK only partially blocked the Annexin V/PI staining observed
after HVEM stimulation (data not shown). In addition, we
analyzed by flow cytometry (Fig. 4C) the activation of the
effector caspases 8 and 9. Both caspases 8 and 9 were activated in
response to treatment of CLL cells by anti-HVEM mAb, with
similar kinetics: 071.5% (t 5 3 h), 5.472.5% (t 5 6 h),
1673.2% (t 5 12 h), 24.873.1 (t 5 24 h) for caspase 8, and
0.8872.1% (t 5 3 h), 4.772.6% (t 5 6 h), 18.474.1% (t 5 12 h),
31.475.1% (t 5 24 h) for caspase 9.
HVEM disrupts the mitochondrial membrane potentialand increases Bax expression
Figure 5A shows that treatment of B-CLL cells with anti-HVEM
mAb resulted in a loss of the mitochondrial membrane potential,
as assessed by the significant decrease of the DiOC2(3) green
fluorescence (7078% in activated cells with resting cells set to
Figure 4. HVEM mAb induces activation of caspases 3, 8, and 9. (A) B-CLL cells were pre-treated with 20mM of the pan-caspase inhibitor Z-VAD-FMK for 30 min at 371C, and then either left untreated or stimulated with anti-HVEM mAb. Untreated B-CLL cells served as a control. After 24 h,cells were collected, permeabilized, and stained for active caspase 3 (left panel). Data represent the mean percentage of cells positive for activecaspase 37SEM (seven independent experiments). In parallel, cells of each condition were analyzed with Annexin V/PI double staining (rightpanel). The bars depict the mean percentage of dead cells observed7SEM (seven independent experiments). Statistical significance in both panelswas calculated versus the control condition with a Wilcoxon test: �po0.05; ��po0.01. (B) B-CLL cells from two different patients (UPN10 and 11)were either left untreated or stimulated with anti-HVEM mAb. After 24 h of incubation, cells lysates were prepared and immunoblotted with mAbrecognizing cleaved caspase 3 or b-actin (to control for protein loading). Jurkat cells unstimulated or treated with the anti-Fas mAb CH11 were usedas positive control. As described in the Materials and methods, each band was quantified and normalized to correct for RNA loading. Ratios ofstimulated condition versus unstimulated condition for the 19 and 17 kDa bands are, respectively: JA16 (1.35 and 11 (the 17 kDa band in the controlcondition was not detected and the ratio not quantified)), UPN10 (0.76 and 1.57) and UPN11 (1.2 and 1.73). (C) B-CLL cells were either left untreatedor stimulated with anti-HVEM mAb, then incubated with a specific caspase 8 or 9 fluorescent inhibitor at the indicated time points for 1 h at 371C,and before flow cytometric analysis. Data represent the mean percentage of cells positive for active caspase 8 or 9 (after subtraction of thebackground corresponding to the untreated condition)7SEM observed for cells from six different patients.
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100%, po0.05). Interestingly, the HVEM-mediated mitochon-
drial depolarization was not inhibited by the pan-caspase
inhibitor Z-VAD-FMK (7175% DiOC2(3) positive cells), suggest-
ing that this step is independent of caspase activation.
The balance between Bax and Bcl-2 proteins is important for
the maintenance of mitochondrial activity. Using flow cytometry
analysis, we show that under resting conditions, Bcl-2 was highly
expressed in B-CLL cells as it was described in previous studies
[31]. HVEM stimulation did not change Bcl-2 expression, but in
contrast, induced a strong significant increase in Bax cytosolic
levels: 3974% positive cells, HVEM treated condition compared
with 2272%, resting condition (Fig. 5B).
HVEM-cell death increases FADD expression andpartially depends on TRAIL pathway
Based on the known mechanisms of apoptosis, we analyzed
the expression of Fas-associated death domain (FADD)
containing protein, the major adaptor molecule involved
in Fas and TNFR-mediated cell death [32]. Interestingly,
treatment of B-CLL cells with anti-HVEM mAb caused a
major increase in FADD expression by Western blotting analysis
(Fig. 6A). The stimulation caused substantial induction
of the 27 kDa band in CLL samples from two different
patients (2.39-fold increase for UPN10 and fivefold for
UPN11).
