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Blood, Vol. 95 No. 12 (June 15), 2000:
pp. 3900-3908
NEOPLASIA
Biologic response of B lymphoma cells to anti-CD20 monoclonal
antibody rituximab in vitro: CD55 and CD59 regulate complement-mediated
cell lysis
Josée Golay,
Luisella Zaffaroni,
Thomas Vaccari,
Manuela Lazzari,
Gian-Maria Borleri,
Sergio Bernasconi,
Francesco Tedesco,
Alessandro Rambaldi, and
Martino Introna
From the Istituto Ricerche Farmacologiche "Mario Negri,"
Milan; the Hematology Division, Ospedali Riuniti, Bergamo; and the
Department of Physiology and Pathology, University of Trieste, Trieste,
Italy.
 |
Abstract |
The chimeric anti-CD20 MAb rituximab has recently become a treatment
of choice for low-grade or follicular non-Hodgkin's lymphomas (FL)
with a response rate of about 50%. In this report, we have investigated the mechanism of action of rituximab on 4 FL and 1 Burkitt's lymphoma (BL) cell lines, 3 fresh FL samples and normal B
cells in vitro. Rituximab efficiently blocks the proliferation of
normal B cells, but not that of the lymphoma lines. We did not detect
significant apoptosis of the cell lines in response to rituximab alone.
All cell lines were targets of antibody-dependent cellular cytotoxicity
(ADCC). On the other hand, human complement-mediated lysis was highly
variable between cell lines, ranging from 100% lysis to complete
resistance. Investigation of the role of the complement inhibitors
CD35, CD46, CD55, and CD59 showed that CD55, and to a lesser extent
CD59, are important regulators of complement-mediated cytotoxicity
(CDC) in FL cell lines as well as in fresh cases of FL: Blocking CD55
and/or CD59 function with specific antibodies significantly increased
CDC in FL cells. We conclude that CDC and ADCC are major mechanisms of
action of rituximab on B-cell lymphomas and that a heterogeneous
susceptibility of different lymphoma cells to complement may be at
least in part responsible for the heterogeneity of the response of
different patients to rituximab in vivo. Furthermore, we suggest that
the relative levels of CD55 and CD59 may become useful markers to
predict the clinical response.
(Blood. 2000;95:3900-3908)
© 2000 by The American Society of Hematology.
| |
Introduction |
Rituximab is an anti-CD20 chimeric monoclonal antibody
containing the human IgG1 and  constant regions1 and is
the first MAb approved for the treatment of low-grade and follicular
non-Hogdkin's lymphoma (FL).2-7 In phase I studies,
rituximab induced a rapid depletion of CD20+ normal and
lymphoma cells.1-3 Phase II trials with low-grade or
follicular lymphomas showed a 50% response rate,5,8
whereas intermediate- to high-grade lymphomas showed a lower response rate.6 Little is known about the reason for the
heterogeneity of the response of different patients. Lack of response
does not seem to relate to CD20 levels because CD20 is highly expressed in most cases. A high tumor burden leading to high antibody clearance rate and low median serum antibody levels has been suggested as one
possible mechanism for lack of response.5,9 Other factors have not been investigated to date.
The biologic basis for choosing the CD20 antigen as a target was its
B-cell specificity, its expression at high levels during nearly all
stages of B-cell differentiation and the fact that the antigen is not
internalized, down-modulated, or shed by anti-CD20 antibodies.1,10 Its mechanism of action in vivo has not yet been fully clarified: It is likely to include complement-mediated cytotoxicity (CDC) and/or antibody-dependent cytotoxicity (ADCC), because an equivalent IgG4 version of the antibody lost the capacity to deplete normal B cells in vivo in nonhuman primates.11
However, different murine anti-CD20 monoclonal antibodies, in
particular the 1F5 and B1 antibodies that recognize close but distinct
epitopes on the CD20 molecule, have been shown to have other biologic
activities on B cells (reviewed by Tedder and Engel10). 1F5
activates normal B cells from the G0 to the G1 phase of the cell cycle,
but blocks their differentiation to immunoglobulin
secretion.12,13 Furthermore, 1F5 can deliver activation or
proliferation signals to some leukemic cells.13,14 The B1
antibody on the other hand blocks mitogen-induced proliferation and
does not activate B cells.15,16 B1 inhibits B-cell
proliferation induced by Staphylococcus aureus cowan I strain bacteria (SAC) or Epstein Barr virus (EBV) but has
little or no activity on anti-µ or T-cell derived signals, or on
phorbol myristic acetate (PMA)-induced
proliferation.15-17 B1 has been suggested to
act in late G1, blocking G1-S phase transition.10,17 In
addition, 1F5 or rituximab may have either a proapoptotic or antiapoptotic activity on neoplastic or normal B cells,
respectively.8,18-20
CD20 is a 33 to 37 kd phosphoprotein that forms tetramers and can act
as a Ca++ channel21 (reviewed by Tedder and
Engel10). It is phosphorylated in both normal and
neoplastic B cells.22,23 It is unclear whether Ca++ fluxes or the tyrosine kinases associated with CD20
are involved in signaling B-cell activation or inhibition of
proliferation.10,24 B-cell activation by 1F5 is accompanied
by induction of c-myc and B-myb.25,26
In this report, we have investigated the different biologic activities
of rituximab in vitro against normal B cells as well as several cell
lines and fresh lymphoma samples, including possible effects on
proliferation, apoptosis, CDC, and ADCC. This work was aimed at
defining the likely mechanism of action of rituximab on lymphoma cells
and the molecular basis for the heterogeneity of the response of
different patients to this agent.
