interactions via chemokine metabolism Anergic T

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2003 102: 2173-2179 Prepublished online May 29, 2003; doi:10.1182/blood-2003-02-0637

Anergic T cells exert antigen-independent inhibition of cell-cell interactions via chemokine metabolism Martha J. James, Lavina Belaramani, Kanella Prodromidou, Arpita Datta, Sussan Nourshargh, Giovanna Lombardi, Julian Dyson, Diane Scott, Elizabeth Simpson, Lorraine Cardozo, Anthony Warrens, Richard M. Szydlo, Robert I. Lechler and Federica M. Marelli-Berg

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From bloodjournal.hematologylibrary.org by guest on July 13, 2011. For personal use only. IMMUNOBIOLOGY

Anergic T cells exert antigen-independent inhibition of cell-cell interactions via chemokine metabolism Martha J. James, Lavina Belaramani, Kanella Prodromidou, Arpita Datta, Sussan Nourshargh, Giovanna Lombardi, Julian Dyson, Diane Scott, Elizabeth Simpson, Lorraine Cardozo, Anthony Warrens, Richard M. Szydlo, Robert I. Lechler, and Federica M. Marelli-Berg

Due to their ability to inhibit antigeninduced T-cell activation in vitro and in vivo, anergic T cells can be considered part of the spectrum of immunoregulatory T lymphocytes. Here we report that both murine and human anergic T cells can impair the ability of parenchymal cells (including endothelial and epithelial cells) to establish cell-cell interactions necessary to sustain leukocyte migration

in vitro and tissue infiltration in vivo. The inhibition is reversible and cell-contact dependent but does not require cognate recognition of the parenchymal cells to occur. Instrumental to this effect is the increased cell surface expression and enzymatic activity of molecules such as CD26 (dipeptidyl-peptidase IV), which may act by metabolizing chemoattractants bound to the endothelial/epithelial cell

surface. These results describe a previously unknown antigen-independent antiinflammatory activity by locally generated anergic T cells and define a novel mechanism for the long-known immunoregulatory properties of these cells. (Blood. 2003;102:2173-2179)

© 2003 by The American Society of Hematology

Introduction T-cell anergy has been defined as a “cellular state in which a lymphocyte is alive but fails to display certain functional responses (including cell division and interleukin 2 [IL-2] production) when optimally stimulated through both its antigen-specific receptor and any other receptor that is normally required for full activation.”1 Anergic T cells can be generated in vitro and in vivo by various mechanisms,1,2 all involving partial or inappropriate stimulation. While losing their proliferative and effector potential, anergic T cells have long been known to be able to exert immunoregulation. The potential for anergic T cells to act as suppressor cells came first from a superantigen in vivo model in which anergic T cells acted as efficient suppressor cells in an antigen-nonspecific manner.3 More recently, murine anergic T cells either generated in vivo or rendered anergic in vitro with immobilized anti-CD3 and adoptively transferred have been shown to prolong skin allograft survival in an antigen-specific manner.4,5 In the human system, anergic CD4⫹ T cells were shown to exert contact-dependent and antigen-specific suppression in vitro.6,7 The mechanism by which anergic T cells exert their immunoregulatory properties appears to be indirect by altering the antigen presenting cells (APCs) immunogenicity8,9 in a cell-cell contactdependent manner. The molecular basis for this effect is still unknown and it has been hypothesized to involve the induction of “regulatory” molecules on the anergic T cells, capable of delivering negative signals to the APC.2 It has recently become clear that the initial stages of T-cell activation are mediated by antigen-independent interactions, which establish areas of focal contact between T cells and APCs.10,11 Such interactions are initiated by chemoattractant-induced cell polarization and subsequent redistribution of adhesion molecules on the

T-cell surface. These, in turn, allow T-cell receptor (TCR) interactions with the major histocompatibility complex (MHC)–peptide complexes displayed on the APC and lead to the formation of highly organized structures named immunologic synapses, which efficiently transduce antigen-initiated signals. Thus, one way of diminishing the efficiency of T-cell activation is the disruption of antigen-independent contact with the APC. In this study, we have explored the possibility that anergic T cells may act by altering the adhesive interactions between T cells and other cell types using in vitro and in vivo models of tissue infiltration by T cells. We observed that the ability of endothelial and epithelial cells to mediate migration of lymphocytes and other leukocytes, an adhesion-dependent function, is altered following noncognate interactions with anergic T cells. Similarly, in vivo contact with anergic T lymphocytes prevents infiltration of the peritoneal membrane by antigen-specific T cells in a murine peritonitis model. This effect appears to be mediated by increased surface expression by anergic T cells of the dipeptidyl-peptidase IV, CD26, which might act by metabolizing chemotactic cytokines on the endothelial/epithelial cell surface, thus altering adhesion-dependent migration, and ultimately, tissue infiltration.

Materials and methods Mice Male and female 4- to 8-week-old C57BL/6 mice and CBA/Ca mice were purchased from Olac Harlan (Bicester, United Kingdom) and used at 6 to 8 weeks of age.

From the Department of Immunology, Faculty of Medicine, Cardiovascular Medicine Unit, Transplantation Biology, Medical Research Council Clinical Research Centre; and Department of Hematology, Imperial College London, Hammersmith Hospital Campus, London, United Kingdom.

Reprints: Federica Marelli-Berg, Department of Immunology, Division of Medicine, Faculty of Medicine, Imperial College London, Hammersmith Campus, Du Cane Road, London W12 0NN, United Kingdom; e-mail: [email protected].

Submitted February 27, 2003; accepted May 16, 2003. Prepublished online as Blood First Edition Paper, May 29, 2003; DOI 10.1182/blood-2003-02-0637.

The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.

Supported in part by the British Heart Foundation (Grant PG/2001014).


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Peptides, antibodies, and reagents inhibitor12

The CD26 ectopeptidase human peptide YY (3-36) was purchased from Bachem UK (Meyerside, United Kingdom). The following monoclonal antibodies (mAbs) were used: antihuman CD3 (OKT3; American Type Culture Collection [ATCC], Rockville, MD), antihuman CD28 (CD28.2; BD Biosciences, Oxford, United Kingdom), antihuman CD26 (M-A261; Serotec, Kidlington, United Kingdom), antimouse CD4 (YTS 191,13), antimouse CD26 (H194-112; BD Biosciences), antimouse CD3 (2C11; BD Biosciences), and antimouse CD28 (MCA1363; Serotec). The human chemokine CXCL-12 (stromal-derived factor-1␣ [SDF-1␣]) was purchased from Pepro Tech (Peterborough, United Kingdom). Endothelial cells and epithelial cells Endothelial cells (ECs) and renal tubular epithelial cells (RTECs) were isolated from human umbilical cord veins and tissue, respectively, and cultured as previously described.14,15 Prior to use in functional assays, human umbilical vein endothelial cells (HUVECs) and RTECs were treated with either 300 U/mL interferon-␥ (IFN-␥) for 72 hours or with 10 ng/mL tumor necrosis factor ␣ (TNF-␣; Pepro Tech) to optimize migration. T-cell clones and lines Human CD4⫹ T-cell clones, HC3 and 7P.61, specific for the influenza hemagglutinin peptide HA100-115 and HA307-319 and restricted by DRB1*0101 and DRB1*0701, respectively, were generated and cultured as described.16,17 Alloreactive CD4⫹ T-cell lines were generated by stimulation of purified peripheral blood CD4⫹ T cells with allogeneic irradiated (60 Gy) peripheral blood mononuclear cells (PBMCs) and maintained in culture by weekly restimulation in the presence of recombinant IL-2 (rIL-2; 10 U/mL; Roche, Mannheim, Germany) in RPMI 1640 medium supplemented with 10% human serum (HS). The murine CD8⫹ C6 T-cell clone, specific for HY peptide epitope TENSGKDI presented by H2-Kk,18 and the murine CD4⫹ T-cell clone B9, specific for HY peptide epitope NAGFNSNRANSSRSS and restricted by H2-Ab,19 were maintained in culture by forthnightly stimulation with peptide-pulsed splenocytes and rIL-2 (20 U/mL; Roche) in RPMI 1640 medium supplemented with 10% fetal calf serum (FCS; Globepharm, Guildford, United Kingdom), and 50 mM 2-mercaptoethanol (2-ME) (Gibco, Paisley, United Kingdom). Isolation of human peripheral blood granulocytes Heparinized blood from healthy adult volunteers was layered on a Ficoll-Paque gradient (Pharmacia, Uppsala, Sweden) and centrifuged for 20 minutes at 800g. The plasma and the interphase consisting mainly of lymphocytes and monocytes were discarded. Erythrocytes were eliminated from the pellet by rapid osmotic lysis. The remaining cell suspension contained more than 99% granulocytes. Cell viability was more than 98%, as determined by trypan blue exclusion (data not shown).

vivo experiments described in this paper. As both resting and activated T cells displayed the same behavior in all the experiments performed, they will be referred as to “responsive” T cells throughout the text. Lymphocyte transmigration assays and chemotaxis assays Transmigration assays were carried out using HUVEC or RTEC monolayers (5 ⫻ 104 cells/12-mm–diameter well) grown on a 3-␮m–pore polycarbonate transwell (Costar, High Wycombe, United Kingdom), as previously described,22 and incubated overnight with anergic or responsive T cells (5 ⫻ 104/well). Following removal of the T cells, fresh T cells (5-7 ⫻ 105/ well, with different antigen specificity) or granulocytes (106/well) were added into each insert and left to migrate through the monolayers. The number of migrated T cells or granulocytes was determined by counting the cells present in the well media over the next 24 hours. Results are expressed as percentage of transmigrated cells in 3 independent counts. For chemotaxis assays with CXCL-12, naive CD4⫹ CD45RA⫹ T isolated by negative immunomagnetic selection were seeded in the top chamber of a 3-␮m–pore polycarbonate transwell (Costar).23 Chemotaxis medium (RPMI 2% FCS) containing CXCL-12 (50 ng/mL), or chemotaxis medium alone, was added to the bottom chamber of the transwell, while the cell suspension was added to the top chamber. The number of migrated cells was evaluated as described in “Lymphocyte transmigration assays and chemotaxis assays.” Peritoneal membrane infiltration by antigen-specific T cells Male CBA/Ca mice were given an intraperitoneal injection of IFN-␥ (600 U; Pepro Tech), which induces a localized inflammatory response (F.M.M.B., unpublished observations, January 2001). As a control, an untreated female mouse was used. HY-specific C6 CD8⫹ T cells (5-7 ⫻ 106) previously labeled with the PKH26 Cell Linker (Sigma, Poole, United Kingdom) were injected intraperitoneally 48 hours later. After a further 24 hours, mice were killed and infiltration of the peritoneal membrane by labeled T cells was visualized by wide-field fluorescence microscopy (see “Wide-field fluorescence microscopy and flow cytometry”). The presence of fluorescently labeled cells in the peritoneal lavage, draining lymph nodes, and the spleen was assessed by cytofluorymetric analysis. In some experiments, responsive or previously anergized unlabeled CD8⫹ C6 (antigen-specific) or CD4⫹ B9 (irrelevant) T cells (4-5 106) were injected intraperitoneally 24 hours prior to the injection of C6 T cells. To assess the functional role of T-cell surface CD26, samples of anergic T cells were pretreated with anti-CD26 (5 ␮g/mL), anti-CD4 (5 ␮g/mL), or an isotypematched antibody (5 ␮g/mL) for 30 minutes at room temperature (RT) and washed prior to injection. Alternatively, anergic T cells were incubated with human peptide YY (3-36, 30 nM) for 30 minutes at RT and washed before injection. Wide-field fluorescence microscopy and flow cytometry

