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. Bohnhorst2,*
*
Institute of Immunology and
Centre for Rheumatic Disease, Rikshospitalet, University Hospital, University of Oslo, Oslo, Norway
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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–4), suggesting that some GC
founder cells are circulating. No PB B cells resembled the Bm3 and Bm4
GC cells. Remarkably, some cells with the
CD38-IgD+ phenotype, previously known as naive
Bm1 cells, expressed CD27. The CD38-IgD+
subpopulation therefore includes both naive Bm1 cells and
IgD+ memory B cells. This new classification of B cell
developmental stages reveals disturbances in the proportions of B cell
subpopulations in primary Sjögren’s syndrome (pSS) patients
compared with healthy donors and rheumatoid arthritis patients.
Patients with pSS contained a significantly higher percentage of B
cells in two activated stages, which might reflect a disturbance in B
cell trafficking and/or alteration in B cell differentiation. These
findings could be of diagnostic significance for
pSS. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Analyses of B cells in the bone marrow and secondary lymphoid tissues
have revealed a broad range of cell surface markers defining B cell
subpopulations, but only a few of these (mainly IgM and IgD) have been
used to analyze B cell subpopulations in PB. Mutational analyses of PB
B cell subpopulations defined by IgM and IgD have led to the conclusion
that PB B cells consist of unmutated naive B cells (
60%) and
somatically mutated memory B cells (40%; Ref. 16).
Recently, CD27 was found to be a useful marker for PB B cells that have
gone through somatic hypermutation and was proposed as a new marker for
memory B cells (17, 18, 19, 20, 21). This led to division of memory B
cells into four subpopulations, all expressing CD27 but differing in
their expression of IgM and IgD (19).
In secondary lymphoid tissue, CD38 and IgD have been useful in classifying important developmental stages from naive to memory B cells (Bm1–Bm5) (11, 12). Cell surface markers such as CD10, CD44, CD77, CD23, IgM, and CD95 have been used in combination with CD38 and IgD for defining seven tonsillar B cell subpopulations (13, 14, 22). Analyses of mutational frequency, functional properties, and Ig class switching have shown that the Bm1–Bm5 classification is an apposite method of defining peripheral B cell developmental stages (11, 13, 14, 22). Immunohistochemical staining of tonsillar sections have confirmed that the Bm1–Bm5 B cell subpopulations are localized in different functional areas of the tonsil (14). Thus, using this Bm1–Bm5 classification to recognize B cell developmental stages in PB would be of a great advantage. Furthermore, the use of CD27 could possibly solve a limitation of the Bm1–Bm5 classification in not having a positive marker for memory B cells.
The aim of this study was to analyze both PB and tonsillar B cells using the Bm1–Bm5 classification system in combination with CD27 to compare the proportion of B cell subpopulations in these two compartments. The generation of Ag-specific memory B cells and plasma cells takes place in several B cell differentiation stages (13). We were thus particularly interested in comparing the normal PB B cell subpopulations with those seen in autoimmune diseases such as rheumatoid arthritis (RA) and primary Sjögren’s syndrome (pSS). Possible alterations in the PB B cell subpopulations in these diseases could be of use as diagnostic aids and provide information on the disease pathology.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PB samples were collected from 12 healthy donors (10 female, 2 male; mean age 53 years; range 27–66 years). Tonsils were obtained from five patients with recurring tonsillitis after routine surgical removal at the Lovisenberg Hospital, Oslo, Norway.
PB from 11 patients (9 female, 2 male; mean age 49 years; range 20–73
years) with pSS were collected during routine examination at The
National Hospital (Oslo, Norway). All these patients fulfilled the
European criteria for pSS (23, 24), and clinical,
laboratory, and medication data are included in Table I
. Finally, PB were obtained from nine
patients with RA (eight female, one male; mean age 45 years; range
25–73 years), fulfilling the revised American College of Rheumatology
criteria for RA (25). The laboratory, clinical, and
medication data for each RA patient are shown in Table II. Most of the PB samples were taken in
connection to routine examination at the Center for Rheumatic Disease,
at the National Hospital, or in connection to orthopedic surgery
(patients R2 and R3) at the Diakonhjemmet Hospital (Oslo, Norway).