Then we examined whether HVEM-induced cell death
could be mediated indirectly through the activation of other
TNFR/ligand systems. We analyzed B-CLL cells for expression of
death receptors and their ligands (Fas and FasL, DR4-DR5, and
TRAIL) before and after HVEM stimulation (Fig. 6B). The
expression level of FasL and DR4 was weak, and DR5
was not detected on each of the samples (n 5 5) before stimula-
tion. Expression of FasL, DR4, and DR5 did not change after
HVEM stimulation (data not shown). In contrast, HVEM
ligation induced a significant increase in TRAIL expression with a
pre-treatment mean MFI value of 179758 to 375781 after
treatment (po0.05), and a non-significant increase of Fas, with
pre-treatment mean MFI 70722 to 104724 after HVEM stimu-
lation. Of note, TRAIL mRNA was among the 322 genes that
we tested by QRT-PCR and although the TRAIL gene was
expressed in CLL, it was not significantly increased follow-
ing HVEM stimulation, suggesting a post-transcriptional
modification.
To further address the role of FasL and TRAIL in HVEM-
mediated cell death, we generated artificial cytotoxic effectors
cells from Cos cells that expressed FasL, TRAIL, or both FasL and
TRAIL. Expression of both FasL and TRAIL following transient
transfection was verified by flow cytometry (data not shown). In
addition, the transfected effector cells were examined for their
capacity to induce apoptosis of Jurkat T cells. We cocultured each
of Cos-effector cell populations with B-CLL cells in the presence
or absence of HVEM mAb for 24 h (Fig. 6C). We found that FasL,
TRAIL, combined FasL and TRAIL induced apoptosis of 47710,
2577, 45717% of B-CLL cells, respectively, compared with
control condition, within 24 h of coculture. Combination of
HVEM mAb with FasL effector cells induced killing of B-CLL cells
that was higher than that observed with HVEM mAb alone
(80720 and 5575%, respectively). In contrast, combination of
TRAIL effector cells with HVEM mAb did not induce more
apoptosis than HVEM mAb alone (53716% compared with
5575%, respectively).
In addition, we examined the effect of blocking the FasL or the
TRAIL pathways on HVEM-induced cell death. B-CLL cells were
pre-incubated with a blocking anti-TRAIL mAb or Fas-Ig protein,
or not, then treated with HVEM mAb within 24 h and evaluated
for Annexin V/PI staining. We found, as shown in Fig. 6D, that
HVEM killing of CLL was significantly blocked by anti-TRAIL
mAb, partially for seven patients and totally for one patient tested
(5279% of dead cells in blocking condition versus 100%
in HVEM mAb condition; po0.01). In contrast, the pre-incuba-
tion with Fas-Ig protein had no effect on HVEM-mediated cell
death.
Figure 5. HVEM mAb disrupts the mitochondrial membrane potentialand increases Bax expression. (A) B-CLL cells were pretreated with20 mM of the pan-caspase inhibitor Z-VAD-FMK for 30 min at 371C, andthen either left untreated or stimulated with anti-HVEM mAb.Untreated B-CLL cells served as a control. After 24 h of incubation,cells were collected, incubated with 50 nM of DiOC2(3) for 30 min at371C, and analyzed for the loss of mitochondrial membrane potential(Dcm) by flow cytometry detecting DiOC2(3) fluorescence. Datarepresent the mean percentage of cells positive for DiOC2(3) greenfluorescence7SEM for cells from six different patients, with thepercentage of DiOC2(3) green fluorescence in the control conditionset to 100%. �po0.05 with a Wilcoxon test. The positive control was thedepolarizing agent carbonyl cyanide m-chloro phenyl hydrazone(4%71% cells positive for DiOC2(3), data not shown). (B) B-CLL cellswere either left untreated or stimulated with anti-HVEM mAb. After24 h of incubation, cells were collected, permeabilized, and stainedwith anti-Bax or anti-Bcl-2 mAb, and analyzed by flow cytometry.Results are expressed as the mean percentage of positive cells (aftersubtraction of the background corresponding to the isotype-matchedcontrol)7SEM observed for six different patients, �po0.05 with aWilcoxon test.