Anergy induction in human and murine T cells For anergy induction by costimulation-deficient TCR triggering, T cells (alloreactive human CD4⫹ T-cell lines and the B9 murine CD4⫹ T-cell clone) were plated in 24-well plates (5 ⫻ 105/well) in the presence of plastic-bound anti-CD3 (1 ␮g/mL OKT3 or 10 ␮g/mL 2C11) in a total volume of 500 ␮L.20 Anergy was also induced by T-T antigen presentation. Human CD4⫹ T-cell clones (HC3 and 7P.61) were incubated overnight (106 T cells) in the presence of 0.7 ␮g/mL cognate HA peptides17 while C6 T cells (5 ⫻ 105) were incubated with 100 nM HY peptide in 24-well plates.21 Following 24-hour incubation, T cells were harvested and viable cells recovered by density gradient prior to use in functional assays. Induction of anergy was assessed by measuring 3H thymidine incorporation following antigenic rechallenge. Resting lymphocytes and T cells activated with plastic-bound anti-CD3 (OKT3; 1 ␮g/mL for human T cells or 2C11 [10 ␮g/mL] for murine T cells) plus anti-CD28 (CD28.2; 5 ␮g/mL for human T cells and MCA1363 [5 ␮g/mL] for murine T cells) were used as a control in all the in vitro and in

Peritoneal membranes were laid onto polysine microscope slides (VWR International, Lutterworth, United Kingdom), left to dry overnight, and then mounted in Vectorshield mounting medium for fluorescence with DAPI (4,6 diamidino-2-phenylindole; Vector Laboratories, Peterborough, United Kingdom). Slides were visualized with a Coolview 12–cooled camera (Photonic Science, Newbury, United Kingdom) mounted over a Zeiss Axiovert S100 microscope equipped with Metamorph software (Zeiss, Welwyn Garden City, United Kingdom). A ⫻ 10 and a ⫻ 40 0.6 objectives and a standard epi-illuminating rhodamine fluorescence filter cube were used and 12-bit image data sets were generated. Tissue infiltration was quantified by randomly selecting ten ⫻ 40-magnified fields and assessing the number of fluorescent cells in each field. For flow cytometry, 105 cells were incubated with the indicated fluoresceinated mAb at 4°C for 30 minutes. An isotype-matched irrelevant antibody was used as a control. Stained cells were analyzed using a FACSCalibur (Becton Dickinson, Mountain View, CA).

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Statistical analysis In the in vitro experiments, comparisons between groups were made using the Student t test. To assess statistical significance in the in vivo experiments, the Mann-Whitney U test was used. All reported P values are 2-sided.

Results Parenchymal cells fail to establish effective cell-cell interactions following exposure to anergic T cells

The cellular ability to establish effective cell-cell interactions following exposure to anergic T cells was tested by an in vitro model of lymphocyte transmigration. Cytokine-treated EC/RTEC monolayers grown on transwells (5 ⫻ 104/well) were incubated overnight with either anergic (5 ⫻ 104/well) or responsive (105/ well) CD4⫹ T-cell lines or clones, or in medium alone. The number of responsive T cells used was doubled to compensate for the fraction of T cells that would have migrated through the monolayer overnight (about 50% of the initial input on average22), while the majority of anergic T cells remained in the top chamber, having lost their migratory ability.22 Following thorough removal (97% recovery on average) of the T cells, fresh T lymphocytes of similar or different specificity (7 ⫻ 105/well), or granulocytes (106/well) were seeded onto the EC monolayers. Cell migration was monitored over the next 6 to 26 hours. The following combinations were used: anergic 7P.61 CD4⫹ T-cell and responsive 7P.61, HC3, or alloreactive CD4⫹ T-cell lines, anergic allospecific CD4⫹ T-cell line and responsive 7P.61, or granulocytes. 7P.61 T cells were rendered anergic by T-T presentation, while the alloreactive T-cell lines were incubated on plastic-bound anti-CD3. As it is shown in Figure 1 both T cells (Figure 1A) and granulocytes (Figure 1B) migrated less efficiently through EC monolayers exposed to anergic T cells. Incubation with responsive T cells often led to increased migration, particularly at early time points. This effect was cell-contact dependent (Figure 1C). The EC monolayer was not disrupted by anergic T cells (data not shown). In addition, culture of human RTEC monolayers with anergic T cells diminished the ability of the epithelium to mediate T-cell migration (Figure 1D). The inhibition was equally exerted by T cells rendered anergic by T-T presentation and costimulation-deficient TCR triggering and did not require cognate recognition of ECs by the anergic T cells (data not shown). Consistently, similar results were obtained using TNF-␣–treated MHC class II–negative ECs (data not shown). To investigate whether the observed effect was reversible, ECs were activated via CD40 ligation24,25 during exposure to the anergic T cells. As shown in Figure 1E-G, CD40 ligation led to a partial recovery of EC function (Figure 1E). Ligation of CD40 on EC incubated with either responsive T cells or medium did not modify their function (Figure 1F and 1G, respectively). Anergic T cells prevent tissue infiltration in vivo

Preliminary to these experiments we observed that the anergic murine C6 or B9 T cells, similarly to their human counterpart, were capable to inhibit EC function in vitro (data not shown). As our working hypothesis is that during inflammation locally generated anergic T cells modulate the ability of surrounding parenchymal cells to establish functional cell-cell interactions that support leukocyte infiltration, a model of peritoneal membrane infiltration in which anergic T cells are administered locally by intraperitoneal

Figure 1. Anergic T cells inhibit parenchymal cell–mediated migration. (Panels A-B) EC monolayers (5 ⫻ 104/transwell) were incubated for 16 hours with either anergic (F; 5 ⫻ 104/transwell) or responsive (E; 105/well) alloreactive CD4⫹ T-cell lines or in medium alone (‚). T cells were subsequently removed and fresh T cells (7 ⫻ 105/well; panel A) or granulocytes (106/well; panel B) were seeded onto the EC monolayers. The results are expressed as percentage of migrated T cells at the given time points and reported as the average of 3 experiments of identical design. The bars show the standard deviations (SD). * indicates statistically significant (at least P ⬍ .01, panel A; at least P ⬍ .02, panel B) versus control cultures (EC monolayers cultured in medium). (Panel C) EC monolayers were incubated overnight with supernatants obtained from cultures of T cells in anergizing (F) or resting (E) conditions or in medium alone (‚). Following 2 washes with warm culture medium, fresh T cells (7 ⫻ 105/well) were seeded onto the EC monolayers. The results are expressed and reported as specified for panels A-B. (Panel D) TNF-␣–treated RTEC monolayers (5 ⫻ 104/transwell) were incubated for 16 hours with either anergic (5 ⫻ 104/well), or responsive (105/well) CD4⫹ T-cell lines or clones, or in medium alone. T cells were subsequently removed and fresh T cells (7 ⫻ 105/well) were seeded onto the RTEC layers. The results are expressed and reported as specified for panels A-B. * indicates statistically significant (at least P ⬍ .05) versus control cultures (RTEC monolayers cultured in medium). (Panels E-G) EC monolayers (5 ⫻ 104/transwell) were incubated for 30 minutes at RT with an antihuman CD40 mAb (5 ␮g/mL, filled symbols) or with an isotype-matched mAb (open symbols). Excess antibody was then removed by gentle washing. Bound antibody was then cross-linked by addition of goat antimouse Miltenyi microbeads (30 ␮L/␮g mAbs) during incubation with anergic (panel E) or responsive (panel F) 7P.61 T cells or in medium alone (panel G) for a further 16 hours. T cells were then removed and migration of freshly added alloreactive T cells (7 ⫻ 105/well) was monitored. In all these experiments, cell migration was monitored over the next 6 to 26 hours. The results are expressed and reported as specified for panels A-B. * indicates statistically significant (at least P ⬍ .01) versus control cultures (EC monolayers cultured in the presence of anergic T cells).

injection was developed. In this model, HY-specific C6 T cells (5-7 ⫻ 106) that are injected intraperitoneally in CBA male mice previously (48 hours) treated intraperitoneally with IFN-␥ promptly infiltrate the peritoneal membrane (Figure 2B,F). Untreated female


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anti-IL10 mAbs were ineffective (data not shown), antibody blockade of CD26 on anergic T cells prior to incubation with cytokine-treated EC monolayers completely prevented the inhibition of EC-mediated T-cell migration (Figure 3F) and partially that of EC-mediated granulocyte migration (Figure 3H). Antibody treatment of responsive T cells (which express intermediate levels of CD26) prior to incubation with the EC did not elicit any effect (Figure 3E,G). Treatment of anergic T cells with the anti-CD26 mAb did not restore responsiveness or motility23 in these cells (data not shown). Similar results were obtained when murine T cells were analyzed (data not shown). Anergic T cells metabolize chemokines via CD26 Figure 2. Anergic T cells inhibit tissue infiltration in vivo. Male CBA mice were injected intraperitoneally with 600 U IFN-␥. As a negative control, untreated female mice were used (panels A,E). Two days later, mice received an intraperitoneal injection of either PBS (panels A-B,E-F), responsive (3-5 ⫻ 107; panels C,G), or anergic (3-5 ⫻ 107; panels D,H) B9 CD4⫹ T cells (4 ⫻ 106/mouse). After 24 hours, red fluorescence-labeled (PKH26) HY-specific C6 T cells (5 ⫻ 107) were also injected intraperitoneally in all the mice. The following day, the presence of fluorescently labeled cells in the peritoneal membrane (⫻ 10 original magnification; panels A-D) and in the peritoneal lavage (panels E-H) was assessed by wide-field fluorescence microscopy and flow cytometry, respectively. The peritoneal membrane wide-field images and histograms (in which the percentage of positive cells is specified) are taken from a representative experiment; however, the median number and range of labeled cells quantified in 10 randomly selected ⫻ 40-magnified fields of the peritoneal membrane of at least 3 different animals are shown below each image (panels A-D). Similarly, the median percentage of labeled cells recovered in the peritoneal lavage of 3 different samples is specified below each representative histogram. * indicates statistically significant (P ⬍ .0001, panel D; P ⫽ .05, panel H) compared with male CBA mice (positive control).