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Heparin or citrate peripheral blood samples were collected from the donors and centrifuged at 560 x g for 25 min. The buffy coat layer was diluted in a 10% solution of citrate-phosphate-dextrose (Sigma, St. Louis, MO) in PBS (Life Technologies, Paisley, U.K.) and centrifuged for 10 min at 450 x g. The cells were diluted in PBS and layered on Isopaque-Ficoll separation medium (Lymphoprep; Nycomed, Oslo, Norway) and centrifuged for 25 min at 450 x g. T cells were depleted from purified mononuclear cells (MNC) by rosetting with aminoethylisothiouronium bromide (AET)-treated SRBC and a second round of Isopaque-Ficoll separation (26).
The tonsils were minced, diluted in PBS, and filtered through a 70-µm pore size nylon filter (BD Biosciences, San Jose, CA) to deplete larger cells. The filtered cells were washed twice in PB and diluted in PBS. The cell suspension was layered on Isopaque-Ficoll separation medium (Lymphoprep) and centrifuged for 25 min at 450 x g. T cells were depleted from the accumulated MNC by rosetting with AET-treated SRBC and a second round of Isopaque-Ficoll separation.
Monoclonal Abs
Cells were stained with mAb specific for the following human cell surface markers: CD19 (B cell specific: allophycocyanin, IgG1, J4.119), CD38 (R-PE linked to cyanin (PC5), IgG1, 679.1 Mc7), CD95 (Fas; FITC, IgG1, UB2), CD23 (FITC, IgG1, 9P25), IgG1 (allophycocyanin, PC5, and PE, isotypic control, 679.1 Mc7), rat anti-human CD77 (unconjugated, IgM, 38-13) followed by goat anti-rat IgM (FITC, F(ab')2), all from Immunotech (Luminy, France); CD3 (T cell specific: FITC, IgG2a, 33-2A3), IgM (PE, IgG2a, IIE2), IgG1 (FITC, isotypic control, 1B9), IgG2a (PE, isotypic control, 5A7), all from Diatec (Oslo, Norway); IgD (PE, IgG2a, IA6-2); CD27 (PE, IgG1, M-T271), IgG2a (FITC, isotypic control, G155–178), all from BD PharMingen (San Diego, CA); IgD (FITC, F(ab')2, 107), CD10 (PE, IgG1, SS2/36), CD44 (FITC, IgG1, DF1485) from DAKO (Glostrup, Denmark).
Multiparameter flow cytometric analysis
Isolated and T cell-depleted MNC were stained for CD19, CD38,
and IgD in combination with the following cell surface markers: CD10,
CD44, CD77, CD23, IgM, CD27, and CD95 (the CD markers with their
expression and function on B cells are summarized in Table III). To assure there were no T cells in
the CD19-positive B cell populations, we also stained for CD3 in
combination with CD19. The cells were incubated with mAb (20 µl) for
30 min, washed twice in PBS containing FCS (2%), and fixed with
paraformaldehyde (1%). Data were collected using a FACSCalibur
analytical instrument, and four-color analyses were performed by
CellQuest software (BD Biosciences) or WinMDI
(http://tacs.scripps.edu/).
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T cells were depleted from the isolated MNC suspension by rosetting with magnetic Dynabeads coated with a primary mAb specific for human CD3 (Dynal Biotech, Oslo, Norway). The procedure was performed as recommended in the product protocol. The cell suspensions were stained for CD19 and CD27 or CD19, CD38, and IgD, as described above, but the cells were kept on ice and in medium (RPMI containing 10% FCS) instead of fixation. The CD19+ B cells were sorted into CD27+ and CD27- subpopulations and collected into two tubes on a FACSVantage (BD Biosciences).
Molecular analysis of VH region from CD27+, CD27-, Bm1 (CD38-IgD+), and Bm2' (CD38++IgD-) PB B cells
The VH region sequences were obtained by extracting mRNA from the sorted B cells with oligo(dT)25-coated magnetic Dynabeads (mRNA direct kit; Dynal). First-strand cDNA was prepared by using a first-strand cDNA synthesis kit (Amersham Pharmacia Biotech, Piscataway, NJ) and a primer from the IgM constant region priming the 3' end (5'-AAGGGTTGGGGCGGATGC-3').