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HVEM induces TNF-a production: synergization onHVEM-cell death
Grell et al. demonstrated that cytotoxic effects induced by some
members of the TNFR family that lack DD, like TNF-R2, CD40, or
CD30, were mediated by endogenous production of TNF and
autotropic or paratropic activation of TNF-R1. Here, we cocultured
B-CLL cells with LIGHT-transfected cells or control cells for 24h, in
the presence or absence of anti-TNF-a mAb and we analyzed the
supernatants for TNF production by ELISA assay. We observed that
LIGHT significantly increased TNF production compared with control
condition (Fig. 7A), with 35710pg/mL compared with 2176pg/
mL, respectively (po0.05); and this secretion was blocked when
B-CLL cells were cultured in the presence of anti-TNF mAb (data not
shown). Then, we cultured B-CLL cells with LIGHT-transfected cells
or control cells for 24h, in the presence or absence of TNF-a, and the
percentage of dead cells was measured by Annexin V/PI double
staining. We found that TNF-a enhanced HVEM-mediated death,
with a mean increase of 38% (Fig. 7B). The difference between
LIGHT1TNF-a condition versus LIGHT condition was statistically
significant, with po0.01. Notably, addition of antibodies to TNF
could not block HVEM-induced cell death, indicating that induction
of TNF-a is not essential for HVEM killing, but that TNF- produced by
HVEM triggering could amplify HVEM-induced cell death (data not
shown)
Finally, in order to study conditions that might mimic the status of
primary CLL in their bone marrow and lymph node niches, we
performed experiments where CLL cells were stimulated with CD40L.
In brief, B-CLL cells were pre-activated with CD40L-transfected cells
or control cells for 24h. Then, cells were treated with anti-Fas mAb
(CH11 clone), anti-HVEM mAb, or left untreated for 24h, and eval-
uated for Annexin V/PI staining. As shown in Fig. 8, we observed that
CD40L pre-activation sensitizes the cells to Fas killing. In contrast,
HVEM-mediated cell death was not affected by CD40L pre-activation.
Figure 6. HVEM-cell death increases FADD expression and partially depends on the TRAIL pathway. (A) B-CLL cells from two different patients(UPN10 and 11) were either left untreated or stimulated with anti-HVEM mAb. After 24 h of incubation, cell lysates were prepared andimmunoblotted for FADD or b-actin (to control for protein loading). The TF1 erythroleukemic cell line was used as a positive control. As describedin the Materials and methods, ratios of stimulated condition versus unstimulated condition for the 27 kDa band were calculated (2.39 for UPN10 and 5for UPN11). (B) Surface expression of Fas and TRAIL was analyzed by flow cytometry in B-CLL cells either untreated (top row) or stimulated withHVEM mAb for 24 h (bottom row). The open histograms represent the fluorescence of the cells stained with isotype-matched control mAb ofirrelevant specificity. The filled histograms represent staining with specific Fas-FITC (left column) or TRAIL-PE (right column) conjugated mAb. Thefigure shows one representative experiment out of five performed. Mean MFI for Fas: 70722 in control condition versus 104724 in HVEM mAbcondition (n 5 5, ns). Mean MFI for TRAIL: 179758 in control condition versus 375781 in HVEM mAb condition (n 5 5, �po0.05 with a Wilcoxon test).(C) Cos cells were transfected transiently with FasL, TRAIL, or both for 24 h, then B-CLL cells were added to the culture, in the presence or absenceof HVEM mAb for 24 h. Cells were then collected and Annexin V/PI staining detected by flow cytometry. Data are presented as fold inductioncalculated as (percent apoptosis of treatment–percent apoptosis of untreated cells) divided by percent apoptosis of untreated cells � 100 for threedifferent patients. (D) B-CLL cells were left untreated or stimulated with HVEM mAb, in the presence or absence of 100 ng/mL of the blocking anti-TRAIL mAb RIK-2, or Fas-Ig protein for 24 h. Cells were then analyzed by flow cytometry for Annexin V/PI double staining. Data represent the meanpercentage of dead cells7SEM for cells from eight different patients, with cell death induced in the HVEM mAb condition set to 100%. Cell deathinduced in the blocking condition was then calculated relatively as: (percent apoptosis in the blocking condition divided by percent apoptosis ofHVEM mAb � 100). Statistical significance in each condition was calculated versus the HVEM mAb condition, ��po0.01 with a Wilcoxon test.
