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: Daniel.Olive@inserm.fr

& 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.

Eur. J. Immunol. 2009. 39: 2502–2514 Immunomodulation 2503

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).

Eur. J. Immunol. 2009. 39: 2502–2514Christine Pasero et al.2504

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).

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.

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.

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.

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%.

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

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

Median %CD5/CD19 (y) (range) 84 (47–98)

b2microglobulin

43 mg/L 4

o3 mg/L 15

Not evaluated 6

4 Normal values 2

Normal values 16

Not evaluated 7

Lymphocyte doubling time

4 12 months 14

o12 months 6

Not evaluated 5

Treatment

a) IgVH status was available only for three patients (mutated forms).

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: Daniel.Olive@inserm.fr

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

A role for HVEM, but not lymphotoxin-beta receptor, in LIGHT-induced tumor cell death and chemokine...

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