Second-strand synthesis and amplification of VH1, VH3, VH4, and VH6 regions were done by PCR using 2 µl of the single-stranded cDNA product in a final volume of 50 µl containing 200 µM (0.2 mM) concentrations of each dNTP, 40 pmol of the Cµ primer, 40 pmol of one of the VH primers, 5 µl of the 10x cloned PFU DNA polymerase reaction buffer, and 1.25 U PfuTurbo DNA polymerase (Stratagene, La Jolla, CA). Oligonucleotides specific for the 5' end of the VH families were used (VH1, 5'-CCATGGACTGGACCTGGAGG-3'; VH3, 5'-CCATGGAGTTTGGGCTGAGC-3'; VH4, 5'-ATGAAACACCTGTGGTTCTT-3', VH6, 5'-ATGTCTGTCTCCTTCCTCAT-3') (Eurogentec, Brussels, Belgium). PCR was conducted for 41 cycles under the following conditions: 2 cycles (98°C for 30 s, 48°C for 30 s, and 72°C for 45 s); 3 cycles (98°C for 30 s, 50°C for 30 s, and 72°C for 45 s); 3 cycles (94°C for 30 s, 51°C 30 s, and 72°C for 30 s); 33 cycles (94°C for 30 s, 53°C 30 s, and 72°C for 30 s); and a final extension of 72°C for 15 min by using GeneAmp PCR system 9700 (Applied Biosystems, Foster City, CA). PCR products were analyzed on a 1.2% agarose gel; bands were cut out of the gel and concentrated by QIAquick gel extraction kit (Qiagen, Chatsworth, CA). The blunt end PCR products of VH genes (2 µl) were cloned by using a Zero blunt TOPO PCR cloning kit for sequencing with chemically competent Escherichia coli (Invitrogen, Groningen The Netherlands). Plasmid DNA was purified from ampicillin-selected clones by a Wizard cap. Plus SV minipreps DNA purification system (Promega, Madison, WI). Plasmid DNA (1 µl) was checked for insertion of the VH PCR product by restriction analysis by cutting with EcoRI enzyme (10 U; Stratagene). An aliquot (4 µl) of positive restriction analyzed plasmid DNA was sequenced by using BigDye terminator cycle sequencing ready reaction kit (Applied Biosystems), and data were collected by a 377 DNA Sequencer and PBI prism software (Applied Biosystems). Nucleotide sequences were analyzed by Lasergene99 software (DNASTAR), Blast search (http://www.ncbi.nlm.nih.gov/), and V base (http://www.mrc-cpe.cam.ac.uk).
Statistical analyses
Percentages of cells expressing cell surface markers were described as means of the individuals in each group. Variation in each group was defined by SEM. Differences between healthy donors, pSS, and RA patient groups were compared by an unpaired two-tailed t test. When the SD was significantly different between the groups, a Welch-corrected t test was used. A value of p < 0.05 was considered to be statistically significant. The statistical calculations were performed with GraphPad InStat.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Staining for CD38 and IgD gave a subdivision of PB B cells into
five subpopulations compared with six subpopulations among the
tonsillar B cells (Fig. 1). These
subpopulations, named Bm1–Bm5, have been described in tonsils based on
the surface expression of CD38 and IgD in conjunction with CD10, CD44,
CD77, CD23, IgM, and CD95 (11, 13). We therefore compared
the expression of these markers in addition to CD27 on PB B cells with
that on tonsillar B cells, to determine which peripheral B cell
differentiation stages are also found in PB (
Figs. 1–3). The germinal
center (GC) B cell subpopulation
(CD38++IgD-) is
undetectable or absent in PB. However, surprisingly, a small proportion
of GC founder cells
(CD38++IgD+) was found
circulating in the PB. In addition, the staining of CD27 revealed new
information on the previously defined peripheral B cell
subpopulations.