Eur. J. Immunol. 2009. 39: 2502–2514Christine Pasero et al.2508
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Discussion
In this study, we describe a novel potential of HVEM and LIGHT
on anti-tumor control. In 2004, Yu et al. reported that
transducing a transmembrane form of LIGHT into a mouse
fibrosarcoma led to strong anti-tumor responses and regression of
established tumors [23]. In this study we investigated the effects
of LIGHT in hematopoietic malignancies and we chose to focus
on B-CLL, given the strong expression of HVEM in these cells and
our previous results on the importance of LIGHT in the
physiology of B lymphocytes [17]. Thus, to explore the impact
of LIGHT in this lymphoid malignancy, we stimulated B-CLL cells
with an anti-HVEM mAb and performed a QRT-PCR global
analysis of the expression of a large panel of genes. Among the
322 genes tested, only 12 were significantly over-expressed
following HVEM stimulation, suggesting a restricted transcrip-
tional profile for HVEM triggering in B-CLL cells. HVEM ligation
upregulated the expression of two genes coding for chemokines:
IL-8 and IP10, and one chemokine receptor, Chemokine (C-C
motif) receptor 4, each of them critical for the recruitment of
immune effectors. These observations were not surprising since
HVEM–LIGHT signaling was already associated with IL-8 and
TNF-a production in monocytic cell lines [33–35]. At the protein
level, HVEM induced a significant chemokine release of not only
IL-8 but also Rantes. Although Rantes was secreted by tumor
cells, upregulation of its transcript was not detectably high to
reach the level of significance. The production of IL-8 seems to be
a key element in HVEM response, since it was described for
several cell types, including monocytes, neutrophils, and in this
study, in leukemic B-CLL cells. Before analyzing this large
transcriptional response to HVEM stimulation, we expected
upregulation of genes associated with immune activation, given
the established functions of LIGHT and its receptor HVEM in
costimulatory responses. But surprisingly, the nine other genes
significantly induced after HVEM stimulation, Bcl-XS, Bid, BNIP3,
all members of the Bcl-2 family, and FasL, CARD11, and
Cytochrome c, p53, all correspond to genes coding for apoptotic
proteins, most of them pro-apoptotic; and the two others, BIRC4
and IEX-1, belong to the anti-apoptotic subgroup.
We observed that this apoptotic transcriptional profile was
correlated with a functional effect in vitro, as LIGHT-transfected
cells or HVEM mAb both effectively increased the killing of B-CLL
cells. Furthermore, we confirmed that the killing effect of LIGHT
on B-CLL cells was mediated through its interaction with HVEM,
as demonstrated by: (i) the inability of rhLTa1b2, the other LT-bR
ligand, to induce CLL cell death and (ii) B-CLL cells, which did
not express LT-bR, were nevertheless killed by anti-HVEM mAb.
This direct killing effect of HVEM on a lymphoid malignancy was
unexpected because HVEM has been considered until now as a
costimulatory receptor and the pro-apoptotic effect of LIGHT was
proposed to be mediated only via LT-bR [18]. We had previously
shown that LIGHT triggering via HVEM increased the sensitivity
of mantle cell lymphoma to Fas-induced apoptosis, but LIGHT
alone failed to induce apoptosis of these cells [29].
HVEM belongs to the subgroup of TNFR, which do not contain
a DD, and are primarily involved in cell survival and costimula-
tory responses. However, it has been reported that some TNFR
members that do not contain a DD can also induce cell death. For
example, CD27-induced apoptosis of a Burkitt’s lymphoma cell
line [36], CD30, was involved in cell death signaling for thymic
negative selection [37], CD40 ligation caused apoptotic cell death
in transformed cells of mesenchymal and epithelial origin [38],
and LIGHT-induced cell death in some adenocarcinoma through
interaction with LT-bR . Our results add HVEM to this subgroup.
Nevertheless, the absence of DD complicated the understanding
of the pathways involved in the HVEM-mediated tumor cell
death.
Figure 7. LIGHT stimulation of primary B-CLL induces both TNFproduction and increased cell death via TNF-a. (A) B-CLL cells werecocultured with LIGHT or CD32 (negative control) transfected L-cells.Culture supernatants were collected at 24 h and assayed for TNF-asecretion by ELISA. TNF production by the transfected cells alone wasverified (data not shown). The bars depict the mean TNF secretionobserved for cells from seven different patients7SEM. �po0.05compared with control condition with a Wilcoxon test. (B) B-CLL cellswere cocultured with LIGHT-transfected L-cells, in the presence orabsence of 100 ng/mL rh-TNF-a for 24 h. Cells were then analyzed byflow cytometry for Annexin V/PI double staining. The hatched barrepresents the mean increase in cell death in the LIGHT1TNF-acondition, compared with the LIGHT condition (open bar, set at100%)7SEM for cells from seven different patients.
