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NEOPLASIA
From the Hematopathology Unit, Department of
Hematology, Institute of Hematology and Oncology, Department of
Pathology, Postgraduate School of Hematology Farreras-Valentí;
the Institut d'Investigacions Biomèdiques August Pi i Sunyer
(IDIBAPS), Hospital Clínic; and the University of Barcelona,
Barcelona, Spain.
Mechanisms involving the in vitro effect of rituximab in cells from
55 patients with B-cell lymphoproliferative disorders were
investigated. No cytotoxic effect was observed when cells were
incubated with rituximab alone, but in the presence of human AB serum
rituximab induced complement-dependent cell death (R-CDC). A cytotoxic
effect was observed in cells from 9 of 33 patients with B-cell chronic
lymphocytic leukemia, 16 of 16 patients with mantle-cell lymphoma, 4 of
4 patients with follicular lymphoma, and 2 of 2 patients with
hairy-cell leukemia. R-CDC was observed in cells from patients
expressing more than 50 × 103 CD20 molecules per cell,
and directly correlated with the number of CD20 molecules per cell.
Preincubation with anti-CD59 increased the cytotoxic effect of
rituximab and sensitized cells from nonsensitive cases. Neither
cleavage of poly-ADP ribose polymerase (PARP) nor activation of
caspase-3 was observed in R-CDC. In addition, no cells with a
hypodiploid DNA content were detected and R-CDC was not prevented by a
broad-spectrum caspase inhibitor, suggesting a caspase-independent
mechanism. Incubation with rituximab in the presence of AB serum
induced a rapid and intense production of reactive oxygen species
(ROS). R-CDC was blocked by the incubation of cells with
N-acetyl-L-cysteine (NAC) or Tiron, 2 ROS scavengers, indicating that
the cytotoxic effect was due to the generation of superoxide
(O Rituximab is a chimeric monoclonal antibody
directed against CD20, an antigen present both on normal B lymphocytes
and on cells from most B-cell lymphoproliferative
disorders.1 Rituximab is currently employed in the
treatment of follicular lymphoma (FL) either
alone2,3 or in combination with
chemotherapy.4 Moreover, there is an increasing interest
in using rituximab in other CD20+ B-cell
lymphoproliferative disorders, such as mantle-cell lymphoma (MCL) or
B-cell chronic lymphocytic leukemia (B-CLL).5,6 A direct
relationship between the clinical efficacy and the intensity of CD20
expression has been proposed, suggesting that patients with high CD20
expression are more likely to respond to rituximab.
The signaling pathways involved in the effect of rituximab are not
clearly established. Recently, complement-mediated cell lysis has
been proposed as the major and most-efficient effector mechanism of
rituximab.1,7 However, the signal transduction pathways
activated by complement, which precede cell death, have not been fully
determined. Moreover, in vitro studies performed in cell lines suggest
that rituximab binding to CD20 could induce apoptosis, mainly through
caspase activation,8 but other pathways, such as
activation of the Src-family kinases9 and
antibody-dependent cell-mediated cytotoxicity (ADCC),1,7
have also been described.
Complement-mediated cell lysis involves a cascade activation of
proteins leading to the formation of the membrane attack complex, which
produces a direct lysis of the target cells. This lysis is regulated by
membrane-bound regulatory proteins, among which CD55 and CD59 seem to
be the most important. CD55 binds to C3b and C4b and accelerates the
decay of C3 and C5 convertases, whereas CD59 binds to C8 and C9 and
prevents pore formation by the membrane attack complex,10
the final step of complement lysis. According to recent studies, these
proteins could be implicated in the cytotoxic effect of
rituximab.7,11
The aims of this study were to correlate the cytotoxic effect of
rituximab with CD20 expression in a variety of CD20+ B-cell
lymphoid malignancies, and to analyze the signaling pathways involved
in complement-dependent cell death (CDC), focusing on complement
regulatory proteins.