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40% resting B cells
(Bm1, early Bm5, and Bm5) and a surprisingly high percentage (60%) of
activated B cells expressing IgD in addition to CD23 and/or CD38 (Bm2
and Bm2' + Bm34). Tonsils consisted of 30% resting B cells
(Bm1, early Bm5, and Bm5) and 70% activated B cells (Bm2, Bm2' +
Bm34, Bm3, and Bm4) (Fig. 2
). The CD38-IgD+ B cell subpopulation contains a significant IgD+ memory B cell compartment in addition to naive Bm1 cells, whereas the large, activated naive Bm2 subpopulation consists of almost all naive B cells
The CD38-IgD+
subpopulation, previously known as naive Bm1 cells, was found in both
PB and tonsils. Analysis of CD27 expression on these cells suggested
that this subpopulation might also include an
IgD+ memory B cell component, in both blood
(59%) and tonsil (37%) (
Figs. 2–4). We
confirmed that these were memory cells by
showing that CD27+ B cells were clearly
somatically mutated by analyzing the sequences from expressed
VH region genes of seven
IgM+CD27+ PB clones and eight
IgM+CD27- PB clones. All
CD27+ VH sequences showed
somatic mutations (Table IV). The mean
somatic mutation frequency of the IgM+CD27+
B cells was 3.1% (range 0.7–7.4%). Only one of the eight
CD27- B cell clones showed evidence of somatic
mutation. In addition, three sequences obtained from the
CD38-IgD+ B cell
subpopulation (IgM+, CD27 unknown) were all
mutated (mean 5.6%, range 1.0–12.9%; Table V).
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), with PB
cells expressing a significantly lower percentage than tonsil cells (PB
11%; tonsil 21%; p = 0.016). GC founder cells circulate in the PB, whereas GC cells do not circulate
Surprisingly, we identified a small subpopulation (3%) of
CD38++IgD+ or GC founder
cells (Bm2' + Bm34) in 9 of the 12 PB samples. A high percentage
of the CD38++IgD+ cells
expressed CD10 (80% in PB; 83% in tonsil; p = NS) as
expected for GC founder and GC cells. In contrast to the tonsillar
subpopulation, the expression of CD27 and CD77 was almost absent on the
PB GC founder population (CD77 PB 3% vs tonsil 23%, p
= 0.032; CD27 PB 3% vs tonsil 56%, p = 0.003) (Figs. 2A and 4). In addition to the previously described
sequencing analysis of CD27- and
CD27+ PB B cells discriminating between unmutated
and mutated cells, two sequences from the PB Bm2'
subpopulation were found to be unmutated (Table V),
excluding these cells from being memory B cells re-entering the GC. The
absence of CD38++IgD- B
cells in PB, confirms that GC cells do not circulate.
Two IgD- memory B cell subpopulations exist in both PB and tonsil (early Bm5 and Bm5), but a higher percentage of these cells are class switched and express CD95 in PB
The IgD- memory B cells could be divided
into two subpopulations, one subpopulation still expressing CD38
(CD38+IgD-, early Bm5) and
one that had down-regulated the CD38 expression
(CD38-IgD-, Bm5). The
percentages of CD27+ cells were high in these two
subpopulations from both tonsil and PB (Figs. 2 and 4). In addition,
CD10 expression was lost, and CD44 expression was up-regulated in both
PB and tonsil (Fig. 2A). More circulating
IgD- memory B cells are class switched than the
tonsillar IgD-memory B cells, in that IgM was
expressed on a significantly lower percentage of both
IgD- memory B cell subpopulations from PB. The
early Bm5 sub population contained 37% IgM+
cells in PB vs 59% in tonsils (p = 0.046) and for Bm5
the percentage of IgM expressing cells was 29% in PB vs 53% in tonsil
(p = 0.029). Moreover, the percentage
expression of the apoptosis related marker, CD95, was higher in the
circulating Bm5 subpopulation (PB = 18%, tonsil = 3%,
p = 0.0001).
