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Rheumatology Advance Access originally published online on May 30, 2006
Rheumatology 2007 46(1):29-36; doi:10.1093/rheumatology/kel148
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© The Author 2006. Published by Oxford University Press on behalf of the British Society for Rheumatology. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Bone marrow B-lineage cells in patients with rheumatoid arthritis following rituximab therapy

M. J. Leandro1, N. Cooper2, G. Cambridge1, M. R. Ehrenstein1 and J. C. W. Edwards1

1Centre for Rheumatology and 2Department of Haematology, University College London, London, UK.

Correspondence to: Maria J. Leandro, University College London, Centre for Rheumatology, Windeyer Building, Room 317, 46 Cleveland Street, London W1T 4JF, UK. E-mail: maria.leandro{at}ucl.ac.uk


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 Conclusions
 Acknowledgements
 References
 
Objective. To assess the presence and phenotype of B-lineage cells in the bone marrow (BM) of rheumatoid arthritis (RA) patients after rituximab therapy.

Methods. Six patients were studied. BM aspirates were collected 3 months after the treatment and analysed using the four-colour flow cytometry.

Results. CD19+ (B-lineage) cells in BM samples varied from 0.1 to 3.25% in the lymphoid gate. CD34+ cells varied from 1.23 to 4.86%. The proportion of CD34+ cells committed to the B-lineage varied between 0 and 42.19%. Pro-B-cells were undetectable in one case. The majority of B-cell precursors were pro-B-cells in Patients 5 and 6 (50 and 62% of CD19+ cells, respectively), pre-B-cells in Patients 3 and 4 (64 and 70%) and immature B-cells in Patient 1 (44%). Detectable CD20 expression on CD19+ cells was either low or absent. Plasma cells varied from 0.01 to 0.36% of the total nucleated cells. There was a trend towards longer duration of clinical response in patients with evidence of more complete depletion in BM.

Conclusion. In this small cohort of RA patients treated with rituximab, differences in proportion and phenotype of CD19+ BM cells were detected. These differences suggest variation in the degree of depletion achieved and correlate with time to relapse. Although pro-B-cells are not targeted directly by rituximab as they do not express CD20, the levels were unexpectedly low.

KEY WORDS: Rheumatoid arthritis, Rituximab, B-cell depletion, Bone marrow


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 Conclusions
 Acknowledgements
 References
 
Rituximab is a chimaeric monoclonal antibody directed to CD20, a pan-B-cell surface marker, that has proven to be very effective in depleting normal and malignant B lymphocytes in vivo, and is widely used in the treatment of B-cell malignancies, particularly B-cell non-Hodgkin's lymphoma [1]. In the last 7 yrs, rituximab has also been used in the treatment of several autoimmune diseases with promising results [6, 2–5]. Rituximab is known to induce an almost complete depletion of normal B-cells in the peripheral blood that lasts usually between 6 and 9 months, but little is known about the quantitative and qualitative aspects of the depletion of normal B-cells achieved in solid tissues, including bone marrow (BM), lymph nodes and spleen [6, 7]. Studies in primates suggest that the depletion of normal B-cells in solid tissues is significant but not complete and that it varies from site to site and in different individuals even if treated with the same dose [8, 9].

There are indications that the degree and duration of B-cell depletion in the peripheral blood induced by rituximab may significantly influence the clinical response to treatment in patients with rheumatoid arthritis (RA) and systemic lupus erythematosus [10, 11]. In RA, failure to achieve at least 97% depletion of B-cells in the peripheral blood, lasting more than 3 months, was associated with no response to treatment. Also, patients who relapsed clinically at the time of B-cell return to peripheral blood showed a tendency to repopulate with a higher frequency of B-cells with a memory phenotype when compared with patients who only relapsed later (Leandro et al. submitted for publication). This suggests that relapse at the time of B-cell repopulation of the peripheral blood may be related to less efficient depletion of memory B-cells in solid lymphoid tissues.