Patients
Reagents
Isolation of cells Mononuclear cells were isolated from peripheral blood samples by centrifugation on a Ficoll/Hypaque (Seromed, Berlin, Germany) gradient and cryopreserved in liquid nitrogen in the presence of 10% dimethyl sulfoxide (DMSO). Cells from 5 cases (1 B-CLL, 3 MCL, and 1 FL) were obtained from lymph node biopsy or spleen after repetitive infiltration with RPMI 1640 culture medium (GibcoBRL, Paisley, Scotland).Cell culture Lymphocytes were cultured immediately after thawing at a concentration of 5 × 106 cells/mL in RPMI 1640 culture medium supplemented with 10% heat inactivated fetal calf serum (BioWhittaker, Verviers, Belgium), 2 mM glutamine, and 0.04 mg/mL gentamicin at 37°C in a humidified atmosphere containing 5% carbon dioxide. Factors were added at the beginning of the culture.Analysis of cell viability by annexin binding Exposure of phosphatidylserine residues was quantified by surface annexin V staining as previously described.13 Briefly, cells were washed in binding buffer (10 mM HEPES, pH 7.4, 2.5 mM CaCl2, 140 mM NaCl), resuspended in 200 µL and incubated with 0.5 µg/mL annexin V-fluorescein isothiocyanate (FITC) (Bender Medsystems, Vienna, Austria) for 15 minutes in the dark. Cells were washed again and resuspended in binding buffer. A quantity of 5 µg/mL propidium iodide (PI) (Sigma Chemical) was added to each sample prior to flow cytometric analysis (FACScan; Becton Dickinson). Ten thousand cells were acquired per sample using CELLquest software and data were analyzed with the Paint-a-gate Pro software (Becton Dickinson). All experiments were performed in duplicate. In some experiments, cells were labeled simultaneously with annexin V-FITC and anti-CD3-PE, in order to separately analyze the cytotoxic effect on B and T lymphocytes.Propidium iodide DNA staining Quantification of apoptosis by PI staining and fluorescence-activated cell sorting (FACS) analysis was performed as previously described.14 Briefly, cells were harvested and fixed in 70% ethanol. Cells were centrifuged, washed in phosphate-buffered saline (PBS) and resuspended in 0.5 mL PBS containing PI (5 µg/mL) and RNase (100µg/mL) (Boehringer Mannheim, Mannheim, Germany). Tubes were incubated for 30 minutes at 37°C and placed at 4°C in the dark overnight prior to flow cytometry analysis.Assessment of mitochondrial transmembrane potential and reactive oxygen species production Changes in mitochondrial transmembrane potential (  m) were evaluated by staining with 1 nM
3,3'-dihexyloxacarbocyanine iodide (DiOC6[3]; Molecular
Probes, Eugene, OR). Reactive oxygen species (ROS) production was
determined by staining with 2 µM dihydroethidine (DHE; Molecular
Probes). Cells were incubated with the dyes for 15 minutes at 37°C,
washed, resuspended in PBS and analyzed by flow cytometry. A decrease
in the signal of DiOC6[3] (FL1) was indicative of
abnormalities in m and appearance of FL2 signal was
indicative of ROS production. Ten thousand cells were acquired in a
FACScan flow cytometer. All experiments were performed in duplicate.
Kinetic studies of ROS generation Cells were preincubated for 5 minutes with 50 µg/mL rituximab, and cell acquisition in a FACScan was started after incubation with DHE for one minute. AB serum was added after the first minute of cell acquisition and ROS production was recorded for 15 minutes. Kinetics of ROS generation were analyzed using the free software WinMDI 2.8 version (http://archive.uwcm.ac.uk/uwcm/hg/hoy/software.html).Flow cytometric detection of the active form of caspase-3 Cells were fixed and permeabilized using the Cytofix/Cytoperm kit (Pharmingen, San Diego, CA) for 20 minutes at 4°C, pelleted and washed with Perm/Wash buffer (Pharmingen). Cells were then stained with the polyclonal antibody against the active form of caspase-3 (Pharmingen) (0.25 µg/L × 106 cells) for 20 minutes at room temperature, washed in Perm/Wash buffer, stained with goat anti-rabbit-FITC (SuperTechs, Bethesda, MD), and analyzed in a FACScan.Quantification of CD20, CD55, and CD59 membrane proteins Cells from patients or healthy donors (n = 19) were incubated with saturating amounts of monoclonal antibodies against CD20 (Leu16), CD55 (143-30), and CD59 (MEM 43) for 45 minutes at 4°C, washed twice in PBS and incubated with goat anti-mouse-FITC (Dako, Globstrub, Denmark) in the same conditions. The beads of QIFKIT (Dako) were processed in parallel following the manufacturer's recommendations. The mean FL1 channel for every bead population was used to calculate a standard curve and the number of molecules per cell was obtained by