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Blood First Edition Paper, prepublished online April 23, 2013; DOI 10.1182/blood-2013-02-482570
IMMUNOBIOLOGY
Rituximab causes a polarisation of B cells which augments its therapeutic
function in NK cell-mediated antibody-dependent cellular cytotoxicity
Dominika Rudnicka1, Anna Oszmiana2, Donna K. Finch3, Ian Strickland3, Darren J.
Schofield3, David C. Lowe3, Matthew A. Sleeman3, Daniel M. Davis1,2*
1
Division of Cell and Molecular Biology, Imperial College London, Sir Alexander
Fleming Building, London, SW7 2AZ, UK
2
Manchester Collaborative Centre for Inflammation Research, Core Technology
Facility, University of Manchester, Oxford Road, Manchester, M13 9PT, UK
3
MedImmune Ltd, Milstein Building, Granta Park, Cambridge, CB21 6GH, UK
*Address correspondence to: Daniel M. Davis, Division of Cell and Molecular
Biology, Sir Alexander Fleming Building, Imperial College London, SW7 2AZ, UK.
Tel: +44-207-594-5420; Fax: +44-207-594-3044; E-mail: [email protected]
Running title: CD20 reorganisation augments ADCC
1
Copyright © 2013 American Society of Hematology
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Key points
•
Rituximab causes a polarisation of B cells, involving a reorganisation of CD20,
ICAM-1 and moesin, and orientation of the MTOC.
•
The polarisation of B cells induced by rituximab augments its therapeutic role
in triggering ADCC by effector NK cells.
2
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Abstract
Rituximab, which binds CD20 on B cells, is one of the best characterised antibodies
used in the treatment of B cell malignancies and autoimmune diseases. Rituximab
triggers Natural Killer (NK) cell-mediated antibody-dependent cellular cytotoxicity
(ADCC) but little is known about the spatial and temporal dynamics of cell-cell
interactions during ADCC - nor what makes rituximab potent at triggering ADCC.
Here, using laser scanning confocal microscopy, we found that rituximab caused
CD20 to cap at the B cell surface, independent of antibody cross-linking or
intercellular contact. Unexpectedly, other proteins, including ICAM-1 and moesin,
were selectively recruited to the cap of CD20 and the MTOC became polarised
towards the cap. Importantly, the frequency at which NK cells would kill target cells
via ADCC increased by 60% when target cells were polarised compared to being unpolarised. Polarised B cells were lysed more frequently still, when initial contact with
NK cells occurred at the place where CD20 was capped. This demonstrates that the
site of contact between immune cells and target cells influences immune responses.
Together, these data establish that rituximab causes a polarisation of B cells and this
augments its therapeutic function in triggering NK cell-mediated ADCC.
3
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Introduction
Depletion of malignant or autoreactive B cells plays an important role in the
treatment of B-cell lymphomas and autoimmune diseases1-2. Rituximab is a
chimaeric human-mouse antibody that targets CD20, a pan-B cell surface marker,
and mediates depletion of these cells3. CD20 is highly expressed on the surface of B
cells as well as the majority of B-cell lymphomas4 but absent from the
haematopoietic stem cells, differentiated plasma cells and other healthy tissues
allowing for a specific targeting of desired cells. Furthermore, it is not shed or
internalised from the surface of the cells upon antibody binding5 making it a good
target for efficient induction of effector mechanisms that mediate depletion of B cells.
The exact mechanism of rituximab-mediated B cell depletion in patients is not fully
understood. Rituximab can potentially trigger three effector functions: programmed
cell death6; induce complement-dependent cytotoxicity; or activate immune cells,
including NK cells, which express Fc gamma receptor III (CD16) to mediate ADCC78
. The respective importance of these mechanisms may vary in different
environments. Evidence that ADCC is important for the activity of rituximab in vivo is
that mice deficient in activating Fc receptors responded poorly to antibody
treatment7. Similarly for humans, patients with high affinity CD16 polymorphism
responded better to rituximab treatment than those with low affinity receptor8. There
is also evidence that macrophages and neutrophils conjugate with antibodyopsonised targets forming ADCC synapses in vivo in mice9. In humans, NK cells are
considered to be the main mediators of ADCC and indeed, NK cells efficiently kill B
cells opsonised with rituximab in vitro and in vivo10-13.
However, few studies have used microscopy to visualise what happens during
ADCC14. Here we employed high-resolution microscopy to study the sequence of
events when NK cells attach to and then kill target cells opsonised by rituximab.
