In humans, up to 40% of peripheral B cells express CD27 and have hypermutated variable regions in their Ig genes. The CD27+ B cells are considered to be derived from germinal center following specific antigenic stimulation. Actually, somatic hypermutation in Ig genes and CD27 expression are hallmarks of memory B cells. However, the blood IgM+IgD+CD27+ B cells were recently associated to splenic marginal zone B cells and proposed to be a subset distinct from germinal center-derived memory B cells showing premutated Igs. The results presented herein further weaken this bona fide association because B cells expressing surface IgG, but not CD27, were found in human blood. Representing 1–4% of all peripheral B cells and ∼25% of the IgG+ blood B cells, this population expressed mutated IgG genes showing antigenic selection characteristics but with lower mutation frequencies than that of CD27+IgG+ B cells. However, their morphology and phenotype were similar to that of CD27+IgG+ cells. Interestingly, the proportion of IgG2 over IgG3 transcripts was opposite in CD27IgG+ and CD27+IgG+ cells, suggesting distinct functions or origins. Overall, these findings extend the memory B cell reservoir beyond the CD27+ compartment and could provide further insights into B cell disorders of unknown etiology.

Memory B cells can be distinguished from naive B cells by the presence of somatic hypermutation in their Ig variable gene sequences (1). Phenotypic characteristics can also be used to identify this B cell subset. Previously, IgD-expressing cells were classified as naive cells and IgD cells as memory B cells (2, 3). However, the description of an IgD+ subset expressing somatically mutated Ig genes discontinued its use for the identification of memory B cells (4, 5). Currently, CD27, belonging to the TNFR family (6), is widely used as a marker of human memory B cells because its expression correlates with the presence of somatic mutations in Ig genes (4, 5, 7, 8). Based on CD27, IgD, IgM, and CD5 expression, peripheral B cells can be subdivided into four CD27+ memory subsets (IgM+IgD+, IgM+IgD, IgG+/IgA+, and IgMIgD+ cells) and two CD27 naive subsets (IgM+IgD+CD5+ and IgM+IgD+CD5), representing 40 and 60% of peripheral B cells, respectively (4).

During immune responses to T-dependent Ags, immunological memory is established mainly via the generation of memory B cells (9, 10). Following an Ag encounter, naive B cells can differentiate into low-affinity Ig-secreting cells or mature within germinal centers into high-affinity memory cells expressing Ig of various isotypes (11). Activated T cell help, mainly by the engagement of B cell-expressed CD40 by T cell-expressed CD154 (12), is necessary for memory cell generation, including germinal center formation, somatic hypermutation, and isotype switching (reviewed in Refs. 11 and 13). The importance of CD40 stimulation in these events has been demonstrated in patients suffering from X-linked hyper IgM syndrome, in which CD40-CD154 interaction is abrogated (reviewed in Ref. 14). In these patients, even though germinal center formation and memory B cell responses are severely impaired, a peripheral B cell fraction expressing CD27 persists (15, 16, 17), suggesting that this marker does not exclusively identify classical germinal center-derived memory B cells. Recently, this CD27+ compartment was recognized as the peripheral counterpart of splenic marginal zone B cells (16, 18). Although expressing mutated Ig genes, this subset differs from classical memory cells, since in this population, hypermutation can occur in absence of CD40-CD154 interactions (16, 17, 18). Rather, as observed in other species, a developmental mechanism increasing repertoire diversity early in life has been postulated (16, 18). Also, unlike classical memory cells, this CD27+ subset is dedicated to immune responses to T-independent Ags (16, 19, 20), during which they likely act as innate effectors in the first line of defense but not as memory cells (16, 21, 22, 23). These findings reinforce the notion that CD27 expression does not necessarily correlate with memory B cells.

Alternatively, the requirement for CD27 expression on classical memory cells has never been firmly established in humans, nor associated with a well-defined function (24). Indeed, we previously reported that prolonged CD40 stimulation triggered naive B cells to switch to IgG and to express CD27 even in absence of somatic hypermutation, suggesting that these latter events could be independent (25). In mice, rather than a memory marker, CD27 appears to be a surrogate for recent B cell activation and is not absolutely necessary for secondary responses (26).

Overall, questions are still raised concerning the exact identification of human memory B cells based on CD27 expression, and the present study further questions this association. Indeed, a new B cell population expressing hypermutated IgG in the absence of CD27 expression was detected in human blood. Representing 1–4% of all peripheral B cells, these cells correspond to ∼25% of IgG+ blood B cells and are morphologically and phenotypically similar to CD27+IgG+ B cells. The identification of this subset extends the human peripheral memory B cell compartment and could suggest novel mechanisms underlying B cell disorders of unknown origin such as hairy cell leukemia (27, 28).