We aimed to study whether the cells of B-cell lineage, including plasma cells, were present in BM aspirates of patients with RA 3 months after the treatment with rituximab. The cells of B-cell lineage in BM aspirates were studied using flow cytometry and a panel of monoclonal antibodies that allowed the distinction between the different CD20– and CDD20+ B-cell precursors, mature naïve B-cells and plasma cells. For ethical reasons, pre-treatment samples for comparison were not collected and the study focus was on qualitative aspects. Differences between patients were studied as well as the correlation between BM findings and time to B-cell repopulation of the peripheral blood, and clinical response to treatment, particularly the time of relapse.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 Conclusions
 Acknowledgements
 References
 
Patients
Six patients with active refractory RA entered the study. All patients were treated in the Department of Rheumatology at University College London Hospitals on the basis of clinical need. The study was approved by the hospital ethics committee. All patients gave informed consent before entering the study. Three patients were male and 3 female. The mean age was 57 yrs (range 37–62) and the mean disease duration was 19 yrs (range 10–27). All patients had erosive disease and were rheumatoid-factor-positive. Patients received two infusions of 1000 mg of rituximab given 2 weeks apart under steroid cover (two doses of 100 mg of methylprednisolone intravenously). All patients were being re-treated with rituximab, a mean of 23 months after the previous treatment (range 10–46). The patients were assessed before treatment, at 1 and 3 months after treatment. Before treatment and at 1 month after treatment, whole peripheral blood was collected for B-lymphocyte immunophenotyping. Three months after treatment, a BM aspirate was collected. A peripheral blood sample was collected at the same time of the BM aspirate to evaluate the degree of depletion of B-cell subpopulations in the peripheral blood and to help in excluding significant contamination of the BM sample with peripheral blood. A BM sample available from a lymphoma patient treated 2 yrs before with rituximab, with no evidence of disease at the time of sampling and full peripheral blood B-cell repopulation, was used for qualitative comparison with the RA patients as an index of the B-cell lineage profile expected at full reconstitution. Details of individual patients at entry to the study are given in Table 1.


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  		TABLE 1. Patients’ clinical characteristics

 
Monoclonal antibodies
Four-colour immunophenotyping of cells of B-cell lineage in BM aspirate was performed using matched combinations of anti-human murine monoclonal antibodies directly conjugated to fluorescein isothiocyanate (FITC), phycoerythrin (PE), phycoerythrin-Texas Red® (PE-TR), peridinin-chlorophyll a complex (PerCP) or allophycocyanin (APC). Combinations of anti-{kappa}(FITC), anti-{lambda}(FITC), anti-CD10 (APC), anti-CD19 (PE and APC), anti-CD20 (FITC and PerCP), anti-CD34 (PerCP), anti-CD38 (APC) and anti-CD138 (FITC) were used. For analysis of B-cells in the peripheral blood, combinations of anti-IgD(FITC), anti-CD19 (PE), anti-CD20 (FITC), anti-CD27 (FITC) and anti-CD38 (PE-Cy5) were used. All antibodies were purchased from Pharmingen (BD biosciences, San Diego, California).

Sample preparation
Twelve millilitres of BM aspirate were collected in tubes containing EDTA. Sample preparation was carried out within 2 h after sample collection. Red-cells were lysed by adding 30 ml of red-cell lysis buffer (Pharmingen, BD Biosciences) to 6 ml of BM aspirate diluted 3: 1 with phosphate-buffered saline (PBS) with 20% heat-inactivated foetal calf serum. The mixture was gently mixed and incubated for 6 min at room temperature. After centrifugation, the cell pellet was washed twice on PBS by centrifugation at 300g for 5 min and then resuspended on cold PBS with 2% heat-inactivated fetal calf serum. The cells were incubated with each monoclonal antibody combination at 4°C for 20 min (5 µl of cell suspension with 2 µl of each antibody or equivalent for 106 cells, as recommended by the manufacturer for the relevant batches). The samples were subsequently washed twice in cold PBS by centrifugation at 300 g for 5 min. The cells were then fixed by incubation with 50 µl of PBS with 2% paraformaldehyde for 5 min at room temperature. The cells were washed twice by centrifugation at 300 g for 5 min, and resuspended in 200 µl of cold PBS. The samples were kept protected from light at 4°C until analysis by flow cytometry. Analysis was carried out either on the same day or the day after the sample collection and preparation. Cell viability was checked before incubation with antibodies by the trypan blue assay. Peripheral blood samples were prepared using the same technique except that they were not diluted before red-cell lysis.