mice, which displayed minimal C6 infiltration (the T cells being recoverable from the peritoneal lavage), were used as a control (Figure 2A,E). Anergic murine T cells were generated by T-T peptide presentation (clone C6, data not shown and Chai et al21) or by culture in the presence of immobilized anti-CD3 (clone B9, data not shown and Marelli-Berg et al22). As shown in Figure 2D and 2H, injection of anergic T cells (3-5 ⫻ 106) 24 hours prior to injection of the C6 T cells prevented infiltration of the peritoneal membrane by the C6 T-cell clone, and a large proportion of the labeled T cells remained in the peritoneal cavity. This was not due to competition, as responsive T cells, which promptly infiltrate the tissue, had a minimum effect on tissue infiltration by the labeled T cells (Figure 2C,G). In addition, we had previously established that anergic T cells are unable to infiltrate the peritoneal membrane while remaining localized in the peritoneal lavage (data not shown). Labeled T cells were not detectable under any of these conditions in lymph nodes or spleen (data not shown). In keeping with our in vitro observations, the specificity of anergic T cells was irrelevant to the effect observed as tissue infiltration was prevented to the same extent by either C6 (antigen-specific) or B9 (irrelevant) anergic T cells (data not shown). Increased expression of dipeptidyl peptidase IV (DPPIV)/CD26 by anergic T cells is instrumental to the inhibition of EC function

In a search for “inhibitory” molecules expressed by anergic T cells we observed that a proportion of these displayed membrane-bound transforming growth factor ␤ (TGF␤) and IL-10 (Figure 3A-B) and that the surface enzyme DPPIV/CD26 was strongly up-regulated on the whole T-cell population (Figure 3C). Similarly, CD26 expression was induced (although at low level) in anergic but not in responsive C6 and B9 T cells, which remained negative (Figure 3D). In functional assays, while neutralizing anti-TGF␤ and/or

CD26 has been shown to metabolize and inactivate chemokines such as CXCL-12.26-28 We first tested the possibility that anergic T cells display

Figure 3. Inhibition of EC function by anergic T cells is mediated by CD26. (Panels A-D) Responsive (u) or anergic (䡺) human alloreactive T-cell lines were incubated with FITC-conjugated anti-TGF␤ (panel A), anti–IL-10 (panel B), and anti-CD26 (panel C) mAbs. Cells were then analyzed by flow cytometry. The average median fluorescence intensity of CD26 expression by anergic T cells in 3 separate experiments was 162.68 ⫾ 57.72 SD, while that of responsive T cells was 30.1 ⫾ 3.8 SD. This difference was statistically significant (P ⬍ .02). Expression of CD26 by murine responsive and anergic T cells was also compared (clone B9; panel D). The average median fluorescence intensity of CD26 expression by murine anergic T cells in 3 separate experiments was 5.6 ⫾ 0.8, while that of responsive T cells was 1.5 ⫾ 2.1 SD. This difference was statistically significant (P ⬍ .05). (Panels E-H) Responsive (105/well; panels E,G) and anergic (5 ⫻ 104/well; panels F,H) T-cell lines were treated for 30 minutes at 4°C with an anti-CD26 antibody or with an isotype-matched control mAb and washed prior to being added to TNF-␣–treated EC monolayers grown on 12-mm–diameter transwells (5 ⫻ 104/well). As a control, EC monolayers were incubated in medium alone. T cells were subsequently removed and fresh 7P.61 T cells (7 ⫻ 105/well; panels E-F) or granulocytes (106/well; panels G-H) were seeded onto the EC monolayers. The results are expressed as percentage of migrated T cells at the given time points and are reported as the average of 3 experiments of identical design. The bars show the SD. * indicates statistically significant (at least P ⬍ .03, panel F; at least P ⬍ .03, panel H) versus control cultures (EC monolayers cultured with anergic T cells).


increased chemokine metabolism. CXCL-12 (50 ng/mL) was incubated overnight at 37°C, 7% CO2 in chemotaxis medium alone, or in the presence of either responsive (106/mL) or anergic (106/mL) human alloreactive CD4⫹ T cells. In addition, chemotaxis medium alone, or containing either responsive or anergic T cells incubated overnight at 37°C 7% CO2 were used as a control. Supernatants were then obtained from each sample by centrifugation and their ability to induce migration of CXCL-12–responsive CD45RA⫹ T cells (2 ⫻ 106/well)23 was measured. Incubation in the presence of anergic T-cell lines completely abrogated the chemotactic activity of CXCL-12, while resting T cells had only a modest effect (Figure 4A-C). To establish whether increased CD26 activity was instrumental to this effect, responsive and anergic T cells were incubated with a blocking anti-CD26 antibody (1 ␮g/mL) for 30 minutes and washed prior to incubation with CXCL-12, as described in this paragraph. As it is shown in Figure 4D, loss of chemotactic activity by CXCL-12 was prevented by CD26 blockade on anergic T cells, while CD26 blockade on responsive T lymphocytes had little effect on CXCL-12–mediated chemotaxis. This effect was still evident 24 hours later, although reduced due to a loss of chemotactic gradient (Figure 4E). This observation, however, ruled out the possibility that a factor released by the anergic T cells might either be toxic to naive T cells or alter their motility. CD26 enzymatic activity mediates inhibition of tissue infiltration by anergic T cells

To establish the role of CD26 activity in the anti-inflammatory effects of anergic T cells in vivo, 3 ⫻ 106 to 5 ⫻ 106 anergic CD4⫹ B9 T cells were incubated either with a blocking anti-CD26 mAb or with an isotype-matched mAb for 30 minutes at RT before intraperitoneal injection into mice treated intraperitoneally with IFN-␥ 24 hours earlier. To exclude the possibility that CD26 signaling modified other intrinsic properties of anergic T cells, the naturally occurring peptide YY peptide (3-36) containing the X-X-Pro N-terminal motif known to inhibit DPPIV enzymatic activity was also employed12 to treat anergic T cells prior to injection (30 nM for 30 minutes at room temperature). The following day, PKH26-labeled C6 T cells (5-7 ⫻ 106) were injected intraperitoneally. Following a further 24-hour incubation, the presence of labeled T cells in the peritoneal membrane and lavage was analyzed as described in “Materials and methods.” As shown in Figure 5, pretreatment of anergic T cells with anti-CD26 mAbs (Figure 5D,J) or


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with the YY peptide (Figure 5F,L) prevented the inhibitory effect of anergic T cells on C6 T-cell infiltration of the peritoneal membrane, which appeared extensive. Pretreatment with an isotype-matched mAb resulted in minimal interference with the anti-inflammatory effect of anergic T cells (Figure 5E,K). The possibility that antibody binding to T cells could interfere with their effect on the peritoneal membrane permeability was excluded by pretreating the cells with an anti-CD4 antibody that binds CD4⫹ B9 T cells. This treatment did not affect the anti-inflammatory activity of anergic T cells (data not shown). AntiCD26 antibody or YY peptide treatment did not lead to the recovery of responsiveness or motility by anergic T cells (data not shown).

Discussion Anergic T cells can exert immunoregulation by reducing the immunogenicity of APCs they come into contact with.4,5,7 The molecules involved in anergic T-cell–mediated immune regulation remain undefined. In undertaking the present study we reasoned that, in a manner similar to the antigen-independent stage of T-cell activation and synapse formation,10,11 leukocyte transmigration and tissue infiltration require effective adhesive interactions with the vascular endothelium as well as other parenchymal components. This led to the hypothesis that contact with anergic T cells might prevent/disrupt the ability to establish cell-cell interactions. We here report that exposure to anergic T cells reduces substantially the ability of parenchymal cells to establish effective cell-cell interactions with leukocytes. This effect does not require cognate recognition of the parenchymal cells as it occurs following interaction with anergic T cells with irrelevant specificity and correlates with up-regulation/induction of CD26 (DPPIV) expression, which confers on anergic T cells the ability to metabolize chemokines. CD26 is a surface-bound dipeptidyl-peptidase capable of cleaving N-terminal dipeptides from polypeptides, including chemokines,26,27 with a proline or an alanine at second position. Albeit extensively studied, the role of CD26 in T-cell biology has not yet been clarified. CD26 has been attributed direct costimulatory activity as well as protection of T cells from adenosine-induced apoptosis.26,27 More recently, CD26 has attracted great interest due to its ability to inactivate or alter the specificity of many chemokines, including CXCL10 IFN␥-inducible protein 10 kDa (IP-10) and CXCL12,28 and has been described to be preferentially expressed by T helper-1 (Th-1)29 and T regulatory-1 (Tr-1) cells.30

Figure 4. Anergic T cells metabolize chemokines via CD26. CXCL12 was added to chemotaxis medium (RPMI 2% FCS) at a concentration of 50 ng/mL and incubated overnight at 37°C 7% CO2 in the absence of T cells (panel A) or in the presence of responsive (panel B) or anergic human CD4⫹ T cells (panel C). As a control, chemotaxis medium (RPMI 2% FCS) alone, or containing either responsive (106/mL) or anergic (106/mL) human T cells, was used. The results are expressed as percentage of migrated naive T cells after 6 hours and are reported as the average of at least 3 experiments of identical design. The bars show the SD. * indicates statistically significant (P ⬍ .005) versus control cultures (CXCL12 incubated with anergic T cells). (Panels D-E) In some experiments responsive (R) and anergic (A) T cells (106/sample) were treated with an antihuman CD26 antibody (5 ␮g/mL) for 30 minutes and washed prior to incubation with CXCL12 as described in “Anergic T cells metabolize chemokines via CD26.” Supernatant was then obtained from each sample (as specified below each column) by centrifugation and the chemotactic activity was assessed monitoring the migration of purified CD45RA⫹ T cells (2 ⫻ 106/well) through a transwell after 6 (panel D) and 24 (panel E) hours. The results are expressed as specified for panels A-C. * indicates statistically significant (P ⬍ .006, panel D; P ⬍ .001, panel E) versus control cultures (CXCL12 incubated with anergic T cells).