The following Abs conjugated to allophycocyanin, PE, FITC, PE-cyanin 5.1 (PC5), PerCP-Cy5.5, and Alexa Fluor 488 (Alexa-488) were used for flow cytometry analysis or cell sorting: PerCP-Cy5.5- or allophycocyanin-anti-CD19, FITC-, PE-, or PC5-anti-CD27; FITC- or allophycocyanin-anti-IgG, FITC-anti-IgG2, FITC-anti-IgG3, FITC-anti-IgM, allophycocyanin-anti-CD5, allophycocyanin-anti-CD38, PE-anti-IgD, PE-anti-CD126, PE-anti-CD138, Alexa-488-anti-mouse CD45R (B220); and PerCP-Cy5.5-, allophycocyanin-, PE-, PC5-, FITC-, or Alexa-488 isotype controls were used in triple or quadruple staining procedures. Most Abs were mouse monoclonal IgG1 from BD Pharmingen, except PC5 Abs from Beckman Coulter and FITC-anti-IgG2 and anti-IgG3 from Sigma-Aldrich. Alexa-488 Abs were rat monoclonal IgG2a (Caltag Laboratories), and FITC-anti-IgM were polyvalent goat IgG (The Jackson Laboratory).

This study has been reviewed and approved by the Héma-Québec Ethical Committee. Blood samples were collected from healthy individuals after informed consent, and PBMC were isolated by density centrifugation as previously described (25) and stored frozen. B cells were purified by negative selection with CD19 mixture according to the manufacturer’s instructions (Stem Sep; Stem Cell Technologies). Purified B cells were >95% CD19+ cells as determined by flow cytometry analysis. CD19+CD27IgG, CD19+CD27IgG+, and CD19+CD27+IgG+ B cells were separated, using PE-anti-CD27 and FITC-anti-IgG, by cell sorting using a BD FACSCalibur low-speed cell sorter (BD Biosciences) or an Epics Elite ESP (Beckman Coulter). Sorted CD27+IgG+ cells were >95%, CD27+ and CD27IgG+ cells were >95% negative for CD27. IgG2 and IgG3 expression was evaluated in costaining with allophycocyanin-anti-IgG, which recognized all human IgG subclasses. No steric hindrance was observed in these costaining procedures since IgG2- and IgG3-positive cells were both stained by the allophycocyanin-anti-IgG. ATP-binding cassette transporter activity (ABC B1)3 was evaluated as described elsewhere (29), using MitoTracker Green (MTG) (Invitrogen Life Technologies). Briefly, PBMC were incubated at 37°C in IMDM containing 10% FBS and 100 nM MTG for 30 min and in absence of MTG for 40 min and then analyzed for CD19, CD27, and IgG expression. Flow cytometry analysis and staining were done as previously described (25), using FACSCalibur flow cytometer with CellQuest Pro software (BD Biosciences). All analyses with PBMC were done on ≥150,000 events. Data were subsequently analyzed with a computer program (FCS express II; De Novo Software).

Sorted CD19+CD27IgG+ B cells, CD19+CD27+IgG+ cells, and CD19+CD27IgG cells were spun onto glass slides, stained with Wright-Giemsa stain (Sigma-Aldrich), and examined by light microscopy using a Nikon Eclipse TE 2000-S microscope (Nikon) at ×400 magnification. Digital pictures were acquired using a Retiga 1300 camera (QImaging) and the SimplePCI 5 Software (Compix).