Flow-cytometry analysis
Data was acquired on a FACSCalibur (BD Biosciences Immunocytometry Systems) flow cytometer. Cellquest software was used (BD Biosciences Immunocytometry Systems). A sample with unstained cells was used as a negative control to compensate for the autofluorescence of cells. Samples incubated with only one antibody were used to compensate for the overlap between the different fluorochrome spectra. For BM samples, data was acquired until 40 000 events were collected in a lymphoid gate defined by a low to moderate forward-angle and low right-angle light-scattering properties (Fig. 1) [12]. For peripheral blood samples, a total of 20 000 events were collected in a lymphocyte gate similarly defined. Pro-B-cells were defined as CD19+CD10+CD34+{lambda}L/{kappa}L– cells, pre-B-cells as CD19+ CD10+CD34–{lambda}L/{kappa}L– cells, immature B-cells as CD19+ CD10+CD34–{lambda}L/{kappa}8461521L+ cells, mature B-cells as CD19+ CD10–CD34–{lambda}L/{kappa}L+ cells, all within the lymphoid gate and plasma cells as CD19+/–CD138+CD38+++ cells gated for the expected right-angle scatter (Fig. 2A) [12–19]. Results were expressed both as proportion of positive cells in total events and, where relevant, proportion of positive cells in lymphoid gate and total number of positive cells per 40 000 events in lymphoid gate.


Figure 1
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  		FIG. 1. Lymphoid gate used for analysis.

 

Figure 2
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  		FIG. 2. Gating for plasma cell analysis and phenotype of plasma cells in one of the samples. (A) Expected right-angle scatter profile for plasma cells in the bone marrow set around cells expressing high levels of CD38; (B) Expression of CD138 vs CD19 on CD38+++ cells with the expected right-angle scatter profile of plasma cells (Patient 1).

 
Statistical analysis
Limited descriptive statistics of results (median and range) were used, given the small number of samples. Correlations were determined by calculating Spearman rank correlation coefficient.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 Conclusions
 Acknowledgements
 References
 
B-cell depletion in peripheral blood
The total number of peripheral blood B-cells decreased a median of 97% at 1 month and a median of 99% at 3 months following the treatment with rituximab. At 3 months, the peripheral blood samples from all patients remained depleted of B-cells, showing a median of 0.75 x 106/l CD19+ cells, with more than 75% having a memory or plasma cell precursor phenotype (IgD–, CD27+) (Table 2).


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  		TABLE 2. Detailed findings on bone marrow sample and peripheral blood analysis 3 months after rituximab therapy

 
Presence of cells of B-cell lineage in BM aspirates 3 months after the treatment with rituximab
Despite similar degrees of depletion in the peripheral blood, the frequency of the cells of B-cell lineage in BM aspirates varied between patients (Table 2, Fig. 3A). Frequency of CD19+ cells in the lymphoid gate varied from 0.1 to 3.25% (median 1.04%). Patient 2 showed almost absence of CD19+ cells (0.1%), while Patients 1 and 3–6 showed signs of B-cell development to different degrees. There was no correlation between total B-cell numbers in the peripheral blood at baseline or at 3 months and total numbers of CD19+ cells in BM aspirate samples at 3 months.


Figure 3
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  		FIG. 3. Proportion of B-lineage cells in the lymphoid gate and phenotype of B-cells and B-cell precursors in different samples. (A) Proportion of CD19+ cells in the lymphoid gate in Patients 1–6 (%); (B) relative proportion of different B-lineage cell subpopulations expressed as a percentage of the total of CD19+ cells in the lymphoid gate in Patients 1 and 3–6.

 
Frequency of CD34+ cells (marker of stem cells and early precursors) in the lymphoid gate varied from 1.23 to 4.86% (median 1.77%) (Table 2). The proportion of CD34+ cells committed to the B-cell lineage (expressing CD19) varied between 0 and 42.19% of the total number of CD34+ cells in the lymphoid gate (median 15.08%).

Different B-cell precursor subpopulations predominate in different patients
Pro-B-cells were undetectable in Patient 2. The majority of B-cell precursors were pro-B-cells in Patients 5 and 6 (50 and 62% of CD19+ cells, respectively), pre-B-cells in Patients 3 and 4 (64 and 70%) and immature B-cells in Patient 1 (44%) (Table 2, Figs. 3B and 4). In Patients 1 and 3–6, the ratios between the frequencies of more mature B-cell precursors (pre-B-cells and immature B-cells) and more immature B-cell precursors (pro-B-cells) were 2.09, 2.33, 3.39, 0.58 and 0.58, respectively.