Unexpectedly, we found that rituximab induces polarisation of CD20, ICAM-1,
myosin and the MTOC; and such polarised cells are preferentially killed by effector
NK cells – especially in interactions where NK cells initially contact B cells where
CD20 has been capped. These data are important in establishing properties that a
therapeutic antibody should have to be optimal in triggering ADCC.
4
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Methods
Cells and constructs
Daudi, 721.221, Raji and Jurkat cells were maintained in RPMI-1640 supplemented
with 10% FCS, 50 U/mL penicillin, 50 µg/mL streptomycin and 2 mM L-glutamine (all
Invitrogen) (referred to later as ‘complete medium’). The plasmid encoding CD20GFP was a kind gift from J. Deans (Calgary, Canada). CD20-GFP was subcloned as
an AgeI/BamHI fragment into the retroviral pIB2 vector, a gift from M. Purbhoo,
Imperial College London.
Peripheral blood NK and B cells were isolated by negative selection from healthy
donor lymphocyte cones purchased from the National Blood Service or fresh blood
using magnetic beads (NK cell isolation kit, B cell isolation kit; Miltenyi Biotec). All
fresh blood donors were healthy and gave informed consent for their blood to be
used in accordance with the Declaration of Helsinki (with ethics approved by The
National Research Ethics Service, Ref 05/Q0401/108). Cells were maintained in
DMEM supplemented with 10% human serum (type AB; Sigma-Aldrich), 30%
nutrient mixture F-12, 2 mM L-glutamine, 1×nonessential amino acids, 1 mM sodium
pyruvate, 50 µM 2-ME, 50 U/mL penicillin and 50 µg/mL streptomycin (all Invitrogen).
Clinical grade rituximab (Rituxan, Roche) was used at 10 µg/mL for 1 hour unless
indicated otherwise. Where indicated, cells were pre-treated with 20 µg/mL CD32
blocking mAb (Clone IV.3, Stemcell technologies) or 10 µM nocodazole (Sigma) in
complete medium for 30 minutes at 37°C prior to incubation with rituximab. Drugs
were then maintained in the medium during incubation with rituximab.
Immunostaining and imaging
For co-localisation experiments, Daudi or primary human B cells were incubated with
10 µg/mL 2H7 or rituximab for 1 hour and fixed with 4% paraformaldehyde (PFA) in
PBS. For intracellular staining, cells were then permeabilised with 0.05%
Saponin/PBS (Sigma) and stained with mAb. For live cell imaging, 5×104 Daudi cells
were pre-incubated with 0.01 – 10 µg/mL rituximab for 1 hour. 1×105 primary NK
cells were pre-incubated with 1 µl/mL LysoTracker Red DND-99 (Molecular Probes)
for 1h. Cells were then mixed together and imaged in eight-well chambers
5
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(Chambered Borosilicate Coverglass, Nunc) pre-coated with 10 μg/mL
fibronectin/PBS (Sigma). Cells were imaged in the presence of 0.5 μM Sytox Blue
(Invitrogen) to visualise cell death. Brightfield and fluorescence images were
obtained by confocal microscopy (Leica SP5 RS) with a 63× water immersion lens
(NA 1.2) with live cell samples maintained at 37°C with 5% (vol/vol) CO2. Time-lapse
imaging was performed over 40-60 minutes with confocal stacks being acquired
every 30-40 s. For quantification of target cell killing, no killing was scored if target
and effector cells parted without target cell death or if the two cells stayed in contact
for at least 20 minutes until the end of the acquisition. Images were analysed
(Volocity, Improvision and ImageJ National Institutes of Health) and co-localisation
between two fluorescence channels assessed by calculating the Pearson’s
correlation coefficient (Image Correlation Analysis plug-in for ImageJ15). Brightness
and contrast were changed in some images for presentation of the figures shown but
analysis used raw images. For flow-based microscopy, Daudi/CD20-GFP cells were
incubated with rituximab for 1 hour and fixed with 4% PFA/PBS, imaged using a
multispectral imaging flow cytometer (ImageStream100, Amnis) and CD20 capping
analysed (IDEAS software, Amnis).
Statistical Analysis
Column Statistics were performed (GraphPad software, Prism) and unless specified
otherwise, mean values and SEM are shown. Data were analysed by one-way
ANOVA test with Bonferroni adjustment. To analyse the MTOC polarisation, a
Kolmogorov-Smirnov test was performed using an application available on-line at
http://www.physics.csbsju.edu/stats/KS-test.html.