interpolation of the mean FL1 channel value for each patient. To quantify the number of complement regulatory proteins on B cells, samples stained with CD55 and CD59 were subsequently incubated with normal mouse serum (Dako) for 5 minutes, followed by CD19-PE monoclonal antibody.Western blot Cells were lysed in 80 mM Tris HCl pH 6.8, 2% sodium dodecyl sulfate (SDS), 10% glycerol, 0.1 M dithiothreitol (DTT) and equal amounts of protein were separated by electrophoresis on 12% polyacrylamide gel and transferred to Immobilon-P (Millipore, Bedford, MA) membranes. The membranes were incubated with polyclonal antibody against poly-ADP ribose polymerase (PARP) (Boehringer Mannheim). Antibody binding was detected using a secondary antibody (mouse anti-rabbit immunoglobulin; Dako) conjugated to horseradish peroxidase and an enhanced chemiluminiscence (ECL) detection kit (Amersham, Buckinghamshire, United Kingdom).Statistical analysis A case was considered sensitive to rituximab when cell death assessed by exposure of phosphatidylserine residues was more than or equal to 15% of cell death observed in control cells. Comparison of CD20, CD55, and CD59 cell levels between healthy donors and patients, and between sensitive and nonsensitive cases were performed using the nonparametric Mann-Whitney test. The effect of blockage with anti-CD55 and/or anti-CD59 was analyzed by means of linear regression models with random effects using a panel data structure, adjusted by normalized values of CD55 and CD59 molecules per cell, with STATA Statistical Software, Release 6.0 (Stata, College Station, TX).
Rituximab induces cell death of tumor B cells in the presence of complement To analyze the in vitro effect of rituximab on the viability of tumor cells from patients with B-cell lymphoproliferative disorders, a dose-response study was first performed in cells from 3 patients. No cytotoxic effect was observed when cells were incubated with rituximab alone (with doses ranging from 10 to 100 µg/mL) for increasing periods of time (up to 5 days). However, when normal human AB serum (AB) was added as a source of complement, rituximab produced a cytotoxic effect after 24 hours of incubation. This effect was dose dependent for both rituximab and AB (with quantities ranging from 1% to 40%) and could be detected using doses as low as 10 µg/mL rituximab and 1% AB. A discriminant cytotoxic effect was observed with 50 µg/mL rituximab in the presence of 10% AB. Thus, this was the combination used in all subsequent experiments.The cytotoxic effect of rituximab in the presence of AB (rituximab+AB)
was characterized by a reduction in the forward scatter (FSC) and an
increase in the side scatter (SSC), indicative of cell shrinkage
(Figure 1A), as well as an exposure of
phosphatidylserine residues, as shown by the increase in annexin
V-labeled cells (Figure 1B), and loss of
Inactivation of the complement by heating AB at 56°C for 30 minutes completely abolished the cytotoxic effect of rituximab. Incubation of cells with AB alone did not induce cell death but, to the contrary, produced a protective effect when compared with cells incubated with medium alone (data not shown). The in vitro effect of rituximab+AB was analyzed on cells from 55 patients. In the above-mentioned conditions, R-CDC was observed in cells from 31 of the 55 patients (56%). The distribution of R-CDC among the different lymphoproliferative disorders was as follows: 9 of 33 in B-CLL (27%), 16 of 16 in MCL (100%), 4 of 4 in FL (100%), and 2 of 2 in HCL (100%). R-CDC was not observed on CD3+ cells from 20 cases as assessed by multiparametric flow cytometry analysis (data not shown), with this indicating that the cytotoxic effect of rituximab was restricted to B cells. R-CDC depends on CD20 expression In order to analyze the differences between cases that are sensitive (cytotoxicity 15%) and nonsensitive (cytotoxicity
< 15%) to rituximab+AB, we quantified the expression of the CD20 antigen. As shown in Figure 2A, a
significant difference (P < .0001) was found between the
CD20 mean values of sensitive cases (100.7 ± 41 × 103
molecules per cell; n = 31) and nonsensitive cases
(18.7 ± 12 × 103 molecules per cell; n = 24). All
the cases expressing more than or equal to 50 × 103
molecules of the CD20 antigen on the cell membrane surface
(CD20++) (n = 29) showed R-CDC, whereas no cytotoxic
effect was observed in cases with CD20 levels less than
40 × 103 molecules per cell (CD20dim)
(n = 22). Interestingly, a direct correlation was found between R-CDC
and the number of CD20 molecules per cell (r = 0.8,
P < .0001) (Figure 2B).