| |
Materials and methods |
Cell cultures
The BJAB Burkitt's lymphoma (BL) line was a kind gift of Dr D. Vercelli (DIBIT, San Raffaele, Milano, Italy). The DHL-4 cell line was
a kind gift of Dr L. Boxer (Stanford University School of Medicine,
Stanford, CA). The Karpas 422, DOHH-2 and WSU-NHL follicular lymphoma
cell lines were purchased from the German Collection of Microorganisms
and Cell Culture (DSM, Braunschweig, Germany). Human resting B cells
were purified from freshly excised tonsils using
aminoethylisothiouronium-treated sheep red blood cells and
discontinuous Percoll gradients (Pharmacia, Uppsala, Sweden) as described.27 Mononuclear cells
from follicular lymphoma patients were purified by
Ficoll-Hypaque gradient centrifugation. All FL cell lines and fresh
samples were verified to carry the t(14;18) translocation by polymerase
chain reaction (PCR). The cells were cultured in RPMI 1640 medium
(Seromed, Berlin, Germany) supplemented with 10% fetal
calf serum (FCS) (Hyclone, Steril System, Logan, UT), glutamine (GIBCO,
Paisley, Scotland), and 50 µg/mL gentamicin (GIBCO).
Antibodies and immunofluorescence
The purified 1F5 anti-CD20 antibody and G28.5 anti-CD40 were a kind
gift of Dr E. A. Clark (University of Washington, Seattle, WA).12 Goat anti-µ F(ab')2 was from
Cappel (Organon Teknika Corp, West Chester, PA). The purified anti-CD19
and CD22 antibodies HD37 and HD239, respectively, were a kind gift of
Dr Moldenhauer (Deutsches Krebsforschungszentrum, Heidelberg,
Germany).28 Rituximab was supplied by Roche Italia (Monza,
Italy). The blocking anti-CD55 antibody BRIC216 was from IGBRL Research
Products, Bristol, UK. The blocking anti-CD46 antibody J4-48 was from
Cymbus Biotechnology Ltd (Chandlers Ford, Hants, UK). CD59
was detected with a goat anti-CD59 antiserum. This complement inhibitor
was isolated from human urine by affinity chromatography as
described,29 using the rat IgG2b MAb YTH53.1 kindly
provided by Prof H. Waldmann (Department of Pathology, University of
Cambridge, UK.).30 The IgG fraction of the anti-CD59
antiserum was purified by chromatography on a protein G-Sepharose
column (Pharmacia) and F(ab')2 fragments prepared by
digestion with pepsin (Sigma Chemical, St Louis, MO) for 4 hours at
37°C at an enzyme ratio of 1/600 (w/w). The
F(ab')2 fragment was purified by fast protein liquid
chromatography on a MonoQ column (Pharmacia) and the purity of the
fragments was checked by SDS-PAGE analysis. The purified
F(ab')2 fragment was used at a 1/100 dilution for
blocking experiments. For phenotype analysis, indirect
immunofluorescence was performed with the above antibodies and
fluorescein isothiocyanate (FITC)-labeled goat antimouse Ig (Becton
Dickinson, Mountain View, CA) or rabbit antigoat Ig (Sigma). Antibodies
against retinoblastoma, cyclin A, cdk2, and cdk4 were from Santa Cruz
(Santa Cruz, CA).