JAMES et al

Figure 5. CD26 activity mediates the anti-inflammatory properties of anergic T cells in vivo. Untreated female mice (negative control; panels A,G) and IFN-␥–treated male mice (positive control; panels B,H) were injected with an equal volume of sterile saline containing no T cells. Alternatively, IFN-␥–treated male mice were injected with either untreated anergic B9 T cells (3-5 ⫻ 106/mouse; panels C,I), or preincubated with an antimouse CD26 mAb (5 ␮g/mL; panels D,J), or with an isotype-matched control (5 ␮g/mL; panels E,K), or with 30nM human YY peptide (3-36) (panels F,L) for 30 minutes at room temperature and subsequently washed. The following day, fluorescently labeled HY-specific C6 T cells (5-7 ⫻ 106) were injected intraperitoneally. Following a further 24-hour incubation, the presence of red-fluorescent T cells in the peritoneal membrane (panels A-F) and lavage (panels G-L), as well as the lymph nodes and spleen (data not shown), was assessed. The median number and range of labeled cells quantified in 10 randomly selected ⫻ 40-magnified fields of the peritoneal membrane of at least 3 different animals are shown below each image (panels A-D), which is taken from a representative experiment. The median percentage of labeled cells recovered in the peritoneal lavage of 3 different samples is specified below each representative histogram (in which the percentage of positive cells is indicated). * indicates statistically significant (P ⬍ .0001 for both panel D and J) compared with anergic T cells treated with isotype-matched control mAbs or with untreated anergic T cells; **, statistically significant (P ⬍ .0001, panel F; P ⬍ .05, panel L) compared with untreated anergic T cells.

Several explanations can account for our observation that although CD26 is expressed at significant levels also by responsive T cells, in this form it is not inhibitory. First, it might be simply a quantitative effect. Second, it is known that CD26 activity is increased by polymerization.31,32 It is possible that this molecule is expressed in a more “active” form by anergic T cells. Third, other DPPIV molecules (such as attractin33) that share antigenic determinants with CD26 (and thus might not be discriminated by antibodies33) can be expressed by anergic T lymphocytes. Finally, CD26 might be required for the inhibitory effect but another factor/molecule specifically expressed by anergic T cells is also needed. Further studies will be required to clarify this issue. The most likely mechanism by which CD26 enzymatic activity exerts its effect in our system is by metabolizing cell surface– bound chemoattractants.34 In support of this hypothesis, we have shown that anergic T cells reduce CXCL12 activity and that this effect is partly prevented by CD26 blockade. Other surface-bound enzymes with similar properties, such as cathepsin-G35 and attractin,36 which are expressed on the T-cell surface where they are metabolically active, might be involved in this effect. Interestingly, the T-cell surface enzyme attractin (DPPT-L) that shares many properties with CD26, including the regulation of T-cell–APC interactions, had been hypothesized to regulate T-cell activation by down-regulating chemokine/cytokine activity by proteolytic modification following the induction of cluster formation.33,36 These opposing effects on T-cell activation are probably determined by the balance of membrane-expressed to soluble attractin.33 How may this mechanism contribute to physiologic immunoregulation in vivo? The hypothesis that we have promoted is that antigen presentation by tissue parenchymal cells leads to the

generation of immunoregulatory anergic T cells and that this is a key event in the control of autoimmunity induced as a byproduct of ongoing inflammation.37 This hypothesis is supported by numerous observations of anergy induction by costimulation-deficient antigenpresenting cells37 including epithelial cells.38,39 This is likely to be a prominent event, in vivo, once tissue inflammation is well established and parenchymal cell MHC class II expression has been induced. In addition, increasing evidence suggests that T-cell anergy may arise in vivo as a result of homeostatic mechanisms involving T-T antigen presentation (R.I.L., unpublished observations, January 1999) and CTLA-4 ligation.40 In this context, newly generated anergic T cells may contribute to the containment of the inflammatory process by metabolizing chemoattractants within the tissue, thus reducing tissue infiltration. Our observations that anergic T cells are immobile and remain localized and die in the site where they have been generated make it unlikely that anergic T cells can exert chemokine metabolism at the luminal side of the endothelium (ie, leukocyte transendothelial migration). Finally, chemokine metabolism may contribute to reducing the immunogenicity of tissue-resident APCs by rendering them unable to induce polarization and adhesion of interacting T cells. This possibility remains to be tested.

Acknowledgments We thank C. Rudd and A. George for critical review of the manuscript and S. Puhalla for her help with wide-field fluorescence microscopy.

References 1. Schwartz R H. Models of T cell anergy: is there a common molecular mechanism? J Exp Med. 1996;184:1-8.

tion of cytotoxic T lymphocyte generation by two distinct suppressor-cell systems. Ann N Y Acad Sci. 1988;532:177-198.

2. Lechler R, Chai JG, Marelli-Berg F, Lombardi G. The contributions of T-cell anergy to peripheral T-cell tolerance. Immunology. 2001;103:262-269.

4. Lombardi G, Sidhu S, Batchelor R, Lechler R. Anergic T cells as suppressor cells in vitro. Science. 1994;264:1587-1589.

3. Battisto JR, Gautam SC, Chow KN. Down-regula-

5. Frasca L, Carmichael P, Lechler R, Lombardi G.

Anergic T cells effect linked suppression. Eur J Immunol. 1997;27:3191-3197. 6. Hirsch R, Eckhaus M, Auchincloss H Jr, Sachs DH, Bluestone JA. Effects of in vivo administration of anti-T3 monoclonal antibody on T cell function in mice, I: immunosuppression of transplantation responses. J Immunol. 1988;140:3766-3772.



7. Chai JG, Bartok I, Chandler P, et al. Anergic T cells act as suppressor cells in vitro and in vivo. Eur J Immunol. 1999;29:686-692.

18. Scott DM, Ehrmann IE, Ellis PS, et al. Identification of a mouse male-specific transplantation antigen, H-Y. Nature. 1995;376:695-698.

8. Taams LS, van Rensen AJ, Poelen MC, et al. Anergic T cells actively suppress T cell responses via the antigen-presenting cell. Eur J Immunol. 1999;28:2902-2912.

19. Scott D, Addey C, Ellis P, et al. Dendritic cells permit identification of genes encoding MHC class II-restricted epitopes of transplantation antigens. Immunity. 2000;12:711-720.

9. Vendetti S, Chai J-G, Dyson J, Simpson E, Lombardi G, Lechler R. Anergic T cells inhibit the antigen-presenting function of dendritic cells. J Immunol. 2000;165:1175-1181.

20. Chai J-G, Lechler RI. Immobilized anti-CD3 mAb induces anergy in murine naive memory CD4⫹ T cells in vitro. Int Immunol. 1997;9:935-944.

10. Krummel MF, Davis MM. Dynamics of the immunological synapse: finding, establishing and solidifying a connection. Curr Opin Immunol. 2002; 14:66-74. 11. Bromley SK, Burack WR, Johnson KG, et al. The immunological synapse. Annu Rev Immunol. 2001;19:375-396. 12. Wrenger S, Reinhold D, Hoffmann T, et al. The N-terminal X-X-Pro sequence of the HIV-1 Tat protein is important for the inhibition of dipeptidyl peptidase IV (DP IV/CD26) and the suppression of mitogen-induced proliferation of human T cells. FEBS Lett. 1996;383:145-149. 13. Cobbold SP, Jayasuriya A, Nash A, Prospero TD, Waldmann H. Therapy with monoclonal antibodies by elimination of T-cell subsets in vivo. Nature. 1984;312:548-551. 14. Frasca L, Marelli-Berg F, Imami N, et al. Interferon-gamma-treated renal tubular epithelial cells induce allospecific tolerance. Kidney Int. 1998;53: 679-689. 15. Jaffe EA, Nachman RL, Becker CG, Minick CR. Culture of human endothelial cells derived from umbilical veins: identification by morphologic and immunologic criteria. J Clin Invest. 1973;52:27452756. 16. Marelli-Berg FM, Hargreaves RE, Carmichael P, Dorling A, Lombardi G, Lechler RI. Major histocompatibility complex class II-expressing endothelial cells induce allospecific nonresponsiveness in naive T cells. J Exp Med. 1996;183:16031612. 17. Sidhu S, Deacock S, Bal V, Batchelor JR, Lombardi G, Lechler RI. Human T cells cannot act as autonomous antigen-presenting cells, but induce tolerance in antigen-specific alloreactive responder cells. J Exp Med. 1992;176:875-880.

21. Chai J-G, Bartok I, Scott D, Dyson J, Lechler R. T:T antigen presentation by activated murine CD8⫹ T cells induces anergy and apoptosis. J Immunol. 1998;160:3655-3665. 22. Marelli-Berg FM, Frasca L, Imami N, Lombardi G, Lechler RI. Lack of T cell proliferation without induction of nonresponsiveness after antigen presentation by endothelial cells. Transplantation. 1999;68:280-287. 23. Colantonio L, Iellem A, Clissi B, et al. Upregulation of integrin alpha6/beta1 and chemokine receptor CCR1 by interleukin-12 promotes the migration of human type 1 helper T cells. Blood. 1999;94:2981-2989. 24. Hollenbaugh D, Mischel-Petty N, Edwards CP, et al. Expression of functional CD40 by vascular endothelial cells. J Exp Med. 1995;182:33-40. 25. Karmann K, Hughes CC, Schechner J, Fanslow WC, Pober JS. CD40 on human endothelial cells: inducibility by cytokines and functional regulation of adhesion molecule expression. Proc Natl Acad Sci USA. 1995;92:4342-4346. 26. De Meester I, Korom S, Van Damme J, Scharpe S. CD26, let it cut or cut it down. Immunol Today. 1999;20:367-375. 27. Gorrell MD, Gysbers V, McCaughan GW. CD26: a multifunctional integral membrane secreted protein of activated lymphocytes. Scand J Immunol. 2001;54:249-264. 28. Lambeir AM, Proost P, Durinx C, et al. Kinetic investigation of chemokine truncation by CD26/ dipeptidyl peptidase IV reveals a striking selectivity within the chemokine family. J Biol Chem. 2001;276:29839-29845. 29. Rogge L, Bianchi E, Biffi M, et al. Transcript imaging of the development of human T helper cells using oligonucleotide arrays. Nat Genet. 2000;25: 96-101.