No modifications were observed in the FSC/SSC pattern or in annexin V binding when cells from 4 CD20dim cases were incubated with rituximab+AB for longer periods of time (up to 5 days) (data not shown). As shown in Table 1, the number of CD20
molecules was significantly lower in B-CLL cells than in normal B
cells, whereas no difference was observed in the other types of
B-lymphoproliferative disorders analyzed. Of note, cells from
nonsensitive B-CLL cases had significantly lower CD20 levels than cells
from sensitive B-CLL cases.
Regulation of R-CDC by CD55 and CD59 To study the role of the complement in R-CDC, we quantified the expression of CD55 and CD59, 2 complement regulatory proteins. The number of CD55 molecules per cell was significantly lower in B-CLL and HCL cells than in normal B cells, whereas no difference was observed in other B-lymphoproliferative disorders (Table 1). Regarding the number of CD59 molecules, only the subgroup of sensitive B-CLL cells expressed lower CD59 molecules per cell than normal B cells. No correlation was found between R-CDC and the number of CD55 or CD59 molecules on the cell surface. Among 4 B-CLL samples that expressed between 40 × 103 molecules per cell and 50 × 103 CD20 molecules per cell (CD20+), only 2 of them were sensitive to rituximab. Interestingly, these 2 cases expressed lower CD59 levels than the 2 nonsensitive cases (Figure 3A).
To better understand the role of CD55 and CD59, the effect of
rituximab+AB was analyzed in the presence of anti-CD59 and/or anti-CD55
in cells from 9 B-CLL patients (Figure 3B). No cytotoxic effect was
observed when cells were incubated with these monoclonal antibodies
alone, or combined with AB, indicating that they cannot activate the
complement by themselves. The statistical analysis of the mean values
obtained for these 9 patients showed that the viability of cells
incubated with rituximab+AB was 80 ± 7% (95% confidence interval
[CI]: 67-93). The addition of anti-CD59 produced a
significant decrease in cell viability by 48 units (95% CI: 38-57;
P < .001), whereas addition of anti-CD55 only
produced a decrease of 8 units (95% CI: 0-17;
P = .078). The combination of the 2 monoclonal
antibodies produced an additional decrease in cell viability of 7.4 units (95% CI: Thus, in B-cell lymphoproliferative disorders, the cytotoxic effect of rituximab is mediated, at least in part, by the complement and is regulated by the number of CD20 and CD59 molecules per cell. Induction of R-CDC in CD20++ cells is not mediated by caspase activation and does not induce DNA cleavage We investigated the implication of caspases in R-CDC in 4 CD20++ cases (3 B-CLL and 1 MCL). Incubation of cells for 24 hours with rituximab+AB did not activate caspase-3, as determined by the absence of the active form of this protease by flow cytometry (Figure 4A). Furthermore, R-CDC was not associated with caspase-mediated nuclear features of apoptosis such as PARP proteolysis or DNA cleavage (appearance of the hypodiploid DNA peak) after 48 hours of incubation (Figure 4B,C). Finally, addition of 200 µM Z-VAD.fmk, a broad-spectrum caspase inhibitor, did not prevent phosphatydilserine exposure, cell shrinkage, and loss ofm induced by rituximab+AB (Figure 4D). To discard the
possibility that caspase activation occurred at earlier time points, we
analyzed caspase-3 activation in 3 B-CLL sensitive cases following
treatment with rituximab+AB for 1, 2, and 4 hours. No evidence of
caspase activation was observed, whereas cells at these time points
showed the typical features of cell death (data not shown). All these
results indicate that the mechanism involved in R-CDC may occur in the
absence of caspase activation.