Figure 8. LIGHT induces cell death of CD40L-activated B-CLL cellsB-CLL cells were pre-activated with CD40L or CD32 (negative control)transfected L-cells for 24 h. Then, cells were either treated with anti-Fas mAb (CH11 clone), anti-HVEM mAb, or left untreated for 24 h.Percent of cell death was evaluated by flow cytometry for Annexin V/PIstaining. The bars depict the mean percentage of dead cells aftersubtracting the value of control condition, observed for two indepen-dent experiments7SEM, with cell death obtained for HVEM mAbcondition set to 100%.
Eur. J. Immunol. 2009. 39: 2502–2514 Immunomodulation 2509
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We found that HVEM stimulation induced activation of
caspases 3, 8, and 9. Pre-incubation of B-CLL cells with the pan-
caspase inhibitor Z-VAD-FMK totally abrogated caspase 3 acti-
vation, but not cell death, suggesting the involvement of a
mechanism independent of caspases. Moreover, HVEM stimula-
tion decreased mitochondrial membrane potential and induced
an upregulation of the pro-apoptotic Bax protein expression. In
contrast, the anti-apoptotic Bcl-2 protein was unaltered. The
mitochondrial activity has been shown to be strictly controlled by
the balance between these Bcl-2 family members [39, 40]. In
addition, the loss of membrane potential was not blocked by the
pan-caspase inhibitor Z-VAD-FMK, suggesting that this step was
also independent of caspase activation. Altogether, these data
suggest that HVEM-induced cell death involves on one hand the
death receptor pathway (activation of caspase 8) and the mito-
chondria pathway (decreased mitochondrial membrane potential
and upregulation of Bax) in an apoptotic mechanism and on the
other hand there is also an unidentified pathway independent of
caspases also leading to cell death.
Previous reports have shown that the TNFR family members
devoid of DD sometimes involved indirect apoptotic mechanisms
through activation of other TNFR members. For example, a report of
Grell et al. showed that the cytotoxic effects induced by TNFRII,
CD40, and CD30 are mediated by endogenous production of TNF-
aand autotropic or paratropic activation of TNFRI [41]. Recent
developments have shed light on the mechanism whereby endogen-
ous TNF production can be switched on, following rapid degradation
of IAP1 and IAP2 proteins triggered by pharmacological compounds
[42, 43]. It remains to be determined whether this pathway can also
be triggered by non-DD TNFR members such as HVEM. In our results,
HVEM stimulation resulted in a major increase in FADD expression.
Since FADD is the major adaptor molecule involved in Fas and
TNFR-mediated cell death, this may suggest an involvement of other
TNFR in HVEM-mediated cell death. We confirmed that HVEM
induced an increase in the surface expression of Fas in CLL, as we
previously described in MCL [29], and interestingly, as we report in
this study, HVEM also upregulated the surface expression of TRAIL.
The combination of HVEM mAb with FasL was more effective than
HVEM mAb alone to induce killing of B-CLL cells. In contrast,
combination with TRAIL did not increase apoptosis above that of
HVEM mAb alone. These data suggest that HVEM TRAIL pathways
may involve shared mediators to induce killing of B-CLL cells. In
addition, pre-treatment of B-CLL cells with anti-TRAIL mAb signifi-
cantly inhibited HVEM-induced cell death. Altogether, these results
strongly suggest that the TRAIL pathway was one of the components
of HVEM-induced killing. In addition, HVEM triggering led to endo-
genous TNF-a production, which was inhibited by anti-TNF-a mAb
and more importantly HVEM stimulation led to increased cell death
following addition of TNF-a. Our results correlate with the TNF
production that was previously observed following CD40, CD30, or
TNFRII triggering. Notably, here blocking anti-TNF antibodies did not
abrogate HVEM-mediated cell death. Therefore, HVEM induces cell
death by (i) upregulation of cytotoxic ligands, such as TNF, and
(ii) sensitization of cells to apoptosis by a signaling pathway, which is
independent of endogenous TNF.