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30. Cavani, A, Nasorri F, Prezzi C, Sebastiani S, Albanesi C, Girolomoni G. Human CD4⫹ T lymphocytes with remarkable regulatory functions on dendritic cells and nickel-specific Th1 immune responses. J Invest Dermatol. 2000;114:295-302. 31. Smith RE, Reynolds CJ, Elder EA. The evolution of proteinase substrates with special reference to dipeptidylpeptidase IV. Histochem J. 1992;24: 637-647. 32. Bermpohl F, Loster K, Reutter W, Baum O. Rat dipeptidyl peptidase IV (DPP IV) exhibits endopeptidase activity with specificity for denatured fibrillar collagens. FEBS Lett. 1998;428:152-156. 33. Duke-Cohan JS, Tang W, Schlossman SF. Attractin: a cub-family protease involved in T cell-monocyte/macrophage interactions. Adv Exp Med Biol. 2000;477:173-185. 34. Salvucci O, Yao L, Villalba S, Sajewicz A, Pittaluga S, Tosato G. Regulation of endothelial cell branching morphogenesis by endogenous chemokine stromal-derived factor-1. Blood. 2002;99: 2703-2711. 35. Delgado MB, Clark-Lewis I, Loetscher P, et al. Rapid inactivation of stromal cell-derived factor-1 by cathepsin G associated with lymphocytes. Eur J Immunol. 2001;31:699-707. 36. Duke-Cohan JS, Gu J, McLaughlin DF, Xu Y, Freeman GJ, Schlossman SF. Attractin (DPPT-L), a member of the CUB family of cell adhesion and guidance proteins, is secreted by activated human T lymphocytes and modulates immune cell interactions. Proc Natl Acad Sci USA. 1998;95: 11336-11341. 37. Marelli-Berg FM, Lechler RI. Antigen presentation by parenchymal cells: a route to peripheral tolerance? Immunol Rev. 1999;172:297-314. 38. Marelli-Berg FM, Weetman A, Frasca L, et al. Antigen presentation by epithelial cells induces anergic immunoregulatory CD45RO⫹ T cells deletion of CD45RA⫹ T cells. J Immunol. 1997;159: 5853-5861. 39. Marelli-Berg FM, Barroso-Herrera O, Lechler RI. Recently activated T cells are costimulation-dependent in vitro. Cell Immunol. 1999;195:18-27. 40. Perez VL, Van Parijs L, Biuckians A, Zheng XX, Strom TB, Abbas AK. Induction of peripheral T cell tolerance in vivo requires CTLA-4 engagement. Immunity. 1997;6:411-417.

From bloodjournal.hematologylibrary.org by guest on July 13, 2011. For personal use only. CLINICAL OBSERVATIONS, INTERVENTIONS, AND THERAPEUTIC TRIALS

Prolonged treatment with rituximab in patients with follicular lymphoma significantly increases event-free survival and response duration compared with the standard weekly ⫻ 4 schedule Michele Ghielmini, Shu-Fang Hsu Schmitz, Sergio B. Cogliatti, Gabriella Pichert, Jo¨rg Hummerjohann, Ursula Waltzer, Martin F. Fey, Daniel C. Betticher, Giovanni Martinelli, Fedro Peccatori, Urs Hess, Emanuele Zucca, Roger Stupp, Tibor Kovacsovics, Claudine Helg, Andreas Lohri, Mario Bargetzi, Daniel Vorobiof, and Thomas Cerny, for the Swiss Group for Clinical Cancer Research (SAKK)

The potential benefits of extended rituximab treatment have been investigated in a randomized trial comparing the standard schedule with prolonged treatment in 202 patients with newly diagnosed or refractory/relapsed follicular lymphoma (FL). All patients received standard treatment (rituximab 375 mg/m2 weekly ⴛ 4). In 185 evaluable patients, the overall response rate was 67% in chemotherapynaive patients and 46% in pretreated cases (P < .01). Patients responding or with stable disease at week 12 (n ⴝ 151) were randomized to no further treatment

or prolonged rituximab administration (375 mg/m2 every 2 months for 4 times). At a median follow-up of 35 months, the median event-free survival (EFS) was 12 months in the no further treatment versus 23 months in the prolonged treatment arm (P ⴝ .02), the difference being particularly notable in chemotherapy-naive patients (19 vs 36 months; P ⴝ .009) and in patients responding to induction treatment (16 vs 36 months; P ⴝ .004). The number of t(14;18)–positive cells in peripheral blood (P ⴝ .0035) and in bone marrow (P ⴝ .0052) at baseline was pre-

dictive for clinical response. Circulating normal B lymphocytes and immunoglobulin M (IgM) plasma levels decreased for a significantly longer time after prolonged treatment, but the incidence of adverse events was not increased. In patients with FL, the administration of 4 additional doses of rituximab at 8-week intervals significantly improves the EFS. (Blood. 2004;103:4416-4423)


Introduction Rituximab is the first commercially available monoclonal antibody for the treatment of lymphoma. It is a chimeric anti-CD20 immunoglobulin (Ig) comprising a human IgG1 and ␬ constant region with murine variable regions. Its mechanisms of action are thought to be complement-mediated lysis; antibody-dependent, cell-mediated cytotoxicity; and induction of apoptosis after binding to the CD20 antigen on the cell surface. As one of the most effective single-agent treatments for non-Hodgkin lymphoma (NHL) developed in the last decade, rituximab has rapidly become a common treatment for this malignancy, either as a single agent or, more recently, in combination with chemotherapy. Rituximab used as a single agent is very active in several types of indolent lymphomas with response rates (RRs) of 45% to 60% even in pretreated patients,1-3 while in more aggressive types of NHL RRs are approximately 30%.4,5 When added to chemotherapy, rituximab significantly improves the RR and in some studies has been shown to improve survival rates compared with chemotherapy alone.6,7 The standard rituximab treatment schedule (375 mg/m2 each week for 4 weeks) was based on the results of 2 phase 1 trials.8,9 Further trials using this schedule in patients with low-grade

lymphoma confirmed its clinical activity, achieving a RR of approximately 50% and a response duration of 10 to 12 months.1,2 However, the relatively low proportion of complete responses (in the range of 5%) gave rise to the suggestion that this standard 375 mg/m2 weekly ⫻ 4 schedule might be enhanced to optimize the clinical efficacy of rituximab. In addition, a significant correlation between the median antibody concentration in blood and response to treatment has been observed.10 Finally, for the majority of anticancer drugs a treatment duration of several months is needed to attain the optimal result, and this could also apply to rituximab. These considerations led us to propose that evaluation of various dosing schedules was warranted and that extended exposure to rituximab would be likely to augment its biologic effect and may thus achieve a higher partial and complete RR, longer time to progression, and longer duration of response. The present trial was therefore designed to test this hypothesis. We compared the standard schedule (375 mg/m2 weekly ⫻ 4) with a prolonged rituximab schedule in which the standard treatment was followed by a single 375-mg/m2 infusion every 2 months for 4 times. The purpose was to maintain a biologically active serum concentration

From the Oncology Institute of Southern Switzerland, Bellinzona, Switzerland; Swiss Institute for Applied Cancer Research (SIAK) Coordinating Centre, Bern, Switzerland; Swiss Reference Centre for Malignant Lymphoma, Kantonsspital, St Gallen, Switzerland; Universita¨tsspital Zu¨rich, Switzerland; Universita¨tsspital Bern, Switzerland; Istituto Europeo di Oncologia, Milano, Italy; Kantonsspital, St Gallen, Switzerland; Centre Hospitalier Universitaire Vaudois (CHUV), Lausanne, Switzerland; University Hospital, Geneva, Switzerland; Universita¨tsspital Basel, Basel, Switzerland; Kantonsspital Aarau, Aarau, Switzerland; and Sandton Oncology Centre, Johannesburg, South Africa.

Supported in part by research funding from Roche Pharma Schweiz AG to the Swiss Group for Clinical Cancer Research (SAKK).

Submitted October 14, 2003; accepted February 8, 2004. Prepublished online as Blood First Edition Paper, February 19, 2004; DOI 10.1182/blood-2003-10-3411.

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A complete list of the members of SAKK appears in the “Appendix.” Reprints: Michele Ghielmini, Oncology Institute of, Southern Switzerland, Ospedale San Giovanni, 6500 Bellinzona, Switzerland; e-mail: mghielmini @ticino.com. The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734. © 2004 by The American Society of Hematology

BLOOD, 15 JUNE 2004 䡠 VOLUME 103, NUMBER 12

From bloodjournal.hematologylibrary.org by guest on July 13, 2011. For personal use only. BLOOD, 15 JUNE 2004 䡠 VOLUME 103, NUMBER 12

for approximately 1 year. The 2-month interval was chosen based on preliminary pharmacokinetics data that demonstrated the persistence of potentially active drug levels in the serum of the majority of patients after this interval.10 To avoid randomizing rituximabresistant patients into the prolonged treatment arm, the randomization was implemented after the end of the standard treatment (week 12 after initiation), when patients had proved not to be resistant to rituximab.

Patients and methods Trial design Trial sites. Patients were enrolled between January 1998 and February 2001 in 25 institutions in Switzerland, Italy, and South Africa. The trial was approved by the local ethics committee of each participating institution and conducted in accordance with the Declaration of Helsinki and currently applicable amendments. All patients gave written informed consent. Treatment. All patients were initially treated with rituximab 375 mg/m2 per week for 4 weeks (“induction” phase). The drug was to be administered every 7 ⫾ 1 days. Patients with stable disease (SD) or who were in partial (PR) or complete remission (CR) at week 12 were randomized in a 1:1 ratio to no further treatment (arm A, “no further treatment”) or to treatment with a single infusion of rituximab 375 mg/m2 at week 12 and again at months 5, 7, and 9 (arm B, “prolonged treatment”). The randomization was stratified according to status of disease at trial entry (first presentation vs refractory or relapsed disease), response to induction treatment (stable disease vs response), and center, using the minimization method.11(pp84-86) Patients were randomized centrally by fax via the SAKK Trials Office in Bern. Upon progression or relapse, further treatment was at the treating physician’s discretion. Trial end points. Event-free survival (EFS) time was the primary end point and was calculated as the time from first induction infusion to progression, relapse, second tumor, or death from any cause. Remission duration was defined in the same way, but restricted to responding patients. Sample size and interim analyses. For the randomized phase of the trial, a group sequential design with 2 interim analyses and 1 final analysis using EFS as the primary endpoint was adopted. The overall type I error probability was 5% and the statistical power was 80% for the 2-sided log-rank test, to detect an increase of EFS from 9 months in the no further treatment arm to 16.2 months in the prolonged treatment arm. A total of 99 events in about 135 patients were required for the final analysis. The first interim analysis was performed after 26 events in 59 patients and the second after 69 events in 132 randomized patients. Neither interim analysis called for early stopping of the trial. Patients Inclusion criteria were a biopsy-proven CD20⫹ follicular lymphoma grade I, II, or III according to the REAL classification, age of 18 years or older, and measurable disease defined as the presence of at least one previously unirradiated lesion with 2 measurable perpendicular diameters of which at least one should be of 2 cm. The interval since the last systemic anticancer treatment should not be shorter than 28 days (6 weeks if treated with nitrosureas). Patients should not have received corticosteroids within 28 days prior to trial entry unless chronically administered at a dosage of no more than 20 mg/day for indications other than lymphoma or lymphomarelated symptoms. Other inclusion criteria were an Eastern Cooperative Oncology Group (ECOG) performance status of 2 or less and, following assessment of the first 100 patients, in some of whom unexpected cardiac events had been observed,12 a cardiac ejection fraction (EF) of at least 50% as determined by echocardiography. Exclusion criteria included transformation to high-grade lymphoma, symptomatic central nervous system disease, prior antibody-based therapy, or a history of significant medical conditions, including previous malignancies within 5 years. Other exclusion criteria were a reduced renal function (creatinine ⬎ 2 times the upper limit of normal [ULN]) or liver function (bilirubin ⬎ 2 ⫻ ULN). Pregnant or