Expression of CD27 on the Bm1–Bm5 subpopulations from PB and tonsil
CD27-expressing B cells were present in both PB and tonsils but at
significantly different percentages in all except one subpopulation
(the early Bm5) (Figs. 2A and 4). The percentage of
IgD+ memory B cells, expressing CD27, within the
CD38-IgD+ subpopulation
was higher in PB than in tonsil, but in PB the percentages were
dramatically lower in the next two B cell subpopulations (Bm2 and Bm2'
+ Bm34). In contrast, the tonsillar B cells showed almost the
same percentage of
IgD+CD27+ memory B cells in
the CD38- and the CD38+
subpopulations, whereas in the Bm2' + Bm34 subpopulations the
percentages of CD27+ cells were high. The
percentages of CD27+ B cells were high in both PB
and tonsil for the early Bm5 and the Bm5 subpopulations.
A significantly higher proportion of activated B cells is found in pSS patients than in RA patents and healthy donors
The CD38 and IgD staining pattern showed that the Bm2
subpopulation was the most abundant PB B cell subpopulation in pSS, RA
patients, and healthy donors (54–75%) (Fig. 5). The four other subpopulations each
accounted for between 2 and 17% of CD19+ PB B
cells. There were remarkable and significant differences in the
relative sizes of the subpopulations among the pSS patients compared
with both healthy donors and RA patients. The Bm2 subpopulation was
significantly larger for patients with pSS than for healthy donors (pSS
75%, healthy donors 59%; p = 0.031). In contrast, the
CD38-IgD+ (including Bm1
and the IgD+ memory B cells) and the early Bm5
subpopulations comprised a significantly lower percentage of PB B cells
in pSS patients (CD38-IgD+
6%, early Bm5 7%) than in healthy donors
(CD38-IgD+ 15%,
p = 0.013; early Bm5 14%, p = 0.031).
There were no significant differences in the sizes of the PB B cell
subpopulations between RA patients and healthy donors. However, the GC
founder, Bm2' + Bm34 subpopulation was observed in only 5 of 9 RA
patients and 9 of 12 healthy donors, whereas it was seen in all 11
patients with pSS.
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). The percentages of
CD38-IgD+ (pSS = 6%,
RA = 17%), early Bm5 (pSS = 7%, RA = 13%), and Bm5
(pSS = 5%, RA = 13%) subpopulations were significantly
lower in pSS than in RA (p = 0.0001, pSS =
p = 0.026 and p = 0.002, respectively),
whereas the Bm2 (pSS = 75%, RA = 54%) and the Bm2' +
Bm34 (pSS = 7%, RA = 2%) subpopulations were
significantly larger in pSS than in RA (p =
0.001, p = 0.033, respectively). Overall there was a
higher percentage of activated B cells (Bm2 and Bm2' + Bm34) in
pSS patients than in healthy donors and RA patients. |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Earlier studies of PB B cells using costaining of IgD and IgM have
identified only naive and memory subpopulations
(IgM-IgD-,
IgM+IgD-, and
IgM+IgD+) (19, 27). Different surface markers have been used to identify B cell
differentiation stages in secondary lymphoid tissue (tonsil). The
tonsillar B cell subpopulations were characterized in a system based on
staining of CD38 and IgD in conjunction with CD10, CD44, CD77, CD23,
IgM, and CD95 (11, 13) and named from Bm1 (naive) to Bm5
(memory). We have used this Bm1–Bm5 classification of B cell
subpopulations to identify and compare circulating B cell
differentiation stages in PB with those in tonsil. The CD38/IgD
staining by itself revealed five PB B cell subpopulations comparable
with five of the six CD38/IgD-defined B cell subpopulations detected in
tonsil. The GC cell subpopulation
(CD38++IgD-) was missing
in PB. In 1994, Pascual et al. (11) defined four
subpopulations only based on CD38/IgD expression, and later Grammer et
al. (28) could divide tonsillar B cells into the same six
subpopulations, as we observed, in addition to two plasma cell
populations (CD38+++). New mAb used for the
staining of CD38 give a better differentiation of CD38 expression and
thus reveal more subpopulations. The present comparisons using
additional surface markers within the CD38/IgD-defined subdivisions
revealed PB B subpopulations not previously known to circulate. In
addition, except for some minor differences in expression of cell
surface markers, the B cell
subpopulations in PB and tonsil seem to be highly homologous (Tables VI and VII).