Proliferation assays
Resting tonsillar B cells were cultured at
1 × 106 cells/mL in 96-well plates. Proliferation
in quadruplicate wells was assessed at different times with a 4- to
16-hour pulse of 0.0185 MBq (0.5 µCi)
3H-thymidine per well. Formalin-killed SAC were from
Calbiochem Behring (La Jolla, CA). Monoparametric analysis on ethanol
fixed cells using propidium iodide was performed as described
previously using a FACStar Plus (Becton Dickinson).27
Antibody-dependent cellular cytotoxicity assays
2 × 106 lymphoma cells were loaded with
51Cr by incubation for 1 hour at 37°C with 3.7 MBq
(100 µCi) 51Cr and then washed 3 times in
phosphate-buffered saline (PBS). Total mononuclear cells were obtained
by Ficoll-Hypaque centrifugation of buffy coats. An enrichment to
15%-30% natural killer (NK) cells was obtained by separation on a
47% Percoll gradient. 104 lymphoma cells were incubated in
duplicate with or without 2 µg/mL rituximab and increasing amounts of
freshly prepared enriched NK cell population for 4 hours at 37°C in
complete medium. The 100 µL of cell supernatant was then collected
and counted in a  -counter.
Complement-mediated cell lysis
Cells at 106/mL were incubated with 2 µg/mL rituximab
and/or blocking antibodies for 10 minutes at room temperature. Five
percent to 50% normal human serum (NHS) was then added, mixed by
pipetting, and incubation was carried out for an additional 1 hour at
37°C. The extent of lysis was measured by direct cell count in a
Bürker chamber of trypan blue-stained samples or by
fluorescence-activated cell sorter (FACS) analysis of acridine
orange-stained cells on a FACStar Plus (Becton Dickinson). Briefly,
lysed cells were mixed with an equal volume of 3 µg/mL acridine
orange solution in PBS and analyzed immediately. Statistical analysis
was performed with the Student t test.
Northern blots
Total RNA was extracted with guanidinium isothiocyanate and purified
on cesium chloride gradients according to standard protocols. Northern
blotting was carried out as described.26 The c-myc, B-myb, cdk2, and cyclin A probes have been described
elsewhere.26,31
Western blots
Total proteins from 1.5 × 106 cells were
extracted in SDS loading buffer and boiled for 5 minutes. Samples were
run in SDS-PAGE gels and elec-troblotted onto nitrocellulose
(Schleicher and Schuell, Dassel, Germany) for 5 hours at 60 V,
according to standard procedures. The blots were incubated with the
relevant antibodies diluted in PBS containing 5% nonfat milk powder
and washed in the same solution containing 0.5% Nonidet P-40.
Secondary peroxidase conjugated antimouse or antirabbit antibodies
were used at 1/1000 (Amersham, Buckinghamshire,
UK). Detection was performed using the enhanced chemiluminescence system (ECL; Amersham).
Apoptosis assays
Apoptosis was measured with the Annexin-V-FLUOS kit (Roche
Diagnostics, Milan, Italy) according to the manufacturer's
instructions and analysis on the FACS.
| |
Results |
Rituximab blocks the proliferation of normal B cells induced by SAC,
not by anti-µ and anti-CD40
The first step was to determine whether rituximab can block
proliferation of B cells. For this purpose we purified resting human B
lymphocytes from tonsils and stimulated them with either SAC bacteria
or with anti-µ and anti-CD40 antibodies. As shown in Figure
1A, rituximab blocked SAC-induced B-cell
proliferation by about 90%, even at the lowest concentration (0.3 µg/mL). An irrelevant IgG1 antibody (My7) had no effect. On the
contrary, anti-µ + anti-CD40 triggered proliferation was little
affected by rituximab, with a maximal inhibition of 15%. These
findings show that rituximab, like B1, is a blocking anti-CD20 antibody and that it interferes with some but not all intracellular
signals.15,17
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| Fig 1.
Inhibition of normal B-cell proliferation by rituximab.
(A) Resting tonsillar B cells were stimulated in vitro with SAC (open
circles) or anti-µ and anti-CD40 (closed circles) in the presence of
the indicated concentrations of rituximab. SAC-stimulated cells were
also incubated with a control IgG1 antibody (10 µg/mL My7 IgG1) (open
square). 3H-thymidine incorporation of quadruplicate wells
was measured at 48 to 66 hours. The data are represented as percentage
of the stimulated controls in the absence of rituximab. (B) Resting
tonsillar B cells were stimulated with SAC and rituximab was added at 2 µg/mL at the indicated times. 3H-thymidine was measured
at 48 to 66 hours.
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To determine at which stage of the cell cycle rituximab acts, the
antibody was added at different times after stimulation of resting B
cells with SAC. As shown in Figure 1B, rituximab was fully effective
when added after up to 12 hours of culture, but its activity decreases
thereafter, suggesting a target of action acting during mid-late G1
phase of the cell cycle. Subsequent cell cycle analysis by flow
cytometry showed that B cells were blocked mostly in G1 because
rituximab led to a 50% decrease in the percentage of cells in the S,
G2, and M phases of the cell cycle (Figure
2).
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| Fig 2.
Rituximab blocks B-cell proliferation before entry into S
phase.
Resting tonsillar B cells were stimulated with SAC in the presence or
absence of 2 µg/mL rituximab. Cell cycle analysis was performed at
the indicated times. The percentage of cells in the different phases of
the cell cycle are indicated.