Finally, most of the studies in CLL are performed in cells from the
periphery that are resting, whereas the CLL cells are activated and
proliferate in the lymph nodes and bone marrow. In order to mimic
these conditions we used in vitro stimulation by CD40L, which
provides a robust activation of CLL cells. Other studies previously
found that this pre-activation with CD40L could sensitize B-CLL cells
to the FasL and TRAIL-mediated apoptosis [44]. Under these condi-
tions, HVEM triggering is still able to induce CLL cell death.
Considering a therapeutic administration of HVEM, we
observed that the HVEM-induced killing compared favorably with
that of the pan-B cell therapeutic mAb rituximab. Further
experiments are required to determine if HVEM could enhance
efficacy of therapeutic agents, such as fludarabine, as it was
demonstrated for rituximab and alemtuzumab mAb [45].
In conclusion, the present data describe the novel ability for
the LIGHT–HVEM couple to induce death of fresh B-CLL tumor
cells together with the production of selected chemokines. Here,
LIGHT induced the cell death of a B lymphoid malignancy
directly through HVEM without the undesirable effect of inducing
proliferation [17]. LIGHT–HVEM signaling may represent a novel
therapeutic target for cancer therapy, as soluble LIGHT or anti-
HVEM mAb could present the attractive advantage of combining
direct killing of malignant lymphocytes expressing HVEM, with
the activation of immune effector cells and their recruitment
through the production of chemokines.
Materials and methods
Cells and reagents
This study was approved by the review board of the Institut Paoli-
Calmettes, Marseille, France. After informed consent in accordance
with the Declaration of Helsinki, peripheral-blood samples were
obtained from untreated patients diagnosed with CLL on the basis of
clinical and immunophenotypic criteria. The mononuclear cells were
isolated by density gradient centrifugation (Lymphoprep, AbCys) and
viably frozen in fetal bovine serum (PAN Biotech) containing 10%
dimethyl sulfoxide (Sigma). Clinical characteristics for 25 CLL
patients are summarized in Table 1, and include Binet stage,
percentage.
Stably transfected cells CD40L, LIGHT, CD40L/LIGHT, GpD,
and BTLA were obtained either by electroporation (960mF, 220 V,
BIO RAD Gene Pulser and Capacitance extender) or transfection
using Fugene-6 (Roche Diagnostics) of LTK murine fibroblasts
(L cells) with pcDNA3.1 vector (Invitrogen, Groningen, The
Netherlands) encoding human CD40L, LIGHT, GpD, or BTLA,
respectively. Expression of the molecule of interest was verified
by flow cytometry using PE-conjugated mAb anti-CD40L (BD
Biosciences), PE-conjugated anti-LIGHT (BD Biosciences), FITC-
conjugated anti-gD, and FITC-conjugated anti-BTLA (both
produced in our laboratory). CD32-transfected fibroblasts were a
kind gift from Schering-Plough (Dardilly, France). Transient
transfected Cos-FasL cells and Cos-TRAIL cells were generated by
Eur. J. Immunol. 2009. 39: 2502–2514Christine Pasero et al.2510
& 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.eji-journal.eu
respectively transfecting monkey kidney Cos cell line with
pcDNA3-h-FasL vector (Cayla) or pORF h-TRAIL vector (Cayla).
The pan-caspase inhibitor N-carbobenzoxy-Val-Ala-Asp fluor-
omethyl ketone (Z-VAD-FMK) was obtained from BD Biosciences.
The anti-HVEM mAb was kindly provided by Synaptex. The anti-
TNF mAb were generated in our laboratory by standard hybri-
doma techniques. Blocking anti-TRAIL mAb RIK-2 was obtained
from BD Biosciences. The recombinant human TNF-a was
purchased from R&D Systems.
Immunofluorescence analysis of cell-surface antigens
Cell surface analysis of B-CLL cells was performed through flow
cytometry with the use of a FACSCanto cytometer and FACSDiva
software (Becton Dickinson, Mountain View, CA). B-CLL cells
were immunostained at 41C for 30 min, with the following mAb:
FITC-conjugated anti-CD19 (Beckman Coulter), FITC-conjugated
anti-HVEM (Medical & Biological Laboratories), PE-conjugated
anti-LT-bR (BD Biosciences), FITC-conjugated anti-Fas (CD95)
(Beckman Coulter), PE-conjugated anti-TRAIL (BD Biosciences),
PE-conjugated anti-FasL (Biolegend), PE-conjugated anti-DR4 or
PE-conjugated anti-DR5 (R&D Systems). FITC- or PE-labeled
isotype-matched Ig were used as negative controls. After
two washings in PBS 2% FBS, cells were analyzed by flow
cytometry.