PROLONGED RITUXIMAB TREATMENT

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lactating females, patients with active opportunistic infections, or patients with known HIV or hepatitis B or C infections were also excluded. Trial assessments Pretreatment evaluation. For all patients, a complete staging, including clinical examination, chest and abdomen computed tomography (CT) scan, bone marrow (BM) aspirate, and trephine biopsy, should be performed not more than 2 weeks before entering the trial. Any other clinically indicated tumor evaluations were at the discretion of the investigator. A central histology review was performed between registration and randomization to confirm the diagnosis. Assessment of clinical response. A formal evaluation of all the involved sites was performed at 12 weeks and at 7, 12, 18, and 24 months, and then yearly or when clinically required. The first 100 patients had an additional staging procedure at week 8, but the first interim analysis showed similar RRs at weeks 8 and 12 and it was decided to limit the first restaging at week 12 for subsequent patients. Re-evaluation of the BM was required only at week 12 and month 12 if involved at trial entry. Blood analyses. Routine blood counts and chemistries were assessed at baseline, before each rituximab administration, and at months 2, 3, 5, 7, 9, and 12. Serum immunoglobulins (IgG, IgA, IgM) were measured at baseline and again at months 3, 7, and 12, while blood samples for immunophenotyping were taken at baseline, week 12, and months 9, 12, 18, and 24. For the analysis of lymphocyte subsets, dimethyl sulfoxide (DMSO)–frozen cells were thawed and washed, stained with fluorescent antibodies, and analyzed on a FacsCalibur machine (Becton Dickinson, San Jose, CA) using the Multiset program. The antibodies used were anti-CD3, -CD4, -CD8, -CD19, -CD16 ⫹ 56, and -CD45. With a 4-color analysis on the lymphocyte gate, the following cells were counted: T cells (CD3⫹), T helper (CD3⫹/CD4⫹), T suppressor (CD3⫹/CD8⫹), natural killer (NK; CD3⫺/CD16⫹56⫹), and B cells (CD19⫹). Molecular analysis. The presence of cells carrying the t(14;18) translocation was investigated at baseline in peripheral blood (PB) and BM using nested polymerase chain reaction (PCR)13; if the result was positive, the analysis was repeated at week 12 and month 12 for BM, while in the PB it was evaluated each time that a radiologic or clinical staging was performed. For the quantification of t(14;18) translocation– carrying cells with a major breakpoint region (MBR), real-time PCR analysis was performed using an ABI Prism 7700 Sequence Detection System (Applied Biosystems, Foster City, CA). For the detection of the t(14;18) translocation, 900 nM of the forward and reverse primer (identical to the internal primers used in the conventional PCR) and 300 nM of 5⬘-FAM-TTTCAACACAGACCCACCCAGAGCC-TAMRA-3⬘ as internal TaqMan probe were used; for the simultaneous amplification of the glyceraldehyde phosphate dehydrogenase (GAPDH) gene, 900 nM 5⬘-CTGGCACCCTATGGACACG-3⬘ as forward primer, 300 nM 5⬘-GGATGAGAAAGGTGGGAGCC-3⬘ as reverse primer, and 300 nM 5⬘-FAM-CCTGCACCACCAACTGCTTAGCACC-TAMRA-3⬘ as internal TaqMan probe were used. A total of 400 ng of genomic DNA, corresponding to 60 000 cells, was used in a final volume of 50 ␮L. For each sample the amplification of the MBR target gene and the GAPDH reference gene was performed in quintuplicates. The thermal cycling conditions comprised an initial uracil DNA glycosylase (UNG) Erase activation step at 60°C for 2 minutes, a denaturation step at 95°C for 10 minutes, and 50 cycles, each one consisting of 15 seconds at 95°C and 1 minute at 60°C. Genomic DNA from the t(14;18)–positive cell line Karpas 422,14 diluted into genomic DNA from t(14;18)–negative mononuclear cells (MNCs), was used to generate a standard curve for the t(14;18) translocation. After demonstrating that the amplification efficiencies of the target and reference gene were approximately equal (data not shown), the comparative ⌬ cycle threshold (⌬Ct) method (⌬Ct ⫽ Ct[t(14;18)] ⫺ Ct[GAPDH]) was used to calculate the t(14; 18)–positive cell numbers in each sample.15 The calculated ⌬Ct standard curve showed an excellent correlation coefficient (r ⫽ 0.998) and slope (⫺3.29). The real-time PCR assay is highly reproducible with an interassay variation of 13% or less. At least 30 t(14;18)–positive cells per 106 MNCs could reproducibly be detected, which is in the range of



GHIELMINI et al

other reports.16 If, in samples with 30 or less t(14;18)–positive cells per 106 MNCs, some of the 5 replicates failed to give a positive signal they were scored as 0.17 Toxicity. The evaluation of toxicity was conducted according to the World Health Organization toxicity grading. Statistical methods The comparisons between treatment arms or groups defined by other factors (as previously treated vs chemotherapy-naive or responders vs nonresponders) were carried out by Wilcoxon rank sum test for continuous variables and by chi-square or Fisher exact test for categoric variables. Odds ratios (ORs) were also calculated for associations between a binary response variable and a binary patient characteristic. Remission duration and EFS time were estimated by the Kaplan-Meier method and compared between groups by log-rank test. Hazard ratios (HRs) were also calculated as the ratio of hazard of experiencing an event in the nonreference group with respect to the reference group. The impact of some potential predictive variables on clinical response and EFS was analyzed by multiple logistic regression and multiple Cox regression, respectively, employing a stepwise model selection procedure. All tests were 2-sided. No adjustment for multiple comparisons in general was performed, except for the primary end point for which adjustment for multiple testing prescribed by the group sequential design was performed. For randomized patients, all between-arm comparisons were carried out by intention-to-treat principle.

Results Patients’ characteristics

A total of 202 patients were enrolled; their characteristics are summarized in Table 1. Seventeen patients were retrospectively judged ineligible for the trial: the pathology review could not confirm the follicular histology in 13 patients, while 4 patients did not have measurable disease as required per protocol. A further 3 patients had their last treatment less than 28 days prior to induction but this was considered a minor protocol violation and they were retained in the analysis. Of the 185 eligible and evaluable patients, 35 were not randomized to the second phase of the trial: 26 due to disease progression during the induction phase, 5 due to major toxicity, and 4 due to other reasons. One patient was randomized prior to being found ineligible. The clinical and pathologic characteristics of the 151 randomized patients (78 in arm A and 73 in arm B) were similar in the 2 arms (Table 1) and, as expected, more favorable at randomization than at baseline. There was a tendency toward lower neutrophil counts at randomization. The proportions of patients having responded to any previous treatments and to rituximab induction treatment were similar for arms A and B. The only significant differences in baseline characteristics between the 2

Table 1. Patients’ characteristics Randomization Induction

No further treatment; arm A

Prolonged treatment; arm B

Categoric variable (% of patients) ECOG performance status 0

147 (73)

70 (90)

60 (82)

1

43 (21)

8 (10)

11 (15)

2

12 (6)

0

1 (1)

Sex Male Female

87 (43)

(37)

33 (45)

115 (57)

(63)

40 (55)

Stage I

6 (3)

3 (4)

3 (4)

II

23 (11)

7 (9)

10 (14)

III

59 (29)

27 (35)

24 (33)

IV

114 (56)

41 (53)

36 (49)

57 (28)

23 (30)

18 (25)

B symptoms Bone marrow involvement

105 (52)

20 (26)

8 (11)

Bulky disease, 5 cm or more

106 (53)

17 (22)

22 (30)

Elevated LDH

74 (37)

14 (18)

16 (22)

Chemotherapy naive

64 (32)

26 (33)

25 (34)

130 (64)

49 (63)

46 (63)

I

68 (34)

24 (31)

31 (43)

II

90 (45)

40 (51)

33 (45)

III

25 (12)

12 (15)

6 (8)

Responders to previous chemotherapy REAL histology grade

Continuous variable, median (range) Age, y*

57 (28-81)

57 (28-81)

56 (31-79)

Disease duration, y*

1.5 (0-21)

1.5 (0-21)

1.1 (0-17)

No. of previous regimens, pretreated patients*

2 (1-4)

2 (1-4)

2 (1-4)

No. of cycles of chemotherapy, pretreated patients*

12 (3-56)

10 (4-33)

10 (3-56)

Interval since last treatment, pretreated patients, mo*

7.5 (0.4-179)

7.2 (0.9-67)

8.5 (0.7-179)

Neutrophil count, ⫻ 109/L

3.6 (0.1-11.1)

3.7 (0.4-7.4)

3.2 (0.1-9.7)

B-cell count, ⫻ 109/L

81 (0-3629)

18 (0-239)

23 (0-225)

132 (8-773)

161 (16-828)

175 (38-469)

25 (1-90)

30 (4-90)

16 (5-90)

NK-cell count, ⫻ 109/L Bone marrow infiltration, involved only, %

Patient populations are as follows: induction, n ⫽ 202; no further treatment, arm A, n ⫽ 78; and prolonged treatment, arm B, n ⫽ 73. ECOG indicates Eastern Cooperative Oncology Group; LDH, lactate dehydrogenase; and REAL, Revised European-American Classification of Lymphoid Neoplasms. *The results for randomized patients reflect characteristics at baseline, not at randomization.