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We also analyzed expression of several cell cycle proteins induced by
SAC. In agreement with a block in mid-late G1, we found that rituximab
inhibited SAC-induced Rb hyperphosphorylation (Figure 3A). Furthermore, induction of cdk2, cdk4,
and cyclin A protein expression, which is associated with G1-S phase
transition, was also blocked. That equivalent amounts of proteins were
loaded in each lane was verified by Ponceau red staining of the blot (data not shown). To determine whether cyclin or cdk2 expression was
inhibited at the transcriptional level, expression of these molecules
was analyzed at the RNA level in Northern blots. As shown in Figure 3B,
rituximab did not significantly block the induction of cyclin A or cdk2
messenger RNA (mRNA) expression. These data show that the antibody
affects cyclin A and cdk2 induction only at the protein level.
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| Fig 3.
Rituximab blocks hyperphosphorylation of Rb and induction
of cell cycle proteins.
Resting tonsillar B cells were stimulated with SAC in the presence or
absence of 2 µg/mL rituximab. (A) Total protein extracts were
prepared from 1.5 × 106 cells for each time point
and analyzed in Western blots with the indicated antibodies. (B) Total
RNA was extracted at the indicated times. RNA (20 µg/mL) was analyzed
by Northern blot analysis using the indicated probes. The lower panel
shows the photograph of the ethidium bromide stained blot. The data are
representative of 2 independent experiments.
|
|
We next wanted to determine whether rituximab has only an inhibitory
effect on B-cell proliferation, like B1, or whether it also has
activating function like 1F5. To measure B-cell activation, we analyzed
the expression of 2 cell cycle regulated genes, c-myc and
B-myb, which are associated with cell activation and have been
previously shown to be induced by the 1F5 antibody.25,26 As
shown in Figure 4, rituximab did not induce
c-myc or B-myb mRNA expression at 3 or 48 hours,
respectively. 1F5, on the other hand, was fully effective as expected
(Figure 4).
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| Fig 4.
Rituximab does not activate B cells.
Resting tonsillar B cells were stimulated with 2 µg/mL rituximab or
1F5 anti-CD20 antibodies. Total RNA was extracted at 3 hours (c-myc,
lanes 1-3) or 48 hours (B-myb, lanes 4-6) and analyzed for expression
of the indicated mRNA by Northern blot. The photograph of the ethidium
bromide stained blots is shown below to demonstrate that all lanes
contained equivalent quantities of RNA.
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We conclude that rituximab, like B1, can efficiently block B-cell
proliferation induced by some (SAC) but not all (anti-µ and
anti-CD40) mitogenic signals,15,17 that it arrests B cells mostly in the G1 phase of the cell cycle, at least in part, through inhibition of retinoblastoma phosphorylation, and that, unlike 1F5, it
has no activating function.
Effect of rituximab on proliferation and/or apoptosis of lymphoma
cell lines
We next investigated whether rituximab also had an antiproliferative
activity on 4 different follicular lymphoma lines (DOHH2, DHL-4,
WSU-NHL, Karpas 422). Proliferation was measured by
3H-thymidine uptake assays 24, 48, and 72 hours after the
beginning of culture. We did not detect any significant effect of
rituximab on proliferation of these cell lines (data not shown). These
data also indicated that rituximab did not induce significant cell death in these lines, which would have been detected as a decrease in
thymidine uptake. However, 2 previous reports had suggested a
proapoptotic effect of rituximab on DHL-4 cells, either alone or in
combination with chemotherapeutic agents.19,20 We therefore went on to verify more extensively this point. DHL-4 cells were cultured in the presence or absence of 2 to 300 µg/mL rituximab for
up to 4 days and live and dead cells counted at regular intervals. As
shown in Figure 5, the growth curve and
number of live cells were not affected by rituximab for up to 48 hours.
Only a small (8%-15%) but reproducible increase in the number of dead
cells could be detected at 48 hours. The number of dead cells did not increase further at 96 hours (data not shown). We also directly measured the number of apoptotic and necrotic cells by Annexin V
binding and propidium iodide staining. As shown in Table
1, rituximab did not induce an increase in
apoptotic or necrotic cells up to 96 hours after the beginning of
culture. On the other hand, staurosporine, a known inducer of
apoptosis, did induce a clear increase in early apoptotic cells at 4 hours. As expected, these cells became necrotic (Annexin V and
propidium iodide positive) at later time points (Table 1). We also
investigated the activation of caspase 3 and 7 using Ac-DEVD-MCA, a
fluorescent substrate for these enzymes.32 We could detect
a 2-fold caspase activation by staurosporine at 4 hours but not by
rituximab at 4, 24, or 48 hours (data not shown). Similarly, we could
detect apoptotic DNA fragmentation 6 hours after staurosporine addition
but not 6 or 12 hours after rituximab (data not shown).