Figure 4
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  		FIG. 4. Expression of surface immunoglobulin light chains and of CD34 vs CD10 on CD19+ cells in the lymphoid gate. (A) Lymphoma patient after peripheral blood B-cell repopulation showing the expected B-lineage cells subpopulations; (B) Patient 4, showing a predominance of pre-B-cells (CD19+CD10+CD34–{lambda}L/{kappa}L–cells); (C) Patient 6, showing a predominance of pro-B-cells (CD19+CD10+CD34+{lambda}L/{kappa}L–cells).

 
Detectable expression of CD20 on B-cells and B-cell precursors in the BM was low or absent
Detectable CD20 expression on CD19+ cells, including immature and mature B-cells, was either low or absent (Fig. 5). Two different antibodies to CD20 gave similar results (data not shown).


Figure 5
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  		FIG. 5. Expression of CD20 vs CD10 on CD19+ cells in the lymphoid gate. (A) Lymphoma patient after peripheral blood B-cell repopulation showing the expected B-lineage cells subpopulations; (B) Patient 1, showing decreased CD20 expression.

 
Plasma cells
The proportion of plasma cells in the total of events collected varied from 0.01 to 0.36% (median 0.02%) (Table 2, Figs. 2 and 6). In the lymphoid gate, the percentage of CD19+ cells that were CD138+ varied between 3.05 and 57.14% (median 9.82%). No correlation was found between the proportion of CD19+ cells in the lymphoid gate or in the total of events collected and the proportion of plasma cells in the total of events collected in these BM samples. The proportion of plasma cells in the total of events collected did not correlate significantly with the serum total immunoglobulin levels at the time of BM sampling (IgA r = 0.12, P = 0.80; IgG r = 0.65, P = 0.18; IgM r = 0.38, P = 0.50).


Figure 6
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  		FIG. 6. Proportion of plasma cells (CD138+CD38+++CD19+/– cells) in total nucleated cells in different patients (%).

 
Clinical correlations
On further follow-up, patients who showed a higher relative proportion of more mature precursors in their BM (Patients 1, 3 and 4) relapsed clinically at B-cell return to the peripheral blood (at 5, 11 and 11 months, respectively) (Table 1). The proportion of plasma cells in the samples was higher in Patient 1, who relapsed earlier than the other patients. The other three patients did not relapse at B-cell return. Patient 2, who showed almost complete depletion, repopulated at 8 months and relapsed clinically at 18 months. Patients 5 and 6 repopulated at 11 and 6 months, respectively, have not yet relapsed (follow-up, 15 and 13 months, respectively).


   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
Conclusions
 Acknowledgements
 References
 
In this cohort of RA patients treated with rituximab, the frequency and phenotype of the cells of B-lineage in BM aspirate samples 3 months after treatment varied between patients despite similar degrees of depletion in the peripheral blood. In children treated for acute lymphoblastic leukemia, several authors have found differences in the pattern of regeneration of precursor B-cells that were related to the intensity of the preceding treatment and presumably to the degree of cell-killing induced [16, 20, 21]. van Lochem and collaborators [20] found that the ratio between ‘more mature’ and ‘immature’ precursor B-cells was <1.0 in regeneration after more aggressive therapy, between 1.2 and 2.8 after less aggressive therapy, and 6–8 after cessation of all treatment. In normal BM samples, the most immature subset is relatively small [16]. In this study, patients who relapsed clinically earlier (Patients 3, 4 and 5), at the time of B-cell return to the peripheral blood, showed higher proportions of more mature B-cell precursors than the patients who relapsed later (Patients 2 and 6). Patient 1, who showed an almost total absence of B-cell precursors, also relapsed later. This suggests that patients who relapsed earlier may have depleted less well in solid lymphoid tissues.

There are potential problems with BM immunophenotyping studies using flow cytometry that involve sampling and dilution of samples with fat and contaminating peripheral blood. For this reason, we have expressed our findings both as percentage of total events collected (nucleated cells) and as percentage of cells in lymphoid gate. These results should be comparable but may be influenced by a low proportion of lymphoid cells in Patient 2 and a high proportion of lymphoid cells in Patient 4 (Table 2). In this study, significant contamination of BM aspirate samples by peripheral blood that influences findings for B-lineage cell populations can be excluded as peripheral blood samples collected at the same time showed almost complete B-cell depletion.