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The lack of a specific memory B cell marker has been a limitation of the Bm1–Bm5 classification system. CD27 has recently been defined as a marker for somatically mutated B cells, and thus memory B cells, by two independent groups (17, 18, 19, 20). Therefore, we analyzed the expression of CD27 on the Bm1–Bm5 classified cells. Surprisingly, CD27 was expressed on more than one-half of the CD38-IgD+ PB B cells, previously defined as unmutated naive Bm1 cells. Sequencing analysis of VH gene segments from CD27+ and CD27- PB B cells of healthy donors confirmed that the CD27 marker discriminates between memory and naive B cells. This indicates that CD38-IgD+ B cells include memory/somatically mutated IgD+ B cells in addition to naive B cells. Moreover, three VH gene sequences obtained from the sorted CD38-IgD+ PB B cells were all mutated. We should also have obtained unmutated sequences from the VH genes analyzed as this population also contains the naive Bm1 cells. However, because CD27+ B cells are over-represented in the PB CD38-IgD+ subpopulation, the sequences are likely to have been obtained only from the CD27+ B cells. Previously, the CD38-IgD+ (Bm1) subpopulation has been found to contain only unmutated cells (11, 12). This discrepancy may be explained by the use of different mAb for CD38 staining in the two studies. The large CD38-IgD+ subpopulation examined in the previous study probably included both the CD38-IgD+ and the CD38+IgD+ (Bm2) subpopulations found in the present analysis. This combined population would therefore have a very low percentage of CD27+ cells. Thus the sequences obtained might only come from CD27- cells, due the low percentage of CD27+ cells. IgD+ memory or somatically mutated B cells have previously been described in both PB (19) and tonsils (21, 29), and these IgD+ memory B cells have possibly been "hidden" in the CD38/IgD classification system.
Some GC founder cells circulate
Surprisingly, some GC founder cells (CD38++IgD+) were identified in PB, indicating that these cells are circulating, contradictory to previous impressions (16). The expression of CD10 on most of the cells in this PB subpopulation corroborates the GC founder cell phenotype, because CD10 is a marker for GC and GC founder cells (22). In addition, all the CD38++IgD+ PB B cells were negative for CD27, confirming that they are not yet GC B cells or somatically mutated memory B cells re-entering the germinal center. Furthermore, two VH gene sequences obtained from Bm2'-sorted cells were found to be unmutated. In contrast, many of the tonsillar CD38++IgD+ B cells express CD27 and CD77, which suggests that these cells are starting to develop into GC cells (CD38++IgD-CD27+CD77+/early Bm3). The presence of circulating GC founder cells may reflect the activation stage of an ongoing immune response. Moreover, this suggests that GC founder cells may be able to migrate between distinct secondary lymphoid tissues via PB. The identification of memory B cells still expressing CD38 (early Bm5) in both PB and tonsil suggests that B cells may also be able leave GC and secondary lymphoid tissues at an earlier differentiation stage than previous believed.
The GC cells are not circulating, and the IgD- memory B cell subpopulations show some differences in the two compartments
Our data indicate that GC B cells are the only peripheral B cell differentiation stage that does not circulate, consistent with other studies showing that GC B cells down-regulate expression of chemokine receptors or lose responsiveness to certain chemokines (30, 31). CD77 is believed to be a regulator of BCR-induced apoptosis (32). Significantly, CD77 was not found on circulating B cells, suggesting that this regulation mechanism is required only in secondary lymphoid tissue. More memory B cells were class switched (IgM-) in PB than in tonsil, suggesting that memory B cells that are not class switched are more prone to die and/or more likely to be activated for a second round of GC reaction. Moreover, a surprisingly high fraction of the memory (Bm5) subpopulation in PB expressed the apoptosis-related marker, CD95 (33, 34), compared with the tonsillar Bm5 cells, suggesting that apoptosis may also be important in the circulating memory B cell compartment. Altogether, our analyses have revealed differentiation stages of peripheral B cells not previously known to circulate. This new knowledge gives an important basis for further analysis on peripheral B cell development in healthy and disease.