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| Fig 5.
Rituximab alone does not induce growth inhibition or cell
death of DHL-4 cells.
DHL-4 cells were plated at 4 × 105 cells/mL in the
presence or absence of 10 µg/mL rituximab. Live and dead cells were
counted at the indicated times by trypan blue exclusion. The results
are representative of 3 independent experiments.
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We conclude that rituximab alone does not inhibit proliferation or
induce significant apoptosis in any of the cell lines examined.
Rituximab can induce ADCC to similar levels in different lymphoma
lines
One proposed mechanism of action of rituximab in vivo is the
induction of ADCC.11 We were therefore interested to
determine whether the lymphoma lines could be lysed by ADCC triggered
by rituximab and whether this response was variable between different cell lines. For this study, 3 FL cell lines and 1 Burkitt's lymphoma line (BJAB) were chosen. ADCC was measured in 3 independent experiments using as effector population, fresh peripheral blood mononuclear cell
fractions enriched in CD16+ NK cells to 15% to 30%. All
cell lines were always tested in parallel to exclude differences
between the effector populations. As shown in Figure
6, 2 of the cell lines were targets for
natural killer activity in the absence of antibody. In particular, the WSU-NHL cell line showed a maximum of 20% killing in the absence of
rituximab. All cell lines showed a 10% to 20% increase in lysis in
the presence of rituximab. We conclude that rituximab can activate the
ADCC of several FL and BL lines and that levels of ADCC do not vary
significantly between the different cell lines.
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| Fig 6.
Rituximab can trigger ADCC of lymphoma lines.
The 51Cr-labeled cell lines were cultured at
5 × 104/mL in the presence of the indicated amounts
of effector cells enriched for NK cells and in the presence (closed
circles) or the absence (open circles) of 2 µg/mL rituximab. The
cells were incubated for 4 hours at 37°C. 51Cr released
in the supernatant was measured as percentage of total 51Cr
released with 1% SDS. The results are the mean and SD of 3 independent
experiments.
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The efficiency of lysis of different FL lines by complement is
heterogeneous
CDC is known to take place in vivo and is likely to be an important
mechanism for the elimination of both normal and lymphoma B cells. We
have therefore tested the capacity of the different lymphoma lines to
be killed by CDC in vitro. As shown in Figure 7, increasing concentrations of human serum
were added as a source of complement in the presence or absence of
rituximab. Serum alone had little toxic effect, except at the higher
concentration (50%), where about 10% to 15% lysis was usually
observed (Figure 7, open squares). In the presence of rituximab, on the
other hand, the different cell lines were lysed in a very heterogeneous
manner: At the extremes, the DHL4 line was completely lysed with only 10% serum, whereas the Karpas 422 showed complete resistance at 25%
and insignificant rituximab-dependent lysis at 50%. The BJAB and
WSU-NHL cell lines showed an intermediate phenotype with an average of
25% and 50% lysis for BJAB and WSU-NHL, respectively, with 25% serum
(Figure 7 and data not shown). Because 50% serum was slightly toxic on
its own, a concentration of 25% was used in subsequent experiments as
a standard efficacious concentration. We also tested the
rituximab-dependent lysis of purified tonsillar B cells. These also
showed clear heterogeneity because 80% were lysed but 20% were
resistant. Similar CDC results were obtained in several experiments and
by evaluating lysis either by a direct count of trypan blue negative
and positive cells or by a FACS analysis of acridine orange-stained
cells (which stains live cells) (see below) or propidium
iodide-stained cells (which marks lysed cells). The results were found
to be highly reproducible using several different batches of NHS (data
not shown).
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| Fig 7.
Complement-mediated lysis of lymphoma lines by rituximab.
The indicated cell lines were incubated at
1 × 106/mL in the presence (closed squares) or the
absence (open squares) of 2 µg/mL rituximab and increasing
concentrations of NHS. Percentage cell lysis was measured after 1 hour
at 37°C by cell count with trypan blue. The data are representative
of at least 3 different experiments.
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The degree of resistance of the lines to complement mediated lysis
correlates with the levels of CD55
To determine the underlying mechanism accounting for the differences
in susceptibility to complement of the lymphoma cell lines, we first
investigated the levels of expression of CD20 itself on the cell
surface. As shown in Figure 8, the 4 cell
lines all expressed CD20 at high levels on more than 80% of the cells. Although a portion of Karpas 422 cells expressed CD20 at slightly lower
levels than the other cell lines, this small difference could not
justify a 100% difference in cell lysis between Karpas 422 and DHL-4.