R-CDC induces production of ROS Since the cytotoxic effect of rituximab+AB was accompanied by a loss inm, we tested whether this phenomenon was
associated with the production of ROS. As seen in Figure
5A, incubation for 24 hours of
CD20++ cells from a representative MCL patient with
rituximab+AB induced the production of ROS, as assessed by staining
with DHE, a compound converted by O
To determine the implication of ROS in R-CDC, we incubated cells with
rituximab+AB in the presence of 2 ROS scavengers. As shown in Figure
5B, incubation of cells from a representative CD20++
patient with Tiron, a pharmacologic cell-permeable scavenger of
O Finally, in order to establish the sequence of cellular changes induced
by rituximab+AB, the kinetics of R-CDC were investigated in 4 CD20++ cases (1 B-CLL, 1 FL, 2 MCL). Addition of AB in
cells preincubated with rituximab resulted in a rapid and intense
detection of ROS, with maximal levels being achieved in less than 15 minutes (Figure 6A). As seen in Figure
6B, ROS production was accompanied by loss in
The results presented in this study show that the cytotoxic effect
of rituximab on primary malignant B cells is, at least in part,
complement mediated and that this process mainly occurs through the
production of ROS. The generation of cytotoxic ROS in cells is due to
the partial reduction of oxygen that leads to the generation of
O In the present study, we found a rapid and selective generation of ROS
after the addition of rituximab and AB serum, since only specific
scavengers of O Consistent with previous reports,11 rituximab alone was unable to induce a cytotoxic effect on primary malignant B cells, the addition of a source of complement being necessary to obtain such an effect. Another mechanism that has been involved in the cytotoxic effect of rituximab is ADCC.1,7 However, a previous report comparing specific rituximab-induced ADCC and CDC in several cell lines showed that CDC is more effective than ADCC.7 Although CDC could be the major mechanism accounting for the elimination of circulating B lymphocytes following rituximab infusion, ADCC may contribute to the elimination of B cells in tissues where interaction between target and effector cells is higher. Incubation of cells with rituximab and AB serum induced the typical
cytoplasmic features of apoptosis: flow cytometric changes in FSC and
SSC indicative of cell shrinkage, exposure of phosphatidylserine residues, and decrease in It has been described that rituximab induces a caspase-dependent apoptosis.8,9,33 However, in all these studies in vitro crosslinking of rituximab with secondary antibodies was performed. The production of antibodies against rituximab has not been described in patients treated with rituximab; therefore, this mechanism is unlikely to occur in vivo.2 Nevertheless, preliminary studies have suggested that caspases may be activated following in vivo infusion of rituximab,34 therefore additional mechanisms could be involved in in vivo caspase activation. Although it has been described that some anti-CD20 antibodies can promote the activation of the Src-family kinases,9 no implication of these kinases has been observed in our study. In fact, incubation with a selective inhibitor of this family of kinases (PP2) did not prevent R-CDC in cells from 4 of our cases (data not shown). Altogether, it is likely that many different pathways of cell death both dependent and independent of apoptosis could mediate the elimination of tumoral B cells in vivo. We have observed that R-CDC directly correlates with the expression of the CD20 antigen in malignant B cells and a certain amount of antigen is required to trigger R-CDC. Therefore, the clinical differences in responses among patients with different types of B-lymphoproliferative disorders could be related, at least in part, to CD20 expression. Another factor influencing the response to rituximab is complement regulatory proteins. The blockage of CD59 with specific antibodies sensitized cells to rituximab, indicating that CD59, the protein controlling the membrane-attack complex, may have an important role in R-CDC. Similar results have been obtained in previous studies,7 in contrast to others that proposed a major role for CD55.11 These differences could be related to the different clones of monoclonal antibodies used to block cellular antigens. The present results, obtained in primary malignant B cells, can serve as the basis for new therapeutic approaches that may improve the efficacy of rituximab in lymphoid malignancies. In this regard, it has been suggested that the up-regulation of CD20 on malignant B cells could enhance the effect of rituximab.35-37 Additional strategies could involve the control of complement regulatory proteins on the surface of neoplastic B cells either by using bispecific monoclonal antibodies against both CD20 and CD59, to avoid side effects of a wide CD59 blockage, or by down-regulating the expression of CD59 with pharmacologic agents.38,39 In conclusion, this report provides evidence that CD20, CD59, and complement contribute to the in vitro cytotoxic effect of rituximab, with this being mediated by a caspase-independent process that involves ROS generation and loss of mitochondrial transmembrane potential. These results may be useful to establish a theoretical basis to improve the efficacy of therapy with rituximab.
We thank L. Quintó from the Unidad de Epidemiología y Bioestadística, Hospital Clínic, for his help in statistical analysis.
Submitted January 18, 2001; accepted May 14, 2001.