Annexin V/PI staining
Following stimulation, cell death was analyzed by Annexin
V-Cy5 (BD Biosciences) and PI double staining (BD Biosciences).
Briefly, 5� 105 cells were washed once with PBS 1% FBS and
resuspended in 100mL 1� binding buffer (BD Biosciences) with
5mL Annexin V-Cy5 for 10 min at room temperature in the dark.
Then 200mL of 1� binding buffer and 3.5 mL of PI were added,
and cells were incubated for 5 min at room temperature in the
dark. Cells were then analyzed on a FACSCanto cytometer
(Becton Dickinson). Data analysis was performed with FACSDiva
software (Becton Dickinson). Dead cells were measured as the
percentage of Annexin V and PI double positive cells.
Determination of caspase, Bax, and Bcl-2 activation byflow cytometry
Caspases 8 and 9 activities were measured at different times
using the following cell-permeable fluoresceinated caspase
inhibitors: FAM-LETD-FMK (caspase 8 inhibitor) and FAM-
LEHD-FMK (caspase 9 inhibitor) (CaspGLOW kits, Biovision).
Cells were stained for 1 h at 371C in the dark, then washed,
resuspended in 500mL buffer, and immediately analyzed by flow
cytometry on a FACSCanto cytometer (Becton Dickinson).
To detect activated intracellular caspase 3, Bcl-2, and Bax,
cells were permeabilized using cytofix/cytoperm buffer (BD
Biosciences) and stained with PE-conjugated anti-active caspase 3
mAb (BD Biosciences), PE-anti-Bax mAb (Santa Cruz Biotech-
nology), or FITC-anti-Bcl-2 mAb (BD Biosciences), and then
analyzed by flow cytometry.
Determination of mitochondrial membrane potential(Dwm)
After stimulation, B-CLL cells were washed, resuspended with
1 mL PBS, and supplemented with 5 mL of 10 mM of the cationic
cyanine dye DiOC2(3) solution, incubated for 30 min (Molecular
Probes) at 371C in the dark. Cells were washed again,
resuspended with 500mL PBS, and analyzed by flow cytometry
to detect DiOC2(3) fluorescence. Such fluorescence decreases
when cells undergo apoptosis. The positive control was the
carbonyl cyanide m-chloro phenyl hydrazone, a mitochondrial
uncoupling agent.
Cytokine and chemokine production
Supernatants were harvested after a 24 h stimulation. The
cytokines were measured using the Cytometric Bead Array
Human Chemokine kit (BD Biosciences), a multiplexed based
immunoassay which allows quantitative detection of several
cytokines (IL-8/CXCL8, Rantes/CCL5, MIG/CXCL9, MCP-1/
CCL2, IP-10/CXCL10) in the same supernatant, as described by
Table 1. Clinical characteristics of the B-CLL patientsa)
Characteristics N
No. of patients 25
Median age (y) (range) 58 (36–72)
Male/female 16/9
Binet stage
A 19
B 5
C 1
Median %CD5/CD19 (y) (range) 84 (47–98)
b2microglobulin
43 mg/L 4
o3 mg/L 15
Not evaluated 6
LDH
4 Normal values 2
Normal values 16
Not evaluated 7
Lymphocyte doubling time
4 12 months 14
o12 months 6
Not evaluated 5
Treatment
Yes 2
No 23
a) IgVH status was available only for three patients (mutated forms).
Eur. J. Immunol. 2009. 39: 2502–2514 Immunomodulation 2511
& 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.eji-journal.eu
the manufacturer. IL-8 and TNF- production were also quantified
using ELISA assays (R&D Systems and BD Biosciences, respec-
tively) according to the manufacturer’s protocols.