Furthermore, the BJAB cell line expressed very high levels of CD20 on
virtually 100% of the cells but is lysed only to about 25%. We
concluded that the susceptibility of the different lines to complement
is not due to antigen density but to other cellular factors. Several
complement inhibitors are known to be present on the surface of
hematopoietic cells.33 The best characterized are CD35,
CD46, CD55, and CD59. The expression pattern of these 4 molecules on
the surface of the lymphoma cell lines was therefore investigated. CD35
was not detectable at significant levels on any of the cell lines (data
not shown). On the contrary, CD46 was present in similar amounts on all
4 lines (Figure 8). The last 2 inhibitors, CD55 and CD59, showed a more
interesting pattern in that they were variably expressed in the
different cell lines. CD55 was expressed at the highest levels in the
Karpas 422 cell line and at the lowest levels in the DHL-4 cell line, with the percentage of positive cells ranging from 95.6 to 20.3, respectively. The WSU-NHL and BJAB cell lines showed intermediate levels of expression (57.7 and 55.0, respectively, Figure 8). These
differences were reproducible (in at least 4 separate experiments). Thus, CD55 levels correlated quite well with the resistance to complement-mediated lysis. CD59 levels were also variable among the
different cell lines. Karpas 422 was essentially negative, whereas the
other cell lines showed intermediate levels of expression (from 13% to
60%) (Figure 8). CD59 levels therefore did not correlate well with the
degree of resistance to complement because the most resistant cell
line, Karpas 422, did not express CD59, whereas the least resistant
(DHL-4) was positive (30%).
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| Fig 8.
Phenotype of the lymphoma cell lines.
Exponentially growing cells were stained by indirect immunofluorescence
for the indicated surface markers. · · · negative control.
__ positive sample. The percentages of positive cells are
indicated in each plot.
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CD55 and CD59 can inhibit complement-mediated cell lysis
To demonstrate directly that CD55 present on the cell lines studied
can indeed inhibit complement, we have carried out a complement lysis
experiment in the presence or absence of antibodies that recognize and
functionally block either to CD55 or to CD46.33 Both
antibodies are murine IgG1 antibodies that cannot therefore activate
complement by themselves. As a control an anti-CD19 IgG1 antibody
(clone HD37) was used. As shown in Figure
9, the anti-CD55, -CD46, or -CD19
antibodies had no effect on complement lysis in the presence of serum
alone (ie, they did not activate complement by themselves).
Interestingly, the anti-CD55, but not anti-CD46 or -CD19 antibodies,
increased the killing of all cell lines in the presence of both
rituximab and complement. This increase was statistically significant
for all cell lines examined (Figure 9). Of note is that the experiment
with the DHL-4 cell line was performed with only 5% serum to obtain
suboptimal levels of lysis in the absence of anti-CD55. An augmented
lysis was reproducibly observed in 5 separate experiments for each cell
line, with a mean increase of 280% for Karpas, 140% for BJAB, 40%
for WSU-NHL, and 20% for DHL-4. Thus, the cells that were most
resistant to complement, Karpas 422 and BJAB, also showed the greatest
response to blocking anti-CD55 antibody. We conclude that CD55 can
regulate complement-mediated lysis and is a most effective inhibitor on the resistant Karpas 422 cell line.
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| Fig 9.
CD55, but not CD46, inhibits complement-mediated lysis.
The indicated cells were incubated in presence (+) or absence ( )
of rituximab and 25% NHS and 10 µg/mL anti-CD55, -CD46, or -CD19
(all IgG1). Cell death was measured after 1 hour at 37°C by cell
count. The results are representative of at least 5 separate
experiments. The statistical significance is shown by asterisks. *
P < .05; ** P < .01; ***
P < .001 (Student t test). ND, not determined.
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Because CD59 has been reported to inhibit CDC in different cell
types,34-37 we have also investigated the functional effect of a blocking anti-CD59 antibody (F(ab')2 antiserum).
The blocking anti-CD59 had no effect of cell viability in the presence
of serum but the absence of rituximab (data not shown), indicating that it was not able to activate complement by itself. The same antibody, however, induced a 6% to 20% increase in complement-mediated lysis triggered by rituximab in all cell lines, except Karpas 422, which does
not express CD59 (Table 2). As shown
previously (Figure 9), the blocking anti-CD46 antibody had no effect.
Interestingly, anti-CD55 synergized with anti-CD59 on the WSU-NHL and
BJAB cell lines, which were lysed completely in the presence of both
blocking antibodies. No such synergy could be observed in DHL-4 or
Karpas 422 (Table 2). The data are representative of at least 3 independent experiments.
We conclude that both CD55 and CD59 are functional on several of the
cell lines examined and that higher expression of CD55 can explain at
least in part the resistance to complement of the Karpas 422 cell line.