arms were in lymphocyte subsets: NK- and B-cell counts were lower in arm A. Description of treatment

Induction treatment. All except 5 patients received 4 infusions of rituximab 375 mg/m2. Four patients received only 1 infusion (1 patient died of a sudden cardiac event, 3 patients experienced infusion-related toxicity), and 1 patient did not receive the fourth infusion due to excessive infusion-related toxicity during the 3 previous infusions. The intervals between infusions in the induction phase were maintained in the planned range except in 2 patients. Reduced doses were administered to 2 patients. Prolonged treatment. The majority of patients in arm B received the treatment according to the protocol. Treatment was discontinued before completing the planned schedule (4 infusions in 6 months) in 18 patients: 2 due to toxicity and 16 due to disease progression or relapse. In 1 patient the third prolonged treatment infusion was given at a reduced dose due to toxicity and in another patient the fourth prolonged treatment infusion was partially administered subcutaneously. Objective response

Induction phase. At week 12 the RR for all 185 evaluable patients who received treatment was 52% (95% confidence interval [CI]: 45%-60%) with 8% CR. There was a significant difference between the RR of the 57 chemotherapy-naive patients (RR 67% with 9% CR) and the 128 previously treated patients (RR 46% with 8% CR) (OR ⫽ 2.34; P ⫽ .0097). In addition to being chemotherapy naive, other favorable predictive factors identified by univariate analyses in evaluable patients included female sex (OR ⫽ 1.85; P ⫽ .040), disease diameter less than 5 cm (OR ⫽ 2.60; P ⫽ .0015), hemoglobin level at least 120 g/L (12 g/dL; OR ⫽ 2.12; P ⫽ .026), platelet count at least 100 ⫻ 109/L (OR ⫽ 2.68; P ⫽ .036), IgA level higher than 1.2 g/L (median) (OR ⫽ 2.24; P ⫽ .021), and IgM level higher than 0.6 g/L (median) (OR ⫽ 3.19; P ⫽ .0012). A multivariate analysis (due to missing values, only 149 patients were included in the model selection procedure) identified 2 significant independent predictors for clinical response (complete response [CR] and PR): previous chemotherapy (OR ⫽ 0.48; P ⫽ .05) and bulky disease (OR ⫽ 0.5; P ⫽ .039). Of the 97 evaluable patients who had BM involvement at baseline, 69 had BM re-evaluation at week 12 with 34 (49%) becoming morphologically negative. It is likely that the lack of BM evaluations for 28 patients has resulted in an underestimation of the CR rate, since patients assessed as achieving CR on the basis of clinical and radiologic evaluation, but who did not have a re-evaluation of a previously positive BM, were considered to have had a PR. Postrandomization phase. Progressive disease was documented at around week 12 in 5 patients (4 in arm A, with no further treatment, and 1 in arm B, with prolonged treatment). The response assessments at week 12 and at months 12 and 24, as well as the overall best response rate (from randomization to last follow-up) for randomized patients, are summarized in Figure 1. In arm A the proportion of patients responding decreased steadily from 67% at week 12 to 44% at month 12 and 28% at month 24, while in arm B the proportion of responders remained initially stable, with 62% at week 12 and 60% at month 12, declining to 45% at month 24. In contrast, the proportion of patients in CR increased constantly in both arms, rising from 8% at week 12 to 19% at month 24 in arm A and from 12% to 29% in arm B. The number of patients improving their response in the first 2 years (from PR to CR, from SD to CR,


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or from SD to PR) was similar in both arms: 21 cases in arm A and 22 cases in arm B. Five patients in arm A and 8 patients in arm B had their first CR documented after month 12. The between-arm difference in RR was not statistically significant at weeks 12 and month 7 but became significant after 1 year (P ⫽ .046). The between-arm difference in CR rate was not statistically significant at any time point. Among responders at week 12, the proportions of patients remaining in CR or PR at later time points were significantly lower in the no further treatment arm compared with the prolonged treatment arm: week 52, 56% versus 80% (P ⫽ .011); month 18, 40% versus 67% (P ⫽ .0097); and month 24, 35% versus 58% (P ⫽ .022). The overall best response was 77% (31% CR) in arm A and 75% (38% CR) in arm B (not statistically significant [NS]). The overall best response for chemotherapy-naive patients was 81% (31% CR) in arm A and 92% (52% CR) in arm B (NS). Among patients who reached a PR or CR, the median time to first response was 10 weeks in arm A and 9 weeks in arm B. The median remission duration among week-12 responders was 16 months in arm A and 36 months in arm B (P ⫽ .0039). The difference in remission duration was considerable and marginally significant for chemotherapy-naive versus pretreated patients (Figure 2A-B). Among patients who had BM involvement at baseline and BM re-evaluation during the postrandomization phase, 13 of 17 (77%) in arm A and 17 of 22 (77%) in arm B became morphologically negative for BM involvement. Event-free survival

The median follow-up time in 126 living patients was 36 months. The median EFS time among 151 randomized patients was 11.8 months in arm A and 23.2 months in arm B (HR ⫽ 0.61, 95% CI: 0.40-0.93; P ⫽ .024, adjusted for multiple testing prescribed by the group sequential design; Figure 3). The difference was more pronounced in chemotherapy-naive patients, with a median EFS of 19 months in the no further treatment arm versus 36 months in the prolonged treatment arm (P ⫽ .009; Table 2). When EFS was stratified by response to the induction phase (Table 2), the prolonged treatment more than doubled the median EFS (from 16 to 36 months; P ⫽ .004) in responding patients, while the effect was not significant in patients with SD at first restaging (from 8 to 11 months; P ⫽ .35). In univariate analyses not taking treatment effect into account, baseline characteristics significantly prognostic for EFS included disease diameter, Ann Arbor stage, previous radiotherapy or

Figure 1. PR and CR rates (at intervals during follow-up and overall) in both study arms. The same proportion of patients responded in each arm and the number of complete responders gradually increased. The response rate remained elevated for a longer time in patients receiving prolonged treatment.



GHIELMINI et al

Table 2. Median event-free survival (months) stratified by previous treatment and by response to induction therapy (151 randomized patients) Standard treatment; arm A; n ⴝ 78

Prolonged treatment; arm B; n ⴝ 73

P

Chemotherapy naive

19

36

.009

Pretreated

10

15

.081

Responders (PR ⫹ CR)*

16

36

.004

8

11

.35

Stable disease

PR indicates partial response; and CR, complete response. *As evaluated at week 12.

Figure 2. Duration of response by study arm. Median duration of response is more than double with the prolonged schedule, independently of previous treatment, in (A) chemotherapy-naive and (B) pretreated patients. Differences are only marginally significant due to the small number of patients in subgroups.

chemotherapy, number of previous chemotherapy regimens, and T-helper cell count. Characteristics at randomization with a prognostic impact on EFS were response at week 12, disease size (⬎ 5 cm), BM infiltration, hemoglobin level, and lymphocyte count. The multivariate analysis to identify factors independently prognostic for EFS in randomized patients (Table 3) confirmed that the prolonged treatment arm highly significantly increased EFS (HR ⫽ 0.40; P ⬍ .0001). Other independent prognostic factors included parameters related to biologic characteristics of the lymphoma (tumors with B-cell lymphoma-2 [bcl-2] immunoreactivity in ⬎ 80% of the tumor cells responded longer), amount of circulating monocytes, patient’s performance status, and response to induction treatment. Molecular response

Induction phase. In 70 patients with a major breakpoint cluster region t(14;18) in PB and/or BM analyzed by real-time PCR (analysis performed in PB only, n ⫽ 22; BM only, n ⫽ 3; BM ⫹ PB, n ⫽ 45), the median number of t(14;18)–positive cells was 374 (range, 0-579 004) per 106 MNCs in PB and 42 (range, 0-707 843) per 106 MNCs in BM. Among the 70 initially t(14;18)–positive cases after induction with rituximab, 56 and 43 patients could be re-evaluated by PCR for t(14;18) in PB and BM, respectively. Of those, 59% of PB samples and 39% of BM samples had become t(14;18)–negative; real-time PCR could be performed in only a subset of cases, and showed that the number of circulating

t(14;18)–positive cells was effectively reduced: 85% of 52 PB samples and 64% of 28 BM samples had a reduction of at least 1 log in t(14;18)–positive cell numbers, and 42% of PB and 46% of BM samples had a greater than 2 log reduction. The number of t(14;18)–positive cells in PB and BM at baseline was significantly predictive of clinical response to rituximab (Table 4). Among patients with initially positive PCR results in PB samples, clearance of circulating t(14;18)–positive cells was significantly associated with clinical response: 84% PCR negative in 32 patients with CR or PR versus 11% PCR negative in 19 patients with SD or progressive disease (PD; P ⬍ .0001). Postrandomization. Of the 70 patients with an initial PCRdetectable t(14:18) translocation, 59 were randomized. In the prolonged treatment arm, 5 of 12 PCR-positive patients became PCR negative, whereas none of 10 PCR-negative patients became PCR positive. In contrast, in the no further treatment arm, 1 of 3 PCR-positive patients became PCR negative (8 PCR-positive patients progressed so rapidly that no follow-up samples could be obtained) and 7 of 15 PCR-negative patients became PCR positive. PCR positivity was highly significantly correlated with disease progression. With both arms pooled together (due to the small number of observations), 11 of 16 patients remaining or becoming PCR positive had PD, whereas only 1 of 23 patients remaining or becoming PCR negative had PD (P ⬍ .0001). Therefore, patients who remained or became PCR positive after induction therapy with rituximab had a higher chance of disease progression in both arms. Toxicity

All 202 patients were evaluable for toxicity. The incidence of nonhematologic and hematologic grade 3 and 4 toxicity is summarized in Table 5. In the induction phase, the majority of nonhematologic toxicities were mild infusion-related symptoms during the first infusion. Sixteen cases of serious adverse events were Table 3. Independent prognostic factors for event-free survival in randomized patients (n ⴝ 121) Factor

Hazard ratio

95% CI

P

0.40

0.26-0.64

⬍ .0001

0.55

0.34-0.87

.01

0.58

0.37-0.91

.017

0.59

0.36-0.97

.039

0.41

0.26-0.65

.0001

Treatment arm Prolonged vs standard treatment Bcl-2 overexpression Greater than 80% vs 80% or less Monocyte count at baseline 0.4⫻109/L or less vs greater than 0.4⫻109/L Performance status at baseline 0 vs 1 ⫹ 2 Response to induction therapy CR ⫹ PR vs SD ⫹ PD Figure 3. EFS of all randomized patients according to study arm. The curves are parallel, suggesting that the gain acquired during the 9 months of prolonged treatment is maintained over time.