Western blotting analysis
Proteins from NP40 lysates were separated by SDS-PAGE in
denaturing conditions, transferred onto membranes, and then
probed with anti-cleaved caspase 3 (Cell Signaling) or anti-FADD
(produced in our laboratory) antibody. The bands were then
visualized with HRP-conjugated anti-mouse or anti-rabbit IgG,
and Western blot chemiluminescence reagent (West Pico, Pierce).
b-actin expression was determined on the same blots after
stripping. Each band was scanned (Powerlook 1000, Umax) and
quantified using the Phoretix 1D Advanced software (Nonlinear
Dynamics). Data were then converted to a fold change ratio
obtained by dividing the stimulated condition values normalized
with b-actin by values determined for unstimulated cells normal-
ized with b-actin. Ratios are indicated in the legends of the
figures.
QRT-PCR
Before RNA isolation, all cells were selected for their ability to be
killed by HVEM mAb. QRT-PCR analysis was performed with the
Applied Biosystems 7900HT Fast RT PCR system. Briefly, total
RNA was isolated from B-CLL 1 day post stimulation and reverse
transcribed. Then, the two following supports were used. In the
first method, 87 gene targets each coding for adhesion,
migration, cytokines, or apoptosis molecules were spotted on
optical plates (Microamp Fast Optical 96 well, Applied Biosys-
tems). QRT-PCR was performed in a 20 mL reaction containing
2� SYBR Green reagent, 200 nM primers, and 0.5 mL cDNA
(equivalent to 10 ng of total RNA) in each well. In the second
method, 96 gene targets each coding for immune molecules were
spotted into TaqMan low-density array (Immune Panel Micro-
fluidic Card, Applied Biosystems). Briefly, 5mL cDNA (equivalent
to 100 ng of total RNA) was mixed with 100 mL of 2� TaqMan
universal Mix (Applied Biosystems) and loaded into one sample
port. Capture of fluorescence was recorded on the ABI Prism
7900HT scanner, and the Ct (threshold cycle) was calculated for
each assay using Sequence Detection System Software 2.1
(Applied Biosystems). Normalization of quantitative-PCR assays
was conducted using b-actin as endogenous control. Data were
then converted to a fold change ratio described with the formula:
2�DDCt, where DCt 5 Ct target–Ct endogenous control, and
DDCt 5DCt stimulated condition –Ct unstimulated condition.
Statistical analysis
Results were compared by the non-parametric Wilcoxon signed-
rank test, to evaluate any statistically significant difference
between HVEM mAb treated condition and untreated condition.
Differences were considered significant when po0.05. p Values
are indicated in the legends of figures.
Acknowledgements: This work was supported by institutional
resources provided by INSERM. C.P. was in part supported for a
PhD fellowship by the Association pour la Recherche sur le
Cancer. We thank Eloıse Perrot for excellent technical assistance,
Dr. Sophie Gomez for providing anti-FADD antibody, Dr. Marc
Lopez for providing anti-HSV gD antibody, Dr. Claude Mawas for
helpful comments. We also thank Michele Batoz of the
‘‘Laboratoire Central d’Immunologie des Hopitaux de Nice
(Inserm UMR576)’’ for supplying optical plate primers.
Conflict of interest: The authors declare no financial or
commercial conflict of interest.
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Abbreviations: AML: acute myeloid leukemias � BTLA: B and T
lymphocyte attenuator � CLL: chronic lymphocytic leukemia � Ct:
threshold cycle � DD: death domain � FADD: Fas-associated death
domain � gD: Glycoprotein D � HVEM: herpes virus entry mediator �IP10: IFN-inducible protein 10 � LT-bR: lymphotoxin-b receptor � MCL:
mantle cell lymphomas � Rantes: regulated on activation, normal T-
cell expressed and secreted � TNFR: TNF receptor � QRT-PCR: real-time
quantitative PCR
Full correspondence: Professor Daniel Olive, INSERM UMR891, Centre de
Recherche en Cancerologie de Marseille, Universite de la Mediterranee,
IFR 137 Institut de Cancerologie et d’Immunologie de Marseille, 27 Bd.
Leı Roure, 13009 Marseille, France
Fax: 133-491-26-03-64
e-mail: [email protected]
Supporting Information for this article is available at
www.wiley-vch.de/contents/jc_2009/39069_s.pdf
Received: 8/11/2008
Revised: 16/6/2009
Accepted: 16/6/2009
Eur. J. Immunol. 2009. 39: 2502–2514Christine Pasero et al.2514
& 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.eji-journal.eu