CD55 and CD59 also affect lysis of fresh FL cells
We next collected blood samples from 3 cases of t(14;18) FL
patients. The phenotype of the purified peripheral blood mononuclear cells (PBMCs) from the patients is shown in Table
3. Patients 1 and 3 contained 96% to 98%
CD20-positive neoplastic B cells and 1% to 3% T lymphocytes. Patient
2 contained only about 50% CD20-positive neoplastic B cells and 37% T
lymphocytes. The cells from all patients expressed the complement
inhibitors CD46, CD55, and CD59 with somewhat variable intensity. We
next measured complement-mediated lysis of the same mononuclear cell
preparations. As shown in Table 4, patient
1 showed about 60% lysis in the presence of both serum and rituximab.
Killing is increased to about 90% in the presence of blocking
anti-CD55 and to 83% in the presence of anti-CD59. Virtually complete
lysis (93%) was obtained with both anti-CD55 and anti-CD59. These
effects were statistically significant. Anti-CD46 had no effect.
Patients 2 and 3 showed a different pattern of lysis: CD20-positive
cells were essentially completely lysed with rituximab and complement
alone. Indeed, blocking the function of CD55 or CD59 or of both
together only marginally increased CDC of these populations, confirming
that most CD20-positive cells were killed by rituximab and complement
alone. To verify the specificity of CDC on blood mononuclear cells, CDC
was also performed on PBMCs from a normal donor. Rituximab and
complement lysed only 5% to 10% of normal PBMCs even in the presence
of blocking CD55 and CD59. This percentage corresponds to the number of
CD20-positive cells in the sample (data not shown).
We conclude that the different FL patients studied showed a different
susceptibility to lysis with rituximab and complement. Interestingly,
lysis of the most resistant FL cells could be increased significantly
with blocking anti-CD55 and/or CD59 antibodies, demonstrating a role
for these inhibitors also on fresh FL cells.
| |
Discussion |
In this work, we have investigated the mechanism of action of
rituximab on 5 human lymphoma cell lines, 4 of the follicular type, and
1 Burkitt, on normal B lymphocytes as well as on 3 fresh cases of
follicular lymphoma. We have examined the potential effect of rituximab
on B-cell proliferation, activation, apoptosis, antibody-dependent cell
mediated cytotoxicity (ADCC) and complement-mediated cytotoxicity (CDC). These studies suggest that complement and complement inhibitors are likely to play a role in the heterogeneity of the response of
different FL patients to rituximab in vivo.
We show here that rituximab inhibits efficiently SAC but not anti-µ
and anti-CD40-induced proliferation of normal B cells. Unlike 1F5,
rituximab did not activate B cells to express c-myc or B-myb. This is
the first demonstration that rituximab is a blocking antibody.
Similarly to B1, it shows specificity for some mitogenic signals and
not others.15,17 The mechanism for this specificity is not
clear at present because the intracellular signals with which B1 or
rituximab interfere have not yet been defined. These may include
Ca++ or kinase-mediated events or both.10,21,24
We have shown here with time course assays and cell cycle analysis that
rituximab blocks cells in mid-late G1 but before entry into S phase. In agreement with these data, we have found that hyperphosphorylation of
retinoblastoma protein, as well as cdk2, cdk4, and cyclin A induction,
is inhibited by rituximab. Interestingly, inhibition of cdk2 and cyclin
A takes place at the protein but not the RNA level, suggesting that
signals involved in the regulation of cdk2 and cyclin A protein
stability or expression are blocked by CD20. The molecular mechanism by
which rituximab inhibits cdk and cyclin A induction and Rb
phosphorylation is beyond the scope of this article but is likely to be
a major cause of the proliferation block induced by the
antibody.38
In contrast to normal B cells, rituximab did not block the
proliferation of 4 FL cell lines. This finding suggests that the signals induced by CD20 either are not active in the cell lines or are
not sufficient to block proliferation. This effect could be related to
the lack of inhibition of growth factor-induced proliferation of
normal B cell by B1 antibody.16 The lack of inhibition of
the cell lines could be a specific property of long-term culture
because the proliferation and/or differentiation of some fresh leukemic
B cells have been shown as being blocked by B1 antibody.13
Thus, the blocking activity of rituximab could also be a relevant
property for neoplastic B cells in vivo.
1F5 and rituximab has been reported previously to induce apoptosis of
Ramos or DHL4 cells, respectively.18-20 The data presented here do not support the conclusion that rituximab on its own has a
significant proapoptotic activity for FL cells, including DHL-4. We
detected a less than 15% increase in cell death after 48 to 96 hours
incubation. Furthermore, Annexin V binding, an early marker of
apoptosis, or caspase 3/7 activity, were not up-regulated 4 to 96 hours
after the addition of rituximab in vitro. The reasons for the
discrepancies with the previous report are not clear but may be due to
culture conditions.20 Altogether our data demonstrate that
rituximab on its own is not proapoptotic, at least in vitro. We cannot
exclude, however, that rituximab has proapoptotic activity against FL
cells in vivo or that, as suggested by others,19 it can
cooperate with other proapoptotic signals, such as those delivered by
chemotherapeutic drugs.