It is interesting that despite the presence of B-cell precursors in the BM in five of the samples, B-cell repopulation of the peripheral blood was only seen 2–8 months later. It is possible that free rituximab is still available and that it prevents full regeneration of the BM until its complete clearance [22]. Our finding that CD20 expression by cells of B-lineage in the BM samples was either low or absent could be explained by bound rituximab still being present and preventing full detection of the CD20 antigen by monoclonal antibodies. The majority of anti-CD20 antibodies available recognizes similar or overlapping epitopes and demonstrates substantial cross blocking [23, 24]. Other authors have also reported that the CD20 antigen cannot be detected by flow cytometry on the surface of B-cells following treatment with rituximab and suggested that this is due to the masking of CD20 by rituximab, and the results reported by Jilani et al. [25] strongly support the case. They have also shown that there was transient down-modulation of CD20 surface expression in some CLL patients treated with rituximab, but there is no data on whether this can happen in normal B-cells and no suggestion that this is a major mechanism for resistance to rituximab treatment in lymphoma.

Some authors have found variable expression of low or intermediate levels of CD20 by pro-B-cells [16], but the majority of pro-B-cells do not express CD20 [17, 26]. Therefore, pro-B-cells should not be depleted by rituximab, and as B-cell formation occurs throughout life in humans, they should be present at least in normal numbers and proportionally increased. In mice treated with anti-CD20 monoclonal antibodies, B-cell precursors not expressing CD20 were not depleted from the bone marrow [27, 28]. Unexpectedly, in Patient 2 no pro-B-cells were detected, and we did not see a consistent proportional increase on pro-B-cells in our patients. This suggests that pro-B-cell survival may be affected following treatment with rituximab, which would help to explain why B-cell repopulation after rituximab treatment takes longer than that after BM transplantation or myeloablative chemotherapy. In patients with relapsed, low-grade non-Hodgkin lymphoma, recovery of B-cell counts in the peripheral blood started 6–9 months after the completion of therapy, and normal levels were obtained after 9–12 months [29]. In a phase II RA study, B-cell depletion lasted more than 6 months [3]. In our own RA cohort, B-cell repopulation of the peripheral blood usually occurs between 6 and 9 months after treatment but can also occur later (up to 12 months) [10]. After autologous BM transplantation, the period of profound peripheral B-cell depletion lasts usually only 1–3 months [30].

It is possible that both prolonged availability of rituximab and some depletion of pro-B-cells explain the differences described above. It is difficult to understand why pro-B-cells survival should be affected by rituximab therapy. Studies in mice suggest that interactions between B-cell precursors may play an important role in promoting their own development, but to our knowledge no similar data exist in humans [31]. Also, in patients with X-linked agammaglobulinaemia, where there is a maturation defect occurring after the pro-B-cell stage, pro-B-cells develop normally and accumulate in the bone marrow [32]. It is possible that human pro-B-cells do express very low levels of CD20 and that rituximab is retained in the bone marrow for prolonged periods of time and mediates specific death signals to developing B-cell precursors, including pro-B-cells. Alternatively, the absence of detectable pro-B-cells in one of the patients may reflect different numbers of B-cell precursors and different capacities for BM regeneration between patients, with time to full generation of new B-cells determined by this regenerative capacity and clearance of rituximab.


   Conclusions
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Conclusions
Acknowledgements
 References
 
In our peripheral blood studies, we observed that patients who relapsed earlier, at the time of B-cell return to the peripheral blood, tended to have higher frequency and higher total numbers of circulating memory B-cells at repopulation, suggesting that the depletion of memory B-cells in peripheral solid lymphoid tissues may have been less extensive in these patients (Leandro et al. in press). The findings on BM aspirate samples presented here suggest a similar picture. In this small cohort there was a trend towards longer duration of clinical response to treatment in patients whose BM findings suggested higher degrees of depletion. Depletion in the bone marrow may be a surrogate for depletion of memory B-cells in solid lymphoid tissues. Nevertheless, our studies suggest that the difference between patients who relapse at the time of B-cell return and patients who relapse only later is not only quantitative but also qualitative. Patients tend to show the same pattern of relapse after repeated cycles of treatment. It is possible that differences between patients in the regeneration capacity of different B-cell populations after treatment with rituximab contribute to this picture.

Formula


   Acknowledgements
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Conclusions
 Acknowledgements
References
 
M.J.L. was supported by a research grant from GlaxoSmithKline Research & Development Limited and has received honoraria from Roche Portugal. J. C. W. E. is in receipt of financial support from Roche Products and GlaxoSmithKline Research & Development Limited. The other authors have declared no conflicts of interest.

The authors have declared no conflicts of interest.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Conclusions
 Acknowledgements
 References
 

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Submitted 13 September 2005; revised version accepted 31 March 2006.
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