Supported in part by research fellowships from the Instituto de Salud Carlos III (B.B.) and Fondo de Investigaciones Sanitarias (FIS) (S.M.); FIS grant numbers 98/0996, 99/0189, and 00/0946; José Carreras International Foundation Against Leukemia (EM/P-01 and CR/P-00); Roche España; and the Asociación Española Contra el Cáncer, Barcelona, Spain.
Roche Spain provides funds for research activities of the Institute of Hematology and Oncology.
B.B and N.V. contributed equally to this study. D.C. and E.M. share the senior authorship of this paper.
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.
Reprints: Emili Montserrat, Institute of Hematology and Oncology, Department of Hematology, Hospital Clínic, Villarroel 170, 08036 Barcelona, Spain; e-mail: emontse{at}clinic.ub.es.
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© 2001 by The American Society of Hematology.
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A. De Milito, E. Iessi, M. Logozzi, F. Lozupone, M. Spada, M. L. Marino, C. Federici, M. Perdicchio, P. Matarrese, L. Lugini, et al. Proton Pump Inhibitors Induce Apoptosis of Human B-Cell Tumors through a Caspase-Independent Mechanism Involving Reactive Oxygen Species Cancer Res., June 1, 2007; 67(11): 5408 - 5417. [Abstract] [Full Text] [PDF]
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Y. Ge, I. Montano, G. Rustici, W. J. Freebern, C. M. Haggerty, W. Cui, D. Ponciano-Jackson, G. V. R. Chandramouli, E. R. Gardner, W. D. Figg, et al. Selective leukemic-cell killing by a novel functional class of thalidomide analogs Blood, December 15, 2006; 108(13): 4126 - 4135. [Abstract] [Full Text] [PDF]
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P. Perez-Galan, G. Roue, N. Villamor, E. Montserrat, E. Campo, and D. Colomer The proteasome inhibitor bortezomib induces apoptosis in mantle-cell lymphoma through generation of ROS and Noxa activation independent of p53 status Blood, January 1, 2006; 107(1): 257 - 264. [Abstract] [Full Text] [PDF]
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R. Niwa, M. Sakurada, Y. Kobayashi, A. Uehara, K. Matsushima, R. Ueda, K. Nakamura, and K. Shitara Enhanced Natural Killer Cell Binding and Activation by Low-Fucose IgG1 Antibody Results in Potent Antibody-Dependent Cellular Cytotoxicity Induction at Lower Antigen Density Clin. Cancer Res., March 15, 2005; 11(6): 2327 - 2336. [Abstract] [Full Text] [PDF]
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M. L. Palomba, W. K. Roberts, T. Dao, G. Manukian, J. A. Guevara-Patino, J. D. Wolchok, D. A. Scheinberg, and A. N. Houghton CD8+ T-Cell-Dependent Immunity Following Xenogeneic DNA Immunization against CD20 in a Tumor Challenge Model of B-Cell Lymphoma Clin. Cancer Res., January 1, 2005; 11(1): 370 - 379. [Abstract] [Full Text] [PDF]
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T. S. Lin, I. W. Flinn, R. Modali, T. A. Lehman, J. Webb, S. Waymer, M. E. Moran, M. S. Lucas, S. S. Farag, and J. C. Byrd FCGR3A and FCGR2A polymorphisms may not correlate with response to alemtuzumab in chronic lymphocytic leukemia Blood, January 1, 2005; 105(1): 289 - 291. [Abstract] [Full Text] [PDF]
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C.-L. Law, C. G. Cerveny, K. A. Gordon, K. Klussman, B. J. Mixan, D. F. Chace, D. L. Meyer, S. O. Doronina, C. B. Siegall, J. A. Francisco, et al. Efficient Elimination of B-Lineage Lymphomas by Anti-CD20-Auristatin Conjugates Clin. Cancer Res., December 1, 2004; 10(23): 7842 - 7851. [Abstract] [Full Text] [PDF]
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G. Cartron, H. Watier, J. Golay, and P. Solal-Celigny From the bench to the bedside: ways to improve rituximab efficacy Blood, November 1, 2004; 104(9): 2635 - 2642. [Abstract] [Full Text] [PDF]
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T. Decker, M. Oelsner, R. J. Kreitman, G. Salvatore, Q.-c. Wang, I. Pastan, C. Peschel, and T. Licht Induction of caspase-dependent programmed cell death in B-cell chronic lymphocytic leukemia by anti-CD22 immunotoxins Blood, April 1, 2004; 103(7): 2718 - 2726. [Abstract] [Full Text] [PDF]