We have next investigated whether the lymphoma lines were targets for
either ADCC or CDC. The results presented show that, whereas the 4 cell
lines studied are all targets of ADCC and show marginal differences in
the extent of lysis, the sensitivity to complement varies dramatically
between the different targets. These differences were highly
reproducible, were not dependent on the batches of serum, and were
therefore a property of the cell lines. The enormous difference of
susceptibility to complement between monoclonal leukemic cell lines is
of particular interest because such differences may be at least in part
the basis for differences in the responses of different patients to
rituximab in vivo. The data presented here on only 3 fresh cases of FL
clearly do not allow us to draw firm conclusions on this point but do support such a hypothesis: Also, the fresh lymphoma cells showed heterogeneity with regard to complement lysis. In this context, it is
worth recording that ADCC and CDC are likely to be responsible for the
in vivo normal B-cell depletion because the equivalent IgG4 anti-CD20
antibody, which lacks both functions, does not deplete B cells in
monkeys.11
Study of the 4 major cell surface complement inhibitors, namely, CD35,
CD46, CD55, and CD59, pointed to a role for CD55 in determining
resistance to CDC: (i) Levels of expression of CD20 were roughly
equivalent in all lines. (ii) CD55 was the only inhibitor whose
expression correlated with resistance to complement. (iii) Blocking of
CD55 function led to a net increase in CDC, particularly in the
complement-resistant cell lines Karpas 422 and BJAB. Indeed, Karpas 422 could be lysed to 50% in the presence of blocking anti-CD55. (iv)
Anti-CD55 antibody was also effective on fresh FL cells: Patient 1 showed only partial lysis (55%-60%) in the absence of anti-CD55 and
was completely lysed (90%-95%) in its presence. Anti-CD46 had no such
effect. CD55 has been shown previously to play a role in inhibiting CDC
of BL-cell lines by the alternative pathway,34 of HIV-1
infected lymphocytes or different tumor cells by the classical
pathway.33,35,39,40 Our data, together with the report that
patients lacking CD55 do not show clinical signs of hemolytic
anemia,18,41 suggest that the inhibition of CD55 during
rituximab treatment could be clinically feasible to increase the
response of resistant patients to the drug.
CD59 blocks the last steps of complement activation and has been shown
to inhibit CDC of some BL-cell lines, erythrocytes,34,40,42 T cells,35 renal carcinoma cell lines,37 and
melanoma lines.36 In agreement with previous reports,
blocking anti-CD59 antibodies increased CDC in all lymphoma cell lines
that express detectable levels of the protein. In addition, in 2 cell
lines (BJAB and WSU-NHL), anti-CD59 antibodies were synergistic with
anti-CD55. Indeed, near complete lysis of both cell lines could be
obtained in the presence of the 2 antibodies. Blocking CD59 also
increased lysis of fresh FL cells, again suggesting the relevance of
this antigen in vivo. A study of a larger group of patients will be required to determine the role of the different surface antigens and is
beyond the scope of this article. However, we cannot exclude that
factors other than CD55 and CD59 contribute to the differences in
complement susceptibility.
To conclude, we suggest that study of the expression of both CD55 and
CD59 on the surface of neoplastic B cells, in addition to that of CD20,
will be important to predict their possible response to rituximab in
vivo. This knowledge may also be important to predict
infusion-related side effects in patients with circulating tumor
cells. In addition, novel and still more efficacious
immunotherapeutic strategies could include either the combined
administration of rituximab, together with blocking anti-CD55/59
antibodies, or the production of bispecific anti-CD20/CD55 or
CD20/CD59 reagents.39
| |
Footnotes |
Submitted June 9, 1999; accepted February 2, 2000.
Supported in part by grants from Roche Italia, the Associazione
Italiana Ricerca sul Cancro (AIRC), The Istituto Superiore di
Sanità (ISS, Rome, AIDS project 30a.0.29), the Consiglio
Nazionale della Ricerche (CNR): target project on biotechnology (no.
97.01278.PF49 and 99.00496.PF4), the Associazione Paolo Belli, Lotta
alla Leucemia, and by Biomed 2 concerted action (no. BMH4-CT96-1005).
Reprints: Martino Introna, Laboratory of Molecular
Immunohematology, Department of Immunology and Cell Biology, Istituto Ricerche Farmacologiche Mario Negri, via Eritrea 62, 20157 Milano, Italy; e-mail: martino{at}irfmn.mnegri.it.
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.
| |
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Top
Abstract
Introduction