Disturbed proportions of PB B cell subpopulations in pSS patients, but not in RA patients
The utility of an expanded classification of PB B cell subpopulations can be addressed by analyses of patients with immunological diseases. RA and pSS are autoimmune rheumatic diseases characterized by chronic inflammation and infiltration of T and B cells in the affected joints of RA patients and the exocrine glands of pSS patients. Ectopic GC-like structures that might play a role in the generation of autoimmune B cells have been identified in the affected tissue of RA (35) and pSS (36) patients.
Remarkably, pSS patients showed different proportions of B cell
subpopulations compared with healthy donors and RA patients. GC founder
cells (Bm2' + Bm34) were detectable in all pSS patients but only
in 5 of the 9 RA patients and 9 of 12 healthy donors. The elevated
presentation of this subpopulation in pSS might be a result of a
frequent B cell activation in this disease. Furthermore, pSS patients
had a higher percentage of activated naive B cells (Bm2) and a lower
percentage of resting B cells (Bm1, the newly identified
IgD+CD27+ memory B cells,
early Bm5, and Bm5) than both healthy donors and RA patients. An
increase of memory B cells (37 , 38) and a
decrease of activated B cells (39) in PB during aging has
been described, but this could not explain the difference between the
groups given that the mean age and range were approximately the same
for all groups.
Rheumatoid factor (RF) is an important diagnostic marker
for both pSS (23, 24) and RA (25), and
elevated plasma levels of RF are associated with a severe disease
prognosis for pSS (40) and RA (41). Recently,
we found that PB B cells from patients with pSS produce RF and
anti-Ro/Sjögren’s syndrome A with a much lower degree
of somatic mutation (42, 43, 44) than RF from RA patients and
healthy donors (45). These findings might be explained by
the higher percentage of activated naive B cells (Bm2) and GC founder
cells (Bm2' + Bm34) found circulating in pSS, resulting from an
hyperactivation of the peripheral B cells. Furthermore, disturbances in
chemokine levels and the expression of their receptors on B cells,
important for GC formation and B cell migration, may influence
peripheral B cell homeostasis in pSS (30, 31, 46, 47, 48, 49).
No significant differences were found in the Bm1–Bm5 B cell subpopulations between RA patients and healthy donors. However, all RA patients in the present study were on least one of the following drugs: nonsteroidal anti-inflammatory drugs (NSAID); corticosteroids, methotrexate; and Salazopyrin. Corticosteroids can affect the expression of cell surface markers (50), and the therapeutic regimens may have masked disturbances in the proportions of B cell subpopulations in the RA group, although we could not see a systematic difference in the three RA patients not receiving corticosteroids or methotrexate. Two of the pSS patients were taking NSAID; however, no difference could be seen in the percentages of B cell subpopulations for those two patients compared with the other pSS patients not receiving NSAID.
A disturbance in B cell trafficking or an increased activation of B cells might explain the altered proportions of PB B cell subpopulations in pSS patients. In addition, there could be an alteration in peripheral B cell differentiation, leading to a bias toward plasma cell differentiation and thus fewer memory B cells in the PB of pSS. Further analysis of both the trafficking and the activation state of the PB B cells in pSS would clarify the role of B cells in the pathology of pSS. The identification of high percentage of activated B cells might be of diagnostic significance in pSS and would be of interest to analyze in other autoimmune disorders.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Janne . Bohnhorst, Institute of Immunology, Rikshospitalet, University Hospital, N-0027 Oslo, Norway. E-mail address: janne.bohnhorst{at}labmed.uio.no
3 Abbreviations used in this paper: Bm, mature B cell; PB, peripheral blood; pSS, primary Sjögren’s syndrome; RA, rheumatoid arthritis; GC, germinal center; MNC, mononuclear cells; AET, aminoethylisothiouronium bromide; RF, rheumatoid factor; NSAID, nonsteroidal anti-inflammatory drugs.
Received for publication September 25, 2000. Accepted for publication July 27, 2001.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, Significant difference (p < 0.05)
between pSS and RA. There were no significant differences between RA
and healthy donors.