CI indicates confidence interval; CR, complete response; PR, partial response; SD, stable disease; and PD progressive disease.



Median t(14;18)-positive cells/106 MNCs Below median, % Above median, % P

Peripheral blood; n ⴝ 65

Bone marrow; n ⴝ 46

374

42

79

74

41

39

.0035

.0052

MNCs indicates mononuclear cells.

B

Number of cells x 106/L

Table 4. Proportion of responders stratified by t(14;18)-positive cell numbers at baseline

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100

A

Prolonged Standard

80 60 A

A

40 B 20 Standard A baseline 2-3

A B

B

B Prolonged

5

9

12

Months since start of treatment

documented: 2 of them were life-threatening events, 1 patient died due to a cardiac event (he had no history of cardiac disease; autopsy revealed subacute myocarditis) and 1 developed acute myeloblastic leukemia. Two further patients had severe infusion-related symptoms resulting in permanent treatment discontinuation. Serious adverse events during prolonged treatment and follow-up period were reported in 4 cases in arm A and 9 cases in arm B. These included 2 malignancies, 1 infection, and 1 neuropathy in arm A, and 2 malignancies, 2 infections, 2 complications of surgery, 1 pneumopathy, and 1 agranulocytosis in arm B. The mean values of cardiac EF assessed in 83 patients at week 12 and in 51 patients at week 52 (20 in arm A and 31 in arm B) were the same as the baseline value (n ⫽ 107). A total of 137 patients were evaluable for late toxicity beyond 1 year. The incidence was 7% in both treatment arms and included 5 cases of infection, 1 of teeth loss, 1 of asymptomatic neutropenia, 2 of leukocytopenia, and 1 of cachexia. The lymphocytes subset analysis (excluding patients with baseline white blood cell count ⬎ 10 ⫻ 109/L who were suspected of having leukemic disease) showed a very early disappearance of circulating B cells, while T-helper, T-suppressor, and NK cells remained approximately stable during the induction phase. The evolution of circulating B lymphocytes for randomized patients over the first year of follow-up is shown in Figure 4. Median B-cell levels decreased rapidly in both study arms during the induction phase. At 1 year, the level rose above the baseline value in arm A while remaining depressed in arm B (P ⫽ .002). At further follow-up (despite the small number of available data) there appears to be a recovery of B cells in both arms (data not shown). The decrease in B cells was accompanied in arm B by a reduction Table 5. Grades 3 to 4 toxicity Adverse event

No. of patients with grades 3 to 4 toxicity (%)

Induction phase, n ⴝ 202 Nonhematologic Overall Hypotension Asthenia Other

19 (9.5) 5 (2.5) 8 (4.0) 13 (6.4)

Hematologic Anemia

4 (2.0)

Leukocytopenia

6 (3.0)

Neutropenia Thrombocytopenia

19 (9.4) 8 (4.0)

Randomized phase, n ⴝ 151 Nonhematologic Arm A, no further treatment

2 (3)

Arm B, prolonged treatment

7 (10)

Hematologic Arm A, no further treatment

13 (17)

Arm B, prolonged treatment

13 (18)

Figure 4. Evolution of B lymphocytes according to study arm. The recovery of B cells starts 2 to 3 months after treatment and continues gradually with no further treatment, while the level of B cells remains suppressed with prolonged treatment.

of the IgM levels at 86% and 72% of baseline level at 3 and 12 months, respectively (P ⬍ .05).

Discussion This trial shows that prolonged treatment with rituximab, compared with the standard rituximab schedule, improves the outcome of patients with follicular lymphoma in terms of both EFS and response duration, without causing additional toxicity. The standard rituximab schedule achieves a response rate of 50% lasting approximately 1 year, while the prolonged schedule nearly doubled the median EFS and remission duration attained with rituximab reaching the order of magnitude seen with chemotherapy or chemoimmunotherapy.7,18 It is noteworthy that patients in both arms continued to convert from partial to complete response even many months after the end of treatment, resulting in similar partial and complete response rates in both arms. Other researchers have investigated the use of prolonged schedules or increased doses of rituximab including increasing the single dose or the number of infusions per week, increasing the number of consecutive weekly infusions up to 8, or repeating the weekly ⫻ 4 schedule every 6 months up to 2 years.19-22 Results from these trials suggest that increasing exposure to rituximab can improve efficacy. The trials with higher doses or more frequent administration were performed in chronic lymphocytic leukemia, demonstrating a RR that was higher than expected and suggesting that in this disease a dose-response or dose intensity–response correlation is likely. The weekly ⫻ 8 dosage was investigated in previously untreated low-grade lymphomas and showed an increased CR compared with the standard schedule, although this was not a randomized population.21 In that trial patients were highly selected with favorable characteristics and were not randomized against the standard schedule. Still, the weekly ⫻ 8 schedule did not seem to increase the overall RR or response duration compared to other series of previously untreated patients. Hainsworth and colleagues22 investigated an extended dosing approach by administering the standard schedule every 6 months in previously untreated patients. The trial showed an increase of the remission rate over time that suggested, as in the present trial, that rituximab-sensitive follicular lymphomas remain sensitive and can be maintained in remission by continuous exposure to the drug. The clinical effect of this schedule is similar to that in our experimental arm (EFS in chemotherapy-naive patients of 34 and 36 months, respectively), although the amount of drug used in the 2 trials was significantly different (20 infusions vs 8 infusions). Whether a continuous rituximab challenge at low dose, as in the present trial,



GHIELMINI et al

is more cost-effective than an intensive rechallenge every 6 months needs to be further investigated. The fact that 8 doses of rituximab given over 8 weeks are less effective on EFS than 8 doses given over 9 months suggests that the increased biological effect of our schedule is due more to the time of exposure to the drug than to peak serum levels. Another strategy that postpones the need for chemotherapy is to re-treat responders with the standard rituximab schedule at the time of their relapse. In a study by Davis and coworkers,23 it was observed that 40% of responders to the first exposure responded again at rechallenge, and the duration of the second response was longer than the first. The efficacy and cost-effectiveness of this strategy compared with those of a prolonged treatment schedule also need to be investigated in further studies. Preliminary data from a randomized trial addressing this question suggest that the time to chemotherapy is similar if patients receive rituximab for a prolonged time or at relapse only.24 Rituximab monotherapy is well tolerated. If compared with the toxicity of other chemotherapy schemes commonly used in follicular lymphoma, such as fludarabine or fludarabine combinations, CVP (cyclophosphamide, vincristine, and prednisone), or CHOP (cyclophosphamide, doxorubicin, vincristine, and prednisone), rituximab treatment, even when prolonged, appears to cause fewer adverse events.25-28 The major benefit from the experimental arm of this trial was achieved in previously untreated patients. Although the subgroup of treatment-naive patients with follicular lymphoma who receive prolonged therapy is small (n ⫽ 30) and retrospectively defined, and the trial was not powered to study the effect of prolonged treatment in untreated patients, it is interesting to note that in this group the median EFS and response duration are doubled with the prolonged schedule. The results are comparable to the most aggressive regimens used for follicular lymphoma such as ProMACE-MOPP (prednisone , high-dose methotrexate, adriamycin, cyclophosphamide, and VP-16–mechlorethamine-vincristineprocarbazine-prednisone), ChVP (cyclophosphamide, low-dose doxorubicin, teniposide, and prednisone), CVP, or fludarabine.18,26-28 If it is confirmed that first-line therapy with 8 infusions of rituximab can achieve a 36-month remission accompanied by little toxicity, it is likely that many patients would prefer this approach compared with the adverse effects of aggressive polychemotherapy, especially considering that a good proportion of responding patients can respond again at relapse.23 In order to optimize the cost-benefit ratio for prolonged treatment with rituximab we tried to identify those patients who are more likely to benefit from such treatment. As expected, the parameters reflecting tumor burden or number of previous treatments are important for both response duration and EFS. Using quantitative PCR as a surrogate marker for tumor burden, we were able to show for the first time that low numbers of circulating lymphoma cells (ie, ⬍ 374 t(14;18)–positive cells/106 blood cells or ⬍ 42 t(14;18)–positive cells/106 BM cells at baseline) are highly predictive for clinical response to rituximab. The prognostic value

of other parameters reflecting the immune status of the patient or biologic characteristics of the tumor is also an important new observation, although the interpretation of these findings is not straightforward. Lymphomas with bcl-2 immunoreactivity in greater than 80% of the tumor cells had a higher probability of responding. The prognostic impact of overexpression of the antiapoptotic bcl-2 protein is unclear, although, in the setting of rituximab treatment, it has been observed that in diffuse large B-cell lymphoma the overexpression of bcl-2 renders tumor cells more sensitive to chemotherapy.29,30 The results of this trial are encouraging and highly significant and support the potential for the prolonged schedule to become a treatment option for patients who have demonstrated sensitivity to rituximab. Continuing treatment for 9 months was an empirical decision, and further extension of regular administration of the drug (ie, until relapse) might substantially ameliorate remission duration. Considering the persistent depletion of B cells and the progressive reduction of serum IgM levels in the experimental arm, we would at present discourage such an approach outside a clinical trial setting. The follow-up trial to be conducted by our group aims to confirm the results of the present trial and explore the possibility of further prolongation of the treatment by comparing the 9-month schedule with a schedule of 2-monthly rituximab therapy administered until the time of disease progression.

Acknowledgments The authors would like to thank Patricia Katz and Corinne Friedly for central data managing, Dina Delle Pezze for the FACS analysis of lymphocyte subsets, and Dr Doris Schmitter and Ursula Bolliger for assistance in molecular biology analysis.

Appendix SAKK is a member of the Swiss Institute for Applied Cancer Research (SIAK).

SAKK center

Chair

Aarau

Dr M. Wernli

Basel

Prof R. Herrmann

Bern

Prof M. Fey

Geneva

Dr A. Roth

Grisons

Dr F. Egli

Lausanne

Prof S. Leyvraz

St Gallen

Prof T. Cerny

Ticino

Prof F. Cavalli

Zurich

Prof H.P. Honegger

SAKK is a member of the Swiss Institute for Applied Cancer Research (SIAK).

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Erratum In the article by James et al entitled “Anergic T cells exert antigenindependent inhibition of cell-cell interactions via chemokine metabolism,” which appeared in the September 15, 2003, issue of Blood (Volume 102:2173-2179), Figure 2B was incorrect. The corrected Figure 2 appears below: