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The Journal of Immunology, 2004, 172: 7432-7441.
Copyright © 2004 by The American Association of Immunologists

CD27 Is Acquired by Primed B Cells at the Centroblast Stage and Promotes Germinal Center Formation1

Yanling Xiao, Jenny Hendriks, Petra Langerak, Heinz Jacobs and Jannie Borst2

Division of Immunology, The Netherlands Cancer Institute, Amsterdam, The Netherlands


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Studies on human B cells have featured CD27 as a marker and mediator of the B cell response. We have studied CD27 expression and function on B cells in the mouse. We find that B cells acquire CD27 at the centroblast stage and lose it progressively upon further differentiation. It is not a marker for somatically mutated B cells and is present at very low frequency on memory B cells. Enrichment of CD27 among centroblasts and the presence of its ligand CD70 on occasional T and B cells in or near germinal centers (GCs) suggested a role for CD27/CD70 interactions in clonal B cell expansion. Accordingly, GC formation in response to influenza virus infection was delayed in CD27 knockout mice. CD27 deficiency did not affect somatic hypermutation or serum levels of virus-specific IgM, IgG, and IgA attained in primary and recall responses. Adoptive transfer of T and B cells into CD27/CD28–/– mice revealed that CD27 promotes GC formation and consequent IgG production by two distinct mechanisms. Stimulation of CD27 on B cells by CD28+ Th cells accelerates GC formation, most likely by promoting centroblast expansion. In addition, CD27 on T cells can partially substitute for CD28 in supporting GC formation.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Expression of TNFR family member CD27 is restricted to naive and activated CD4+ and CD8+ T cells and subsets of B and NK cells. Its membrane-bound ligand CD70 is confined to activated lymphocytes and mature dendritic cells (1, 2, 3, 4). In vitro studies have established that CD27 promotes expansion of newly activated T cells (1, 5, 6). In CD27–/– mice, generation and maintenance of CD4+ and CD8+ effector T cells in response to antigenic challenge is impaired (7). We have recently resolved that CD27 rescues activated T cells from death, and in this way, increments the yield of live T cells upon their clonal expansion at the site of priming. In addition, CD27 exerts a prosurvival effect on CD4+ and CD8+ T cells at tissue sites (8).

In humans, CD27 is induced by B cell receptor triggering and maintained long-term (9, 10). CD27+ B cells are predominantly found in germinal centers (GCs)3 and marginal zones (11, 12). It is considered a marker for memory B cells (13) based on the following observations: 1) among IgM+D+ B cells in blood and marginal zones, CD27+ cells contain somatic Ig gene mutations (12, 14); 2) Ig class switching is more frequent among CD27+ than CD27– B cells (9); 3) upon activation, CD27+ B cells secrete Ig more efficiently than CD27– B cells (15); and 4) cord blood B cells lack CD27, while the percentage of CD27+ B cells in blood increases with age (16). Although these findings are in line with CD27 being a hallmark of primed B cells, they do not classify all CD27+ B cells as memory B cells. In particular, CD27 was recently found at low levels on all GC B cells from human tonsils, and dramatically up-regulated upon their in vitro differentiation into plasma cells (17). In this study, we have analyzed in detail the expression of CD27 on B cells in the mouse and find it in line with a contribution of CD27 to centroblast expansion.

In vitro studies on human B cells indicate that CD27 can promote IgM, IgG, IgA, and IgE secretion (10, 16, 18, 19). Whether CD27/CD70 interactions enhance Ig production by delivering differentiation signals to B cells or by sustaining expansion of differentiating cells is unclear. Jacquot et al. (20) found that CD27 did not drive expansion of activated B cells, but promoted the generation of a plasma cell phenotype and IgG secretion. However, other data argue that CD27 does promote expansion of activated B cells (19, 21). All in vitro studies rely on deliberate stimulation of CD27 with CD70 transfectants. Whether CD27 signals are invoked in vivo will depend on availability of CD70. In both humans and mice, CD70 is induced by Ag receptor triggering in T and B cells (2, 3, 22). In humans, CD70 has been detected on B cells, which may represent recent GC immigrants (23). CD70 expression is very transient and plasma membrane levels are often extremely low, making it difficult to detect (3). In this study, we have used confocal laser microscopy to define at what point during the GC reaction CD27/CD70 interactions might play a part.

Comparing wild-type and CD27–/– mice, we have determined how CD27 contributes to the B cell response to influenza virus. An effect of CD27 deletion was expected, because anti-viral T cell responses in draining lymph nodes and the lung are severely impaired in CD27–/– mice (7). Apart from determining its impact on Ig production, we have studied the role of CD27 in GC formation. Present data suggest that CD27 promotes B cell expansion and/or differentiation into plasma cells. Alternatively or in addition, CD27 may promote the B cell response indirectly by facilitating Th cell expansion. The costimulatory receptor CD28 and its ligands play a key role in GC formation. Mice lacking CD28 function are greatly defective in Th-dependent IgG responses to hapten-protein conjugates, and lack obvious GCs (24, 25, 26, 27). To map a contribution of CD27 to the GC reaction relative to that of CD28, we generated mice genetically deficient for both CD27 and CD28, and performed adoptive transfer experiments with T and B cells from CD28–/– or CD27–/– mice. Collective experiments show that CD27 can support GC formation and Ig production via a major CD28-dependent route that proceeds via CD27 on B cells, and via a minor CD28-independent route that requires CD27 on T cells.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Mice

Wild-type, CD27–/– (7), and CD28–/– mice (24) were on a C57BL/6 background and used for experiments at 6–10 wk of age, in accordance with national and institutional guidelines. Offspring were genotyped by PCR and phenotype was confirmed by flow cytometry.

Flow cytometry

Spleen and lymph nodes were forced through a nylon mesh in IMDM with 8% FCS. Erythrocytes were lysed in 0.14 M NH4Cl, 0.017 M Tris-HCl (pH 7.2). Cells were incubated with Fc Block (2.4G2; BD PharMingen, San Diego, CA), washed in staining buffer (PBS, 0.5% BSA, 0.01% sodium azide), labeled as indicated, and analyzed using a FACSCalibur and CellQuest software (BD Biosciences, Mountain View, CA). Abs used were anti-mouse IgM, -IgD, pooled IgG (-1, -2a, -2b) mAb (Southern Biotechnology Associates, Birmingham, AL); anti-CD3{epsilon} mAb 145.2C11, anti-CD45R/B220 mAb RA3-6B2, anti-CD19 mAb 1D3, anti-CD27 mAb LG.3A10 (6), and anti-CD28 mAb 37.51 (from BD PharMingen). Dr. A. Rolink (University of Basel, Basel, Switzerland) kindly provided 493 mAb (28). Biotinylated peanut agglutinin (PNA) was from Vector Laboratories (Burlingame, CA).

Selection for IgM expression

IgM+ cells were selected by magnetic cell sorting. Spleen cells were incubated for 30 min on ice with 400 µl of anti-mouse IgM-microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany) per 107 cells. After washing, cells were resuspended in IMDM with 4% FCS, followed by positive selection using the autoMACSen (Miltenyi Biotec). The IgM-positive fraction was stained with biotinylated anti-IgD mAb followed by streptavidin (SA)-allophycocyanin, anti-IgM-PE, and anti-CD27-FITC. Alternatively, it was stained with biotinylated 493 mAb followed by SA-allophycocyanin, anti-IgD-PE, and anti-CD27-FITC. The IgM-negative fraction was stained with biotinylated anti-pooled IgG, followed by SA-allophycocyanin, anti-CD19-PE, and anti-CD27-FITC. The IgM-positive fraction contained >98% B cells, and the IgM– fraction contained ~3–4% B cells.

Analysis of somatic hypermutation (SHM)

Wild-type and CD27–/– mice were immunized i.p. with 50 µg chicken {gamma} globulin conjugated to (4-hydroxy-3-nitrophenyl) acetyl in alum. B cells were enriched from spleen by means of MACS using anti-CD19–microbeads (Miltenyi Biotec). For analysis of SHM in CD27–/– mice, the MACS-sorted population was stained with biotinylated PNA, followed by SA-allophycocyanin and PE-conjugated anti-V{lambda}1/2 mAb LS136 (29). PNAhigh V{lambda}1/2+ lymphocytes were sorted in the presence of propidium iodide on a FACStarPlus (BD Biosciences). For analysis of SHM in CD27+ and CD27– GC B cells of wild-type mice, the MACS-sorted population was stained with biotinylated PNA, followed by SA-PerCP, anti-CD19-Cy5, anti-CD27-FITC, and anti-V{lambda}1/2-PE. V{lambda}1/2+ cells, all expressing CD19, were analyzed for PNAhigh and PNAlow phenotype. Within the PNAhigh gate, CD27+ and CD27– cells were individually sorted. Single live cells were sorted into 96-well PCR plates, snap frozen, and stored at –20°C (29). Rearranged V{lambda}1/2 genes were amplified by PCR and 310 nt were sequenced using Bigdye Terminators Ready Reaction Mix (Applied Biosystems, Foster City, CA), in combination with the nested V{lambda}1/2 primer (29). Sequences were analyzed using DNAstar software (Madison, WI).

ELISA

Blood from the tail vein was collected in heparin-treated Microvette (Sarstedt Aktiengesellschaft, Nümbrecht, Germany) and centrifuged to obtain sera, which were stored at –20°C. Density gradient-purified influenza virus was incubated with 1% Triton X-100 at room temperature for 1 h and coated at 4°C overnight onto polystyrene 96-well flat-bottom microtiter plates (Nunc, Roskilde, Denmark) at 2 µg/well in 0.1 M sodium carbonate buffer (pH 9.6) with 0.01% Triton X-100. Subsequent incubations were performed at room temperature with washing in between. Nonspecific binding was blocked with PBS and 1% BSA for 1 h. Wells were incubated with sera, serially diluted in high performance ELISA buffer (HPE; Sanquin, Amsterdam, The Netherlands) with 1% BSA, followed by biotinylated goat anti-mouse IgM, -G1, -G2a, -G2b, -G3, or -A mAb (Southern Biotechnology Associates), diluted 1:2,500 in HPE with 1% BSA, and SA-conjugated HRP (Sigma-Aldrich, St. Louis, MO), diluted 1:10,000 in HPE with 1% BSA. Substrate 3,3',5,5' tetramethylbenzidin (Merck, Darmstadt, Germany) was added at 0.1 mg/ml (100 µl/well), together with 0.06% hydrogen peroxide in 0.1 M sodium acetate (pH 5.5). The reaction was stopped with 2 M H2SO4, and OD450 was read by a Wallac 1420 VICTOR2 multilabel counter (PerkinElmer Life Sciences, Boston, MA). Endpoint titers were expressed as reciprocal log3 of the last dilution, which gave an OD450 of ≥0.1 OD unit above the OD450 of the negative control (pooled serum from nonimmunized mice).

Immunohistology

Spleens were embedded in OCT compound (Tissue-Tek; Miles, Torrance, CA), frozen in liquid nitrogen, and stored at –80°C. Cryostat sections (6 µm) were mounted onto glass slides coated with 3-aminopropyltriethoxy-silane (Sigma-Aldrich). Slides were air dried for 1 h, fixed in ice-cold acetone, dried, and stored at –20°C. Sections were rehydrated in PBS for 30 min and immersed in PBS and 5% BSA for 30 min at room temperature. They were incubated in the dark overnight at 4°C in PBS and 1% BSA with mAb, which was directly conjugated to allophycocyanin, FITC, or PE, or biotinylated and detected by Texas Red (TR)-conjugated SA (Molecular Probes Europe, Leiden, The Netherlands). Abs used were anti-CD45R/B220 mAb RA3-6B2, anti-GL7 mAb GL7, anti-CD3 mAb 500A2, anti-CD4 mAb RM4-5, anti-CD8 53-6.7 (1:200; BD PharMingen), and biotinylated anti-follicular dendritic cell (FDC) mAb FDC-M2 (30) (20 µg/ml; AMS Biotechnology, Oxon, U.K.). Slides were mounted in Vectashield (Vector Laboratories) and observed under a Leica TCS NT confocal laser-scanning microscope (Leica Microsystems, Wetzlar, Germany). For quantitative analysis of GC formation, spleen sections were stained with anti-B220-allophycocyanin and anti-GL7-FITC. For each data point, whole spleen cross-sections were analyzed. The total number of B cell follicles in each section was counted. B220/GL7 double-positive clusters within B follicles were defined as GCs (31). GC formation was expressed as the ratio of GCs per total number of B cell follicles.

Preparation of purified T and B cells and total splenocytes

Splenocytes were passed over nylon wool to remove adherent cells (Polysciences, Warrington, PA). For T cell purification, nonadherent splenocytes were incubated on ice for 30 min with anti-MHC class II mAb M5/114.15.2 (BD PharMingen), followed by 30 min incubation on ice with 100 µl of goat anti-mouse Ig-coated magnetic beads, and 20 µl of sheep anti-rat Ig-coated magnetic beads (Advanced Magnetics, Cambridge, MA) per 107 cells. Beads were removed by magnetic sorting. For B cell purification, nonadherent splenocytes were incubated for 30 min on ice with rat anti-mouse mAb GK1.5 to CD4 and Lyt.2 to CD8, followed by 30 min incubation on ice with 100 µl of goat anti-rat Ig-coated magnetic beads per 107 cells. Purity of the resulting cell populations was checked with anti-CD3{epsilon} and anti-B220 mAbs. Only preparations that contained >98% T or B cells were used.

Virus infection and adoptive transfer

Influenza virus strain A/NT/60/68 was obtained from the Department of Virology at Erasmus University (Rotterdam, The Netherlands) and stored in aliquots at –80°C. Mice were anesthetized and infected intranasally with 25 hemagglutinin units of virus in 50 µl of HBSS, or with 100 U for rechallenge. For adoptive transfer, total nonadherent spleen cells (50 x 106) or purified T or B cells (20 x 106) were resuspended in 200 µl of PBS and injected into the tail vein of recipient mice.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
CD27 is transiently expressed during the GC reaction, primarily at the centroblast stage

A detailed analysis of CD27 expression on B cells in the mouse had not previously been performed. To determine CD27 expression on B cells of naive and memory phenotype, we purified IgM+ B cells from spleens of mice, which had been infected with influenza virus 6 mo earlier. In this population, 0.15% of IgM+D+ cells and 5.65% of IgM+D– cells expressed CD27 (Fig. 1A). Because the IgM+D– cell population in the spleen contains both immature transitional B cells and memory B cells (32), we also used 493 mAb, which detects transitional B cells (28). Within the IgM+D– population, the minor 493+ subset lacked CD27, while 5.6% of the 493–IgM+D– population of memory phenotype B cells expressed CD27 (Fig. 1A, red gate). Within the IgM– fraction, 2.30% of IgG+ cells expressed CD27 (Fig. 1A). We conclude that CD27 is essentially lacking on transitional and naive B cells and present on a few percent of IgM+ and IgG+ memory phenotype B cells.



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  		FIGURE 1. CD27 expression is in line with a contribution to centroblast expansion. A, Pooled splenocytes of two mice infected with influenza virus 6 mo earlier were separated into IgM+ and IgM– subsets by MACS, triple stained with the indicated Abs, and analyzed within the indicated gates for fluorescence intensity. Numbers give the percentage of positive cells in the quadrants. The experiment is representative of three experiments. B, One million pooled splenocytes or DLN cells of four mice obtained at day 8 after infection were stained with PNA, anti-CD19, and anti-CD27 mAb, and analyzed. Within CD19+PNAhigh gates, FSC discriminates centroblasts and centrocytes. Percentages of positive cells per quadrant are indicated. C, At day 8 after virus infection, spleens were isolated, processed for confocal laser immunohistology, and stained with fluorescent Abs. C1 and C2 show the same triple-stained section. In C1, GC cells are detected with anti-GL7-FITC. In C2, B cells are detected with anti-B220-allophycocyanin, and arrows indicate some CD70+ B cells. In C3, another section is shown, where T cells are detected with anti-CD3-FITC. CD70 was detected by anti-CD70-biotin and TR-SA.

 
To examine CD27 expression on GC B cells, we stained spleen and lung-draining lymph nodes (DLN) at day 8 after influenza virus infection. CD19 was used as B cell marker and PNA as GC marker (33). Cells were examined for forward scatter (FSC) profile to discriminate between centrocytes and centroblasts (Fig. 1B, red rectangles). CD27 was found on only 0.2% of PNAlow non-GC B cells and ~13% of PNAhigh GC B cells in both spleen and DLN, indicating that it is acquired during the GC reaction. About half of the CD27+ GC B cells were centroblasts and the other half centrocytes. CD27 was found at 3- to 4-fold higher frequency among centroblasts than centrocytes (Fig. 1B), suggesting that CD27 is most abundant during the expansion phase of primed B cells.

To gain further insight into the GC B cell stage when CD27 is acquired, we examined somatic mutations in Ig genes of CD27+ and CD27– GC B cells of wild-type mice. At day 10 after immunization with haptenated protein Ag, B cells were purified from pooled spleens of two mice and sorted individually on the basis of a PNAhighV{lambda}1/2+CD27+ or -CD27– phenotype. Single-cell PCR with V{lambda}1/2-specific primers allowed determination of the frequency of somatic Ig gene mutations in each subset. For the CD27+ subset, 29 cells were analyzed, of which 4 contained mutations (14%). For the CD27– subset, 42 cells were analyzed, of which 12 contained mutations (29%). The actual mutation frequency, which is the percentage of nucleotides mutated among total nucleotides sequenced from mutated cells, was very similar for CD27+ and CD27– cells, namely 0.67% and 0.70%. These data argue that CD27 expression is not concordant with the presence of somatic mutations.

To examine CD70 expression, we used confocal immunohistology, because CD70 is difficult to detect at the plasma membrane (3). GL7 was used as a marker for GC B cells, because it performs well on fixed tissue (34). Staining for CD70 and GL7 revealed multiple CD70-positive cells in GCs, or in close proximity of GCs, in spleens of influenza virus-infected mice (Fig. 1C). Staining of the same section with B220 defined a significant proportion of CD70+ cells as B cells. CD70 was also found on T cells, which were located in the B cell follicle near or in GCs, typically in clusters (Fig. 1C).

Our data indicate that, in mice, CD27 is essentially lacking on transitional and naive B cells and acquired during the GC reaction at the centroblast stage. CD27 is not a marker for somatically mutated B cells in the mouse and is absent from most memory B cells. Expression of CD27 and CD70 is in line with a role for this receptor/ligand pair during the B cell expansion phase in GCs.

CD27 accelerates GC formation, but does not affect SHM or Ig production

Initial examination of GC formation at day 8 after influenza virus infection failed to reveal a defect in CD27–/– mice. However, when GC formation was followed kinetically, an impact of CD27 was found (Fig. 2A). At day 4 after infection, 32% of B cell follicles in wild-type mice showed GCs, while this was about 2-fold less in CD27–/– mice. At day 6, the frequency of GCs was still significantly lower in CD27–/– mice, but at day 8 it had reached the same level as in wild-type mice. Apparently, CD27 determines the kinetics of GC formation, but is not absolutely required for this response.



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  		FIGURE 2. In CD27–/– mice, GC formation is delayed but serum Ig levels are normal. A, At the indicated days after infection, spleens were isolated, processed for confocal laser immunohistology, and stained with fluorescent Abs to B220 as B cell marker and GL7 as marker for GC cells. The percentage of follicles containing GCs was determined as described in Materials and Methods, using two spleens per time point. Error bar indicates SD. B, At the indicated days after infection, serum titers of virus-specific Ig isotypes were determined by ELISA. At day 56, mice were reinfected (arrows). Each data point represents a measurement on pooled sera from four mice. Results are representative of two independent experiments.

 
To analyze whether CD27 affected SHM, wild-type and CD27–/– mice were immunized with haptenated protein Ag, and single-cell PCR was performed on V{lambda}1/2+ GC B cells isolated 7 days later. Twenty-nine V{lambda}1/2 PCR products derived from pooled cells of two CD27–/– mice were sequenced, of which 34% carried mutations. The actual mutation frequency was 0.60%. These figures are in the wild-type range (29). The V{lambda}1/2 mutations in CD27–/– cells showed an A-T bias, preferred transitions over transversions, and had a preference for RGYW motifs, as is characteristic of SHM in the wild-type situation (35). We conclude that CD27 is not required for the somatic diversification of Ig genes.

To examine the impact of CD27 deletion on Ig production, mice were infected with influenza virus and bled from the tail vein 0, 7, 14, 21, 28, and 56 days later. Influenza virus-specific IgM, -G1, -G2a, -G2b, -G3, and -A isotypes increased in response to infection, with a transient increase in IgM preceding the others (Fig. 2B). No major differences in responsiveness were seen between wild-type and CD27–/– mice. Virus-specific IgG and IgA levels were as high as the plateau level at day 56 after primary challenge (Fig. 2B), and remained so for at least 6 mo in both wild-type and CD27–/– mice (data not shown). Rechallenge at day 56 modestly increased some Ig isotypes, but responsiveness in wild-type and CD27–/– mice was similar (Fig. 2B). In conclusion, CD27 deletion does not have a detectable impact on Ig class switching, generation, and maintenance of virus-specific IgM, IgG, and IgA levels in primary and recall responses in this physiological model.

CD27 supports GC formation and IgG production in absence of CD28

To determine whether CD27 contributes to GC formation downstream of CD28, we examined responses to influenza virus in CD28–/– and CD27/CD28–/– mice. In the spleen of these mice, T and B cell areas were normal in size and appropriately separated (data not shown). To analyze GC formation, sections of spleen, isolated at day 8 after virus infection, were double stained with fluorescent Abs to B220 and GL7. B cell follicles of wild-type and CD27–/– mice contained GCs of similar size and frequency. In spleens of CD28–/– mice, GCs were still detectable, but these were small and low in frequency (Fig. 3A). Interestingly, in mice lacking both CD27 and CD28, GC formation in the spleen was completely abrogated (Fig. 3A). Apparently, CD27 can support GC formation in the absence of CD28, but these GCs are small and occur in low frequency as compared with wild-type.



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  		FIGURE 3. CD27 supports GC formation and IgG production in the absence of CD28. A, Spleens taken at day 8 after infection with influenza virus were stained with anti-B220-allophycocyanin and anti-GL7-FITC to detect GCs. Sections were examined by confocal microscopy. The percentage of follicles containing GCs was determined as indicated in Fig. 2A. B, Virus-specific serum Ig was measured as indicated in Fig. 2B. Data points represent OD450 values in ELISA of pooled sera from three mice per group. IgG1, -G2a, -G2b, and -G3 levels were measured at day 14 after infection when plateau levels were reached (see Fig. 2B). The experiment was repeated with very similar results.

 
At day 14 after influenza virus infection, when plateau levels are reached in IgG production (see Fig. 2B), IgG1, -G2a, and -G2b levels were 8- to 10-fold reduced in CD28–/– mice, as compared with wild-type and IgG3 levels reduced ~4-fold (Fig. 3B). However, significant IgG responses were still detectable. Interestingly, on the CD28–/– background, CD27 deficiency further reduced levels of all virus-specific IgG isotypes ~10-fold (Fig. 3B). Thus, CD27 can support the generation of IgG-secreting plasma cells in the absence of CD28. Apparently, the modest GC formation in CD28–/– mice is sufficient to produce significant IgG levels. In the additional absence of CD27, GC formation in spleens is undetectable and IgG production drops accordingly. The very low residual IgG production in CD27/CD28–/– mice may be attributed to B cell maturation in the small GCs that still form at low frequency in DLN of these mice (data not shown).

A CD28-independent CD27-driven pathway to deliver T cell help to B cells

Our data indicate that CD27 can partially substitute for CD28 to drive GC formation. Subsequent experiments were designed to determine through what cell-cell interactions this was achieved. We used CD27/CD28–/– mice as recipients and investigated to what extent GC formation in the spleen could be restored by adoptive transfer of T cells that were deficient for CD28, but proficient for CD27. As a positive control, CD27/CD28–/– mice were reconstituted with wild-type total nonadherent splenocytes, i.e., T and B cells. This permitted GC formation at a frequency approximating that in wild-type mice (Fig. 4A). Transfer of CD27+/+CD28–/– splenocytes allowed the formation of small GCs at low frequency, exactly as seen in CD28–/– mice (see Fig. 3A). Purified CD27+/+CD28–/– T cells had the same effect, indicating that GC formation in absence of CD28 requires CD27 on T cells, but not on B cells (Fig. 4A). In CD27/CD28–/– mice that received CD27+/+CD28–/– splenocytes, GL7+ B cells clustered next to the FDC network, as in wild-type mice. In addition, CD4+ T cells were present in these areas (Fig. 4B). These findings indicate that CD27 on T cells can promote the formation of small but normally organized GCs in the absence of CD28 (36, 37).



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  		FIGURE 4. GC formation in absence of CD28 is mediated by CD27+/+ T cells. CD27/CD28–/– mice were reconstituted by i.v. adoptive transfer of total nonadherent splenocytes (T and B cells) or purified T or B cells of the indicated phenotypes. Mice were infected at day 2 after transfer and examined 8 days after infection. A, Spleen sections were stained with anti-B220-allophycocyanin and anti-GL7-FITC, and examined by confocal microscopy to determine the percentage and size of GCs. The experiment is representative of two experiments. The cartoons indicate the theoretical possibilities of CD27, CD28, and CD70 expression on T and B cells in the adoptive transfer setting, based on these genotypes of the cells present. B cells from donor (Bd) and recipient (Br) are indicated separately. B, Detection of FDC and CD4+ T cells in GCs of CD27/CD28–/– mice, which had received CD27+/+CD28–/– splenocytes at day 8 after infection. Sections of adoptive transferred and wild-type control mice were stained with FDC-M2-TR (30 ) and anti-GL7-FITC, or with anti-CD4-allophycocyanin and anti-GL7-FITC, as indicated.

 
Production of anti-viral IgM, -G1, -G2a, -G2b, and -G3 was determined at days 7, 14, and 21 after infection in the reconstituted mice, and compared with that in CD27/CD28–/– mice or mice reconstituted with wild-type splenocytes (control). Titers of all IgG isotypes increased to plateau levels in this period, while IgM production peaked at days 7–14 (Fig. 5). Purified CD27+/+CD28–/– T cells restored IgG3 production almost to control levels, while it increased titers of the other IgG isotypes 3- to 6-fold, to a level that was ~3-fold lower than control. Apparently, the modest GC formation in these reconstituted mice allows for significant IgG production. We conclude that there exists a CD28-independent pathway to deliver T cell help for GC formation and IgG production, which requires CD27 on T cells but not on B cells.



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  		FIGURE 5. Ig production in absence of CD28 is mediated by CD27+/+ T cells. CD27/CD28–/– mice were reconstituted with purified T cells from CD28–/– mice or total nonadherent spleen cells (T and B cells) from wild-type mice, as indicated. Mice were infected at day 2 after adoptive transfer and serum Ig titers were determined at the indicated days after infection. Each data point represents a measurement on pooled sera from three mice.

 
CD27 on B cells accelerates CD28-dependent GC formation and facilitates IgG production

Subsequent experiments were designed to determine the contribution of CD27 to CD28-dependent GC formation. Bringing back CD27–/–CD28+/+ splenocytes into CD28–/– mice allowed formation of GCs with wild-type size and frequency within 8 days after infection (Fig. 6A). This was expected, because both CD27 and CD28 were present in the reconstituted situation. Reconstitution of CD27/CD28–/– mice with CD27–/–CD28+/+ splenocytes created a situation where CD28 was present but CD27 was absent. In this case, only small GCs were observed at day 8 after infection, which occurred in ~5-fold lower frequency than in the reconstituted wild-type situation (Fig. 6A).



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  		FIGURE 6. GC formation in presence of CD28 is accelerated by CD27 on B cells. A, CD28–/– or CD27/CD28–/– mice were reconstituted with total nonadherent spleen cells (T and B cells) from CD27–/–CD28+/+ mice, and GC formation was determined at day 8 after infection. B, CD27/CD28–/– mice were reconstituted with CD27–/–CD28+/+ T cells and wild-type (CD27+/+) B cells or CD27–/–CD28+/+ T cells only. GC formation at day 8 is shown. Cartoons represent what is indicated in Fig. 4. C, Quantitative representation of GC incidence at days 8, 14, and 20 after infection in CD27/CD28–/– mice that were reconstituted as indicated. Means ± SD are derived from three mice per time point.

 
Next, we examined whether it was CD27 on T cells or CD27 on B cells that supported CD28-dependent GC formation. CD27/CD28–/– mice received purified CD27–/–CD28+/+ T cells and wild-type (CD27+/+) B cells, or purified CD27–/–CD28+/+ T cells only. In the animals that received both CD27–/–CD28+/+ T cells and CD27+/+ B cells, GCs were of normal size, but somewhat lower in frequency than in the reconstituted wild-type situation at day 8 after infection. However, at day 14 the frequency was normal (Fig. 6, B and C). In mice that received purified CD27–/–CD28+/+ T cells only, a more significant deficit was observed. At day 8 after infection, GCs were small and infrequent (Fig. 6B). Throughout the next 2 wk, GCs increased in size and frequency to finally reach the wild-type situation (Fig. 6C). We conclude that in this adoptive transfer setting, GC formation is slightly delayed when CD28+/+ Th cells act in the absence of CD27 on T cells. In absence of CD27 on B cells, GC formation is dramatically delayed, but not defective. In a wild-type adoptive transfer setting, GC formation is complete at day 8 after infection (Fig. 6C), while this situation is only reached at day 20 in absence of CD27 on B cells. We conclude that GC formation as driven by CD28+ T cells is accelerated in particular by CD27 on B cells, while CD27 on T cells also makes a small contribution.

Virus-specific serum IgG production in CD27/CD28–/– mice reconstituted with CD27–/–CD28+/+ T cells and CD27+/+ B cells was as efficient as the response in wild-type mice (Fig. 7). This emphasizes that CD27 on T cells makes a nondetectable contribution to the B cell response when CD28 is present. However, this experiment clearly revealed a contribution of CD27 on B cells to CD28-dependent Ab production. Although the IgM response was normal in reconstituted mice that lacked CD27 on B cells, the IgG responses were delayed. IgG1 production had reached wild-type plateau levels at day 21, but the other isotypes were still at a significantly lower level at this time point (Fig. 7). This is in line with GC formation just reaching its optimum at day 20 after infection (Fig. 6C). In conclusion, CD28-dependent IgG production does not require CD27 on T cells, but is facilitated by CD27 on B cells.



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  		FIGURE 7. Ig production in the presence of CD28 is facilitated by CD27 on B cells. CD27/CD28–/– mice were reconstituted with purified CD27–/–CD28+/+ T cells, plus purified wild-type or CD27–/– B cells. Mice were infected with influenza virus at day 2 after adoptive transfer, and serum Ig titers were determined at the indicated days after infection. Responses in wild-type mice were analyzed for comparison. Each data point represents a measurement on pooled sera from three mice.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We have found that in the mouse, B cells acquire CD27 during the GC reaction and express it there at the highest frequency. CD27 was detected on ~13% of GC B cells in spleen and DLN at the peak of the primary response to influenza virus. In our hands, anti-CD27-biotin conjugates followed by SA-PE or SA-allophycocyanin gave similar percentages of CD27 expression on B cells as use of directly FITC-labeled anti-CD27 mAb. So, within the limits of detection, the frequency of CD27 expression on GC B cells in the mouse appears lower than in humans, where all tonsillar GC B cells reportedly express CD27 (17). We found CD27 on ~25% of centroblasts and only on ~8% of centrocytes, suggesting that it is gradually lost from B cells after their expansion phase. Within GCs, SHM was less frequent among CD27-positive cells than among CD27-negative cells, in line with its predominant expression during the expansion phase. Among memory B cells, only a few percent express CD27. Consistently, CD27+ B cells do not accumulate in the mouse upon aging. However, the frequency of CD27+ B cells transiently increases upon secondary antigenic challenge (data not shown). Our collective data indicate that in the mouse, CD27 is a marker for recent B cell activation and that CD27/CD70 interactions come into play during the B cell expansion phase in GCs. The mutation status of CD27+ and CD27– B cells within GCs is in line with CD27 acquisition preceding or coinciding with the onset of SHM. In humans, CD27 is also induced upon B cell priming, but it appears to be expressed at a higher frequency among GC B cells and maintained long-term on a much higher proportion of post–GC B cells than in the mouse.

Whether CD27 is engaged will depend on presence of CD70, which both in humans and mice is tightly controlled by antigenic stimulation and of a transient nature (1, 3). CD70 is difficult to document on live cells, but by confocal microscopy, we could detect occasional CD70+ B cells in GC and CD70+ T cells at the border of or inside GCs. This suggests that CD27/CD70 interactions come into play during Th cell-B cell communication just before or during B cell expansion in GCs. Studies on human cells show unambiguously that stimulation of CD27 on activated B cells promotes the generation of plasma cells (1). We have indeed revealed such a pathway in vivo, but in the mouse it is not required for production or maintenance of serum IgG and IgA levels in primary or recall responses to influenza virus (this study) or haptenated protein Ags (our unpublished results). In CD27–/– mice, the only observable B cell phenotype is a delay in GC formation, indicative of less effective expansion of activated B cells in the absence of CD27/CD70 interactions. Our adoptive transfer experiments reveal the same phenotype, with a larger window for read out. They add that efficient GC B cell expansion requires CD27 on B cells, while CD27 on T cells is virtually dispensable for it. Most likely, CD27 on B cells is triggered by CD70 on activated CD28+ Th cells. We have recently established that on activated T cells, CD27 promotes survival and not division (8). Therefore, we postulate that CD27 also promotes survival of activated B cells, and thus facilitates their expansion during the GC reaction. In humans, CD27 is expected to fulfil the same role. Additionally, in humans, CD27 may promote the survival of previously primed B cells outside GCs when CD70 is available.

We have also revealed a CD28-independent pathway where CD27+ T cells provide help to B cells to form small GCs and produce IgG. This may involve interaction between CD27 on Th cells and CD70 on B cells. Alternatively, CD27/CD70 interactions between T cells may support expansion of the Th cell population in the absence of CD28. A third scenario is that CD27 on T cells contributes to the B cell response like OX40, in that it would interact with CD70 on CD4+3– accessory cells and stimulate T cells to migrate into B cell follicles (38). Although a contribution of OX40 to the B cell response has been identified by Ab intervention studies (39, 40), GC formation and IgG production in response to viruses and haptens are normal in OX40-deficient mice (41, 42), while only a minor defect was found in OX40 ligand (OX40L)-deficient mice (43). We have found that Ig production in response to influenza virus is normal in OX40L–/– mice, as well as in mice lacking both OX40L and CD27 (Y.X., unpublished observations). Possibly, the GC B cell response is finely tuned by partially overlapping costimulatory signals by TNFR family members and their ligands. The dominant effect of CD28 on GC formation may be explained by its role in the up-regulation of multiple receptors and ligands of this type.

We have found that CD27 deficiency reduces the survival of virus-specific CD8+ T cells in DLN and lung (8). How it impacts on the CD4+ T cell response to influenza virus cannot be followed in this model with the aid of MHC tetramers. Therefore, we have much less knowledge about it. CD27 deficiency does reduce the accumulation of CD4+ T cells in the lung (7). However, GCs are formed in DLN of CD27–/– mice (data not shown). Ongoing work is focused on the CD4 T cell response to OVA, which we can follow both with MHC tetramers and with TCR transgenic cells. These studies should illuminate exactly how CD27 regulates CD4 T cell function.

The detailed division of labor between different TNFR family members and their selective and transient expression on certain cell types during an ongoing immune response will allow for highly specific intervention in disease states. Because constitutive ligation of these receptors can be highly detrimental due to effects on lymphocyte homeostasis (e.g.,44), such interventions should be transient and properly localized.


   Acknowledgments
 
We thank G. Rimmelzwaan, G. Dingjan, R. Hendriks, and J. Laman for assistance and advice, and R. A. W. van Lier, F. Scheeren, and K. Schepers for advice and critical reading of the manuscript. We also thank the personnel of the histology, confocal microscopy, flow cytometry, and experimental animal facilities of the Netherlands Cancer Institute for excellent technical assistance.


   Footnotes
 
1 This work was supported by The Netherlands Organization for Scientific Research and by the Dutch Cancer Society. Back

2 Address correspondence and reprint requests to Dr. Jannie Borst, Division of Immunology, The Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, The Netherlands. E-mail address: j.borst{at}nki.nl Back

3 Abbreviations used in this paper: GC, germinal center; DLN, draining lymph nodes; FDC, follicular dendritic cells; FSC, forward scatter; HPE, high performance ELISA buffer; OX40L, OX40 ligand; PNA, peanut agglutinin; SA, streptavidin; SHM, somatic hypermutation; TR, Texas Red. Back

Received for publication January 5, 2004. Accepted for publication March 26, 2004.


   References
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
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