6
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Results
Rituximab triggers capping of CD20 at the surface of B cells, independent of
cross-linking
To investigate the effect of opsonisation with rituximab on the organisation of the B
cell surface, three B cell lines – the Burkitt lymphoma B cell lines, Daudi and Raji,
and the EBV-transformed B cell line 721.221 – were transfected to express CD20
with an N-terminal GFP tag. Levels of CD20 expression in these transfectants were
somewhat higher but comparable to the levels of expression of CD20 in WT cells
(supplemental Figure 1). CD20-GFP was frequently distributed evenly (on a
micrometre scale) around the cell surface of all three cell lines prior to the addition of
rituximab (Figure 1A-C, left panels). Incubation with 10 µg/mL rituximab, induced
CD20 to frequently cap on one side of all three cell lines tested (Figure 1A-C, right
panels). Importantly, cross-linking of rituximab by a secondary antibody was not
required to observe this effect.
To test whether or not CD20 endogenously expressed in primary B cells also caps,
peripheral blood B cells were isolated and left untreated or incubated with 10 µg/mL
rituximab, fixed and stained with an antibody targeting the cytoplasmic portion of
CD20. Rituximab caused CD20 to commonly cap to one side of primary B cells
(Figure 1D). Without rituximab, the frequency at which CD20 was capped was
25.7±4.5% and 29.9±1.4% for primary B cells and Daudi/CD20-GFP cells,
respectively (Figure 1E). Upon treatment with 10 µg/mL rituximab, 61.2±4.7%
primary B cells and 68.5±1.6% Daudi transfectants had CD20 capped to one side of
the cell (Figure 1E). Additionally, quantification of CD20 polarisation in Daudi/CD20GFP cells was assessed by flow microscopy allowing unbiased automated
measurements of CD20 capping in a large number of cells (over 350 cells per
condition per experiment). Data obtained by flow microscopy were in agreement with
the results from single-cell quantification; capping of CD20 was observed in
29.6±1.9% untreated cells and increased to 70.6±0.8% in cells incubated with 10
µg/mL rituximab (Figure 1F).
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Quantification of the confocal microscope images taken throughout the volume of the
cell revealed that the majority of the CD20 was accumulated in the cap. 68±2% –
83±1% and 68±2% – 72±2% of total CD20 was accumulated in the cap within Daudi
and primary B cells, respectively (Figure 1G). The amount of protein accumulated in
the cap slightly (but significantly) increased in Daudi cells treated with rituximab but
not significantly in primary B cells. These differences may indicate quantitative
differences in the effects of rituximab on primary cells and immortal cell lines.
To investigate whether or not capping of CD20 was specific for rituximab or could be
triggered by any mAb against CD20 we compared treatment with 10 µg/mL 2H7, a
mAb which targets a similar epitope within CD20 as rituximab16. In contrast to
rituximab, incubation with 2H7 did not increase the number of cells with polarised
CD20 (Figure 2A). One possible explanation for antibodies to vary in their ability to
cause protein capping would be if they were internalised differentially. However, flow
cytometric analysis confirmed that neither rituximab nor 2H7 were internalised to a
significant extent over the time frame of these experiments (supplemental Figure
2A). Taken together, these data establish that a large fraction of cell surface CD20 is
capped to one side of a B cell upon treatment with rituximab specifically.
Fc receptors are not involved in rituximab-mediated capping of CD20
B cells express FcγRIIb (CD32) on their surface and therefore binding of the Fc
portion of rituximab to CD32 could potentially be involved in capping CD20. Indeed, it
has been previously suggested that rituximab can cross-link CD20 and CD3217. To
test this possibility, Daudi cells were pre-treated with a blocking mAb against CD32
for 1 hour prior to incubation with rituximab. Blocking CD32 did not influence the
frequency at which cells were capped by rituximab, indicating that the interaction
between the Fc portion of the antibody and CD32 does not play a role in mediating
CD20 polarisation (Figure 2A). To assess whether or not the bivalency of rituximab
was required to induce CD20 capping, cells were also incubated with a monovalent
version of rituximab IgG (described in supplemental Methods) that bound efficiently
to CD20 (supplemental Figure 2B). Monovalent rituximab IgG was not able to
increase the frequency at which CD20 was capped in B cells (Figure 2A). Taken
together, these data demonstrate that rituximab triggers the capping of CD20,
independent of cross-linking, but requiring the bivalency of the mAb.
8
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Rituximab does not efficiently cap CD20 in T cell transfectants
To investigate whether or not capping of CD20 was specific to B cells or could occur
in other cell types, Jurkat T cells were transfected to express CD20-GFP
(Jurkat/CD20-GFP). The level of expression of CD20 in Jurkat/CD20-GFP and
Daudi/CD20-GFP was comparable (supplemental Figure 3A). Surprisingly, CD20
was never polarised in untreated Jurkat/CD20-GFP and rituximab treatment induced
capping of CD20 in only 12.1±1.6% of cells (Figure 2B-C). In the cells where CD20
was capped, the amount of protein in the cap (58.1±2.5%) was significantly less than
in primary B cells (Figure 2D).The interaction of rituximab with CD20 was preserved
in Jurkat/CD20-GFP cells as they became susceptible to NK cell-mediated ADCC,
albeit to a relatively low extent (supplemental Figure 3B). This data indicates that
rituximab-mediated capping of CD20 is especially pronounced for B cells, likely
requiring cellular proteins that are absent from T cells. This adds further evidence
that rituximab does more than merely bind CD20 passively at the cell surface.
Redistribution to lipid rafts is not essential for rituximab-mediated CD20
capping
Rituximab has previously been shown to redistribute CD20 into lipid rafts18. To test
whether or not recruitment to lipid rafts caused capping of CD20, Daudi B cells were
transfected to express GFP attached to a mutant variant of CD20 (CΔ219-225)
which lacks a membrane-proximal sequence previously established to be important
for lipid-raft redistribution of the protein19. Translocation of this mutated version of
CD20 to the lipid rafts upon antibody binding is reduced by 75% as compared to the
wild type protein19. Here, cells expressing the wild type (WT) or mutated version of
CD20-GFP were treated with rituximab and the localisation of the fluorescent CD20
was compared (Figure 2E). Both the mutant and WT CD20 were equally frequently
capped in Daudi transfectants upon treatment with rituximab (70.4±5.6% vs.
66.9±6.4%, respectively) and the amount of mutant CD20 localised in the cap
(77.9±1.4%) was significantly lower than the amount observed for WT CD20 –
though the difference was very small (Figure 2F). Thus, redistribution of CD20 to
lipid rafts is not essential for capping of this protein caused by rituximab.
9
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Other proteins selectively co-localise with CD20 in the cap
We next set out to test whether or not other proteins co-localised within the cap of
CD20 after rituximab treatment. Primary B cells or Daudi were incubated with
rituximab, then fixed and stained for the localisation of other proteins: (i) the integrin
ICAM-1; (ii) moesin, a member of the ezrin–radixin–moesin family that is involved in
cross-linking plasma membrane proteins such as ICAM-1 with the actin
cytoskeleton20; (iii) CD45, a protein tyrosine phosphatase abundantly expressed on
the surface of B cells, or (iv) surface proteins in general, marked with biotin (Figure
3A-B). Co-localisation of proteins relative to CD20 was analysed by confocal
microscopy and Pearson’s correlation coefficients calculated. Pearson’s correlation
coefficients are between 1 and -1 where 1 indicates high co-localisation and -1
indicates anti-correlation. The adhesion molecule ICAM-1 strongly co-localised with
capped CD20 both in Daudi (Figure 3A,C) and primary B cells (Figure 3B,D) with
correlation coefficient values of 0.73±0.01 and 0.74±0.02, respectively. Staining for
moesin was very weak in Daudi but in primary B cells, this protein also co-localised
with CD20 (correlation coefficient: 0.72±0.02 )(Figure 3B,D).
However, in contrast, CD45 did not cap with CD20 and remained uniformly
distributed throughout the plasma membrane (correlation coefficient: 0.30±0.02 and
0.32±0.02 for Daudi and primary B, respectively). Likewise, surface proteins in
general, visualised by biotinylation followed by staining with fluorescently-labelled
streptavidin, remained homogeneously distributed around the cell surface and the
level of co-localisation with CD20 was not high (correlation coefficient: 0.24±0.02 and
0.28±0.02 for Daudi and primary B, respectively) (Figure 3A-D). Thus, the
enrichment of surface proteins in the cap was selective.
The localisation of surface proteins in rituximab-treated cells is reminiscent of
polarised lymphocytes characterised by a differential localisation of proteins at the
leading edge and uropod - rather than cells in which the antibody has merely capped
its ligand. ICAM-1 and moesin are known to be strongly enriched in the uropod of
lymphocytes 21. Conversely, CD45 has been reported to be uniformly distributed on
the cell surface of polarised lymphocytes22. The chemokine receptor CCR7 is a
surface protein known to localise specifically to the leading edge of migrating
lymphocytes and thus, we assessed the localisation of CCR7 in relation to the cap of
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CD20 in cells treated with rituximab. CCR7 did not co-localise with CD20 and
commonly accumulated at the opposite side of the cell to the cap of CD20, both in
primary B cells and in Daudi, evidenced by a negative correlation coefficient: 0.11±0.03 and -0.20±0.03 for primary B and Daudi, respectively (Figure 3A-D).
Taken together, these data suggest that rituximab causes B cells to adopt a
polarised phenotype.
The MTOC polarises towards the CD20 cap
A characteristic consequence of cellular polarisation is a specific orientation for the
microtubule organising centre (MTOC). Here, to assess the localisation of the MTOC
relative to the CD20-rich cap, Daudi and primary B cells were incubated for 1 hour
with rituximab, and then fixed and stained for α-tubulin (Figure 4A,B). The relative
distance between the MTOC and the centre of CD20-enriched cap was measured
and a polarity index calculated as the ratio between distance of MTOC to the CD20
cap and the cell diameter (Figure 4B). 97% of Daudi and 92% of primary B cells had
a polarity index below 0.5, which indicated that the MTOC was almost always
polarised towards the cap (Figure 4B).
An intact microtubule network is required for CD20 capping
To determine whether the microtubule network was involved in facilitating the
reorganisation of cell surface proteins, cells were treated with the microtubuleperturbing drug nocodazole and analysed for the extent to which CD20 was capped
after treatment with rituximab. The activity of the drug was confirmed since after
nocodazole treatment the MTOC was visually undetected when cells were stained
with an anti-α-tubulin mAb (supplemental Figure 4). The frequency at which
rituximab caused CD20 to cap in cells was 64.3±3.5% and 66.7±1.8% for Daudi and
primary B cells respectively. But this reduced to 31.3±2.5% and 42.3±2.7% when
cells were also treated with nocodazole (Figure 4C-D). Nocodazole did not affect the
frequency at which cells not treated with rituximab sometimes exhibited a cap of
CD20 – 21.9±1.3% and 29.7±4.2% in untreated Daudi and primary B cells
respectively, and for the same cells treated with nocodazole this was 18.3±3.6% and
22.7±8.1% respectively. This indicates that an intact microtubule network is
important for rituximab-mediated CD20 reorganisation.
11
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Polarisation of B cells by rituximab influences the efficiency of target cell
killing
Treatment with rituximab had a profound impact on the organisation of proteins at
the plasma membrane of B cells, and we hypothesised that these changes could
influence the outcome of interactions with NK cells and the efficiency of ADCC. To
assess this, Daudi cells were pre-treated with rituximab prior to co-incubation with
primary NK cells for imaging by laser scanning confocal microscopy at 37°C. Target
cell death was visualised by positive staining for DNA (Sytox Blue). Killing of target
cells by primary unstimulated NK cells in the presence of rituximab was mostly due
to ADCC as in the absence of rituximab almost no killing was observed and blocking
the Fc receptor CD16 inhibited killing via rituximab (supplemental Figure 5). To
investigate whether or not the organisation of CD20 influenced the efficiency of
target cell killing, we characterised each cell-cell interaction according to the
organisation of CD20 on Daudi cells and the site of initial contact with NK cells
(Figure 5A-B and Video 1-3). Conjugates could be assessed as to whether or not
CD20 was uniformly distributed in the plasma membrane in target cells upon contact
with an NK cell. In 82.4% of all conjugates scored (n=119) CD20 was polarised. We
then assessed whether or not target cells were killed by NK cells and found that
unpolarised target cells, with a uniform surface distribution of CD20, were killed
much less efficiently. Specifically, 42.9% of contacts between NK cells and
unpolarised target cells lead to target cell death within 20 minutes while, 68.4% of
contacts with polarised cells, in which CD20 was capped, lead to lysis (Figure 5C).
Thus, polarised B cells were killed more efficiently by NK cells.
Among cells in which CD20 was polarised, there was the possibility that (i) the cap of
CD20 was away from the initial site of the contact with the NK cell (44.9% of
contacts; n=98) or (ii) NK cells initially contacted target cells where CD20 was
capped (55.1%) (Figure 5B, Table 1). When CD20 was capped away from the site of
initial contact with the effector cell, 61.4% of cells were subsequently killed (Figure
5D). However, in conjugates where the NK cell initially contacted the target cells
precisely where CD20 was capped, the frequency of target cell lysis increased to
74.1%.
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From the movies, we also quantified whether or not the time required by the NK cell
to kill the target differed according to the topology of the cell-cell interaction and
whether or not target cells were polarised. The average time from initial contact to
target cell death - indicated by staining with a DNA dye - was between 3.5 and 4.1
minutes in all circumstances and no significant differences were observed (Figure
5E). This time-scale for NK cell-mediated cytotoxicity was similar for lysis of MICAexpressing target cells, killed through engagement of the NKG2D receptor on NK
cells, i.e. independent of ADCC, which took 2.6±0.6 minutes (Figure 5E). Thus, the
time for NK cell-mediated killing is likely relatively fixed by the time needed for cell
biological processes, including cytolytic granule release by the NK cell and apoptosis
in the target cell. Together these data establish that polarisation of B cells does not
alter the time needed for NK cells to kill target cells, but importantly, increases the
probability that the outcome of NK cell surveillance will be target cell lysis.
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Discussion
Rituximab was the first therapeutic mAb accepted for therapy of non-Hodgkin’s
lymphomas over 15 years ago3 and following this success, it has been since
introduced to treatments of other diseases where B cell depletion is desirable 1-2.
However, much debate remains as to its mechanism of action – perhaps involving
multiple lines of attack – one of which is ADCC. Here we investigated the sequence
of events taking place during the process of rituximab-mediated ADCC from the
initial opsonisation of B cells to the eventual killing by NK cells.
First, we found that rituximab mediated capping of CD20 such that the protein
accumulated at one pole of the B cell. This was true across multiple B cell lines, and
primary B cells isolated from healthy donors, which suggests that this phenomenon
is common – though the process remains to be tested in cells isolated from primary
tumours such as lymphomas. Capping of CD20 was also observed in untreated
cells, albeit with a lower frequency, and it remains to be established whether or not
rituximab enhanced this process occurring naturally within B cells or triggered CD20
through an alternative route.
Surprisingly, capping of CD20 occurred independent of antibody cross-linking by a
secondary antibody, as usually required for protein capping. It has been previously
reported that rituximab induces or strengthens the association of CD20 with lipid
rafts 14,18,23-24 and this process has been suggested to be at least partially
responsible for rituximab effectiveness in mediating B cell depletion, especially
through complement-dependant cytotoxicity. The capping of CD20 observed here,
however, was not dependent on an association with lipid rafts, since a mutant variant
of CD20 that does not associate with lipid rafts was still capped by rituximab. In
addition, it has been shown that mAb 2H7 as well as its monovalent Fab fragment
are both able to mediate the association of CD20 with the lipid rafts 23. Here, 2H7, or
a monovalent version of rituximab, were unable to cause CD20 to cap. Thus,
association of CD20 with lipid rafts is not sufficient to cause its polarisation.
Unexpectedly, we found that the action of rituximab was not to merely cluster its
CD20 ligand but to more generally rearrange several proteins at the B cell surface.
Specifically, ICAM-1 and moesin co-localised with CD20 in the cap while others such
as CCR7 segregated away from CD20. Furthermore, intracellular cellular changes
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are caused by rituximab as evidenced by a specific orientation of the MTOC towards
the CD20-rich cap. Indeed, an intact microtubule network was required for CD20
capping. Thus, rituximab mediates a polarisation of B cells.
The mechanism by which this occurs is an important unknown, but we can speculate
a model based on our findings and previous research. CD20 has been previously
shown to reside in the plasma membrane of cells as homo-tetramers 24-25. Thus,
binding to CD20 rituximab may interact with neighbouring tetramers, cross-link them
and bring them together24. Multiple tetramers could be then assembled in a form of
lattice by rituximab leading to the creation of clusters. An additional process,
dependent on the microtubule network, may aid the coalescence of CD20 clusters
into a cap and more broadly, triggers cellular polarisation, likely responsible for
ICAM-1 and moesin being recruited to the cap of CD20 while CCR7 is excluded. It
has been previously shown that rituximab is able to induce signals resembling BCR
stimulation26, perhaps due to a functional association of CD20 with BCR26-27. Thus, it
is possible that rituximab causes B cell polarisation in manner that involves the BCR.
Consistent with this hypothesis is that we found that CD20 was not readily capped by
rituximab when expressed in Jurkat T cells (which obviously lack the BCR).
These data are functionally important because the reorganisation of the B cell
surface by rituximab influences the efficiency of target cell killing by ADCC. Polarised
cells are killed up to 60% more frequently than those with uniform distribution of
CD20; the most efficient killing of target cells takes place when NK cell contacts the
target directly on the side where CD20 is accumulated.
Many factors must be considered in the rational design of antibodies for use in
ADCC. These include their affinity to target antigen, little or no internalisation into
target cells, and efficient engagement of Fc-receptors on effector cells. Here we
describe yet another factor that could be taken into account: changes to the cell
surface organisation of the target cell. It may be important to consider screening
putative therapeutic antibodies for their ability to trigger protein clustering and cellular
polarisation.
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Acknowledgments
We thank J.P. Deans (University of Calgary) and M. Purbhoo (Imperial College
London) for DNA constructs, M. Mehrabi for isolation of PBMCs, M. Spitaler in the
Facility of Imaging by Light Microscopy for help with imaging and B. Kemp and the
IgG Purification Team at MedImmune Ltd for production and QC of reagents.
Research was funded by the Medical Research Council, a PhD studentship from the
Manchester Collaborative Centre for Inflammation Research (to AO), a Wolfson
Royal Society Research Merit Award (to DMD), and a Marie Curie European
Reintegration Grant (to DR).
Authorship Contributions
DR and AO performed experiments and analysed data; DKF and MAS helped design
the experiments and edited the manuscript; IS performed experiments on ImageStream
and analysed data; DJS and DCL created and provided a reagent; DR, MAS and DMD
conceived the project, designed experiments and DR and DMD wrote the
manuscript.
Conflict of Interest
DKF, IS, DJS, DCL and MAS are employees of MedImmune Ltd; a wholly owned
subsidiary of the Astrazeneca.
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Figure legends
Figure 1. Rituximab enhances capping of CD20 on B cell surface. Fluorescent
and bright-field images of B cell lines Daudi (A), 721.221 (B) and Raji (C) expressing
CD20-GFP or primary B cells (D) untreated or incubated with 10 μg/mL of rituximab
(left and right panels, respectively). Primary B cells were additionally labelled with
anti-CD20-AF647 antibody recognising the intracellular portion of the protein. Scale
bars represent 10 μm. (E) Capping of CD20 was quantified on images of
Daudi/CD20-GFP and primary B cells incubated in the absence or presence of 1
μg/mL and 10 μg/mL of rituximab. Graph represents mean ± SEM of three
independent experiments. (F) Capping of CD20 in Daudi/CD20-GFP cells was
quantified by flow microscopy. Cells were incubated alone or with 1 μg/mL or 10
μg/mL rituximab, then fixed and analysed by ImageStream multispectral imaging flow
cytometer. Graph represents mean ± SEM of two independent experiments.(G)
Quantification of the fraction of CD20 localised in the cap in Daudi/CD20-GFP and
primary B cells. 30-32 cells were analysed per condition. Data were analysed by 1way ANOVA with Bonferroni post-test. *** P<.001
Figure 2. Enhancement of CD20 capping is specific for rituximab and B cells.
(A) Daudi/CD20-GFP cells were incubated alone; or in the presence of 10 μg/mL
rituximab (Rtx) or 10 μg/mL mouse CD20-targeting antibody 2H7; or pre-incubated
with 20 µg/mL CD32-blocking antibody followed by incubation with 10 μg/mL
rituximab (Rtx+CD32); or incubated with 20 μg/mL of a monovalent version of
rituximab (Rtx mono). Number of cells with polarisation of CD20 was scored. Graph
represents mean ± SEM of three independent experiments. > 160 cells were
analysed per condition. (B) Fluorescent and bright-field images of Jurkat/CD20-GFP
cells untreated (left panels) or incubated with 10 μg/mL of rituximab with uniform
distribution of CD20 or CD20 capped on one side (middle and right panels,
respectively). (C) Capping of CD20 was quantified on images of Jurkat/CD20-GFP
cells incubated in the absence or presence of 10 μg/mL of rituximab. Graph
represents mean ± SEM of three independent experiments. 112-118 cells were
analysed per condition. (D) Amount of CD20 accumulated in the cap in Jurkat/CD20GFP cells pre-treated with 10 μg/mL rituximab was compared to the amount of CD20
accumulated in the cap in primary B cells pre-treated with 10 μg/mL rituximab. Data
were analysed by unpaired T test (two-tailed). 21-31 cells from 3 or 4 experiments
20
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were analysed per condition. (E) Daudi/CD20-GFP (CD20 WT) or Daudi cells
expressing a mutant form of CD20-GFP (CD20 mut) were left untreated or incubated
with rituximab then fixed and analysed by laser scanning confocal microscopy.
Proportion of cells with polarised CD20 is shown. Graph represents mean ± SEM of
three independent experiments. Data were analysed by unpaired T test (two-tailed).
> 200 cells were analysed per condition. (F) Amount of CD20 accumulated in the cap
in Daudi cells expressing a mutant form of CD20-GFP (CD20 mut) pre-treated with
10 μg/mL of rituximab was compared to the amount of CD20 accumulated in the cap
in Daudi/CD20-GFP (CD20 WT) cells pre-treated with 10 μg/mL rituximab. 31 cells
from 2 or 3 experiments were analysed per condition. Data were analysed by
unpaired T test (two-tailed). ** P<.01, *** P<.001
Figure 3. Other proteins co-localise with CD20 in the cap. (A) Daudi/CD20-GFP
and (B) primary B cells were incubated with 10 μg/mL of rituximab or rituximabAF633, respectively, for 1 hour and then fixed and stained for ICAM-1, CD45, CCR7,
moesin or biotinylated as indicated followed by incubation with secondary
fluorescently labelled antibodies. As a positive control in Daudi/CD20-GFP cells
CD20 was targeted by rituximab AF633 and co-localisation between green and red
fluorescent channels was calculated (A). As a positive control primary B cells were
additionally stained for CD20 using an antibody recognising the cytoplasmic portion
of the protein followed by incubation with secondary fluorescently labelled antibody
(B). Scale bars represent 5 μm. (C-D) Pearson’s correlation coefficients calculated
for co-localisation of CD20 and other cell components as indicated in individual
Daudi (C) or primary B cells (D) are shown. 18-46 cells were analysed per condition.
Figure 4. Microtubule network is involved in rituximab-mediated CD20
polarisation. (A) Fluorescent and bright-field images of Daudi/CD20-GFP cells
incubated with rituximab and labelled for α-tubulin (red). MTOC is identified as the
brightest spot in the red channel. Scale bar represents 10 μm. (B, left panel)
Schematic representation of a cell, in which MTOC polarisation towards CD20enriched region was assessed by calculating the polarity index values corresponding
to the ratio between the distance from MTOC to CD20 cap (a) and the cell diameter
(b). Distribution of polarity indexes in primary B cells (B, top right panel) and Daudi
21
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cells (B, bottom right panel) incubated with rituximab is shown. 74 Daudi and 84
primary B cells were analysed. Proportion of Daudi (C) and primary B cells (D) with
polarised CD20 in untreated cells or after incubation with rituximab and nocodazole.
Graphs represent mean ± SEM of three independent experiments. > 100 cells were
analysed per condition. Data were analysed by 1-way ANOVA with Bonferroni posttest. * P<.05, ** P<.01, *** P<.001
Figure 5. Surface organisation of CD20 influences efficiency of target cell
killing. Daudi/CD20-GFP cells were pre-incubated with 10 µg/mL of rituximab and
then mixed with freshly isolated primary NK cells and imaged for 40 – 60 minutes.
(A) A schematic representation (left panel) and 3D reconstructed snapshots from live
cell microscopy (right panels) of Daudi cells with unpolarised CD20 interacting with
and being killed by an NK cell (CD20 is green, lysotracker used to visualise lytic
granules within NK cells is red, Sytox Blue used to visualise dead cells is white). (B)
A schematic representation (left panels) and 3D reconstructed snapshots from live
cell microscopy (right panels) show examples of target cell killing in the context of
CD20 organisation when CD20 is capped: cells with CD20 capping away from the
site of contact with an NK cell (top panels) and cells with CD20 enriched on the side
of the initial contact with an NK cell (bottom panels). Scale bars represent 10 μm. (C)
Proportion of conjugates with or without capping of CD20, in which killing of a target
cell took place. (D) Proportion of conjugates with capping of CD20 away from the
initial contact or at the contact side, in which killing of a target cell took place. (E)
Time of conjugation between Daudi and NK cell leading to killing of the target was
quantified for each category as well as for Daudi-MICA in conjugates with NK cells.
22
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Prepublished online April 23, 2013;
doi:10.1182/blood-2013-02-482570
Rituximab causes a polarisation of B cells which augments its therapeutic
function in NK cell-mediated antibody-dependent cellular cytotoxicity
Dominika Rudnicka, Anna Oszmiana, Donna K. Finch, Ian Strickland, Darren J. Schofield, David C. Lowe,
Matthew A. Sleeman and Daniel M. Davis
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