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A. P. Mone, P. Huang, H. Pelicano, C. M. Cheney, J. M. Green, J. Y. Tso, A. J. Johnson, S. Jefferson, T. S. Lin, and J. C. Byrd Hu1D10 induces apoptosis concurrent with activation of the AKT survival pathway in human chronic lymphocytic leukemia cells Blood, March 1, 2004; 103(5): 1846 - 1854. [Abstract] [Full Text] [PDF]
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S. S. Farag, I. W. Flinn, R. Modali, T. A. Lehman, D. Young, and J. C. Byrd Fc{gamma}RIIIa and Fc{gamma}RIIa polymorphisms do not predict response to rituximab in B-cell chronic lymphocytic leukemia Blood, February 15, 2004; 103(4): 1472 - 1474. [Abstract] [Full Text] [PDF]
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N. Di Gaetano, E. Cittera, R. Nota, A. Vecchi, V. Grieco, E. Scanziani, M. Botto, M. Introna, and J. Golay Complement Activation Determines the Therapeutic Activity of Rituximab In Vivo J. Immunol., August 1, 2003; 171(3): 1581 - 1587. [Abstract] [Full Text] [PDF]
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O. Manches, G. Lui, L. Chaperot, R. Gressin, J.-P. Molens, M.-C. Jacob, J.-J. Sotto, D. Leroux, J.-C. Bensa, and J. Plumas In vitro mechanisms of action of rituximab on primary non-Hodgkin lymphomas Blood, February 1, 2003; 101(3): 949 - 954. [Abstract] [Full Text] [PDF]
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M. S. Cragg, S. M. Morgan, H. T. C. Chan, B. P. Morgan, A. V. Filatov, P. W. M. Johnson, R. R. French, and M. J. Glennie Complement-mediated lysis by anti-CD20 mAb correlates with segregation into lipid rafts Blood, February 1, 2003; 101(3): 1045 - 1052. [Abstract] [Full Text] [PDF]
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A. D. Kennedy, M. D. Solga, T. A. Schuman, A. W. Chi, M. A. Lindorfer, W. M. Sutherland, P. L. Foley, and R. P. Taylor An anti-C3b(i) mAb enhances complement activation, C3b(i) deposition, and killing of CD20+ cells by rituximab Blood, February 1, 2003; 101(3): 1071 - 1079. [Abstract] [Full Text] [PDF]
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J. C. Byrd, B. L. Peterson, V. A. Morrison, K. Park, R. Jacobson, E. Hoke, J. W. Vardiman, K. Rai, C. A. Schiffer, and R. A. Larson Randomized phase 2 study of fludarabine with concurrent versus sequential treatment with rituximab in symptomatic, untreated patients with B-cell chronic lymphocytic leukemia: results from Cancer and Leukemia Group B 9712 (CALGB 9712) Blood, January 1, 2003; 101(1): 6 - 14. [Abstract] [Full Text] [PDF]
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A. L. Rose, B. E. Smith, and D. G. Maloney Glucocorticoids and rituximab in vitro: synergistic direct antiproliferative and apoptotic effects Blood, August 13, 2002; 100(5): 1765 - 1773. [Abstract] [Full Text] [PDF]
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B. Bellosillo, N. Villamor, A. Lopez-Guillermo, S. Marce, F. Bosch, E. Campo, E. Montserrat, and D. Colomer Spontaneous and drug-induced apoptosis is mediated by conformational changes of Bax and Bak in B-cell chronic lymphocytic leukemia Blood, August 13, 2002; 100(5): 1810 - 1816. [Abstract] [Full Text] [PDF]
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Top
Abstract
Introduction
) and sensitive
(
) patients.
5.7-20.5), this not being significantly different
from the decrease observed with CD59 alone (P = .269).
Finally, the cytotoxic effect produced when blocking with anti-CD59
directly correlated with the number of CD59 molecules per cell
(P < .001).
m,
which was detected as early as 2 minutes after the addition of AB.
Exposure of phosphatidylserine residues was detected simultaneously to
these changes in cells from CD20++ patients with high
expression of CD20 molecules (Figure 
) and hydrogen peroxide
(H2O2),
.
RI/CD64-directed bispecific antibodies in B-cell lymphoma.
Blood.
2000;96:3544-3552