Total RNA was extracted from 6 to 8 × 104 sorted CD19+CD27+IgG+ and CD19+CD27IgG+ B cells using TRIzol reagent according to the manufacturer’s instructions (Invitrogen Life Technologies). The total amount of RNA obtained for each cell sample was used to synthesize cDNAs using oligo(dT) primers and Moloney murine leukemia virus reverse transcriptase (Invitrogen Life Technologies). Reverse transcriptase reaction done with 500 ng of total RNA from CD40-activated B cells (30) was used as positive control. VH and γ 1, 2, 3, and 4 regions of IgG and β-actin were amplified by PCR. Amplification of VH regions was done using 35 cycles of 94°C for 30s, 45°C for 40 s, and 72°C for 60s. Amplification of γ regions was done as above, except that step 2 was 60 s at 65°C for γ 1, 62.5°C for γ 2, 3, and 4, and 55°C for β-actin. One to 2 μl of reverse transcriptase mixture was used with Platinum TaqDNA polymerase (Invitrogen Life Technologies). The primers used for VH were 5′ IgG-specific Cγ (5′-AAGTAGTCCTTGACC) and 3′ VH1-, VH2-, VH3-, and VH4-specific primers described elsewhere (31), and for IgG1, IgG2, and IgG3 amplification, the same 5′ primer was used (5′-TCCACCAAGGGCCCATCG-3′) in combination with the following 3′ primers: CG1z (5′-GCATGTACTAGTTTTGTCACAAGATTTGGG-3′) for IgG1, CG2a (5′-CTCGACACTAGTTTTGCGCTCAACTGTCTT-3′) for IgG2, and CG3a (5′-TGTGTGACTAGTGTCACCAAGTGGGGTTTT-3′) for IgG3 (32, 33). IgG4 amplification was done using 5′ primer (5′-GCTTCCACCAAGGGCCCATC-3′) and in 3′ primer CG4a (5′-GCATGAACTAGTTGGGGGACCATATTTGGA-3′) (32). β-actin primers were described elsewhere (25). Amplicon specificity was confirmed on 1% agarose gel electrophoresis using ethidium bromide staining. Densitometry analysis was done using the Alpha EaseFC software from Alpha Innotech, and relative intensity of gene expression was based on the ratio of γ 1, 2, 3, or 4 to β-actin gene expression. VH amplicons (500–650 nt) were purified from 1% agarose gel using the QIAquick Gel extraction kit (Qiagen) and cloned into pDRIVE vector from the Qiagen PCR cloning kit (Qiagen). Plasmid DNA was purified using conventional techniques (34), and the cloned IgG VH regions were sequenced in both directions using M13 primers by automated fluorescent DNA sequencing (ABI373; Applied Biosystems). Taq polymerase error was estimated at 0.34%, representing one mutation every 294 nt (25).

Mutational analysis of VH IgG transcripts was done using the ImMunoGeneTics (IMGT) database (available at 〈http://imgt.cines.fr〉) (35). The number of mutated nucleotides was determined for each transcript after their alignment with the germline gene showing the highest homology from framework region (FR) 1 to FR3 for a total of 312 nt. The distribution of replacement (R) and silent (S) mutations and the R:S ratios were evaluated at the amino acid level for each gene region and were used as an index of antigenic selection (36, 37, 38, 39). The distribution of R and S mutations within FR1, FR2, and FR3 and CDR1 and CDR2 was evaluated in each region, comprising 26 aa for FR1, 17 aa for FR2, 39 aa for FR3, 12 aa for CDR1, and 10 aa for CDR2. The classification of each transcript to IgG1, IgG2, IgG3, or IgG4 was done using the applet “Blast 2 sequences” from the National Center for Biotechnology Information web site (〈www.ncbi.nlm.nih.gov/blast/bl2seq/wblast2.cgi〉). The 100-nt constant region amplified from the CH1 region of each transcript was sequentially aligned against sequences representing IgG constant region of each subclass (GenBank accession no. J00228 for IgG1, J00230 for IgG2, D78345, and AF237585 for IgG3, and K01316 for IgG4). The single alignment giving the highest homology (99–100%) was used for identification of subclasses.

Statistical significance was evaluated using two-tailed unpaired Student’s t tests or two-tailed Fisher exact tests, using a value of p < 0.05. All results are presented as mean ± SD.

Currently, CD27 expression is associated with memory B cells since all CD27+ B cells express hypermutated Ig genes (4, 5, 7, 8) and peripheral B cells expressing isotype switched Ig are usually considered to express CD27 (4, 5). We have detected, in human blood, a distinct B cell population, which did not correspond to these criteria. Indeed, CD19+CD27IgG+ B cells were detected in PBMC prepared from 15 independent samples (Table I and Fig. 1,A). As expected (5), an average of 9.7 ± 4.3% IgG-expressing cells was observed among these blood samples, but IgG+ cells were further separated into 7.3 ± 3.3% CD27+IgG+ cells and 2.4 ± 1.2% CD27IgG+ cells. The CD19+CD27IgG+ population represented ∼25% of IgG-expressing cells in blood (25.4 ± 5.2%). These results were corroborated using purified CD19+ B cells (Fig. 1,B), whereas CD19+IgG+ cells were gated and analyzed for CD27 expression. Three independent samples were showing an average of 28 ± 7% of CD27 cells among IgG-expressing cells (data not shown). This new population within CD19+IgG+ cells resulted in ∼75% of the cells coexpressing CD27 instead of the expected 100%. Furthermore, no intracellular CD27 expression wasdetected in these CD27IgG+ cells (data not shown). When IgG expression was evaluated within CD19+CD27+ and CD19+CD27 populations, 24.3 ± 8% of CD27+ cells expressed IgG whereas 3.4 ± 2% of CD27 cells were IgG+ (Fig. 1 C and data not shown). Finally, the mean fluorescence intensity (MFI) of IgG expression was similar in CD27+IgG+ and CD27IgG+ cells, being of 186 ± 4 and 191 ± 17, respectively (data not shown).

Table I.

CD19+CD27IgG+ B cells are present in peripheral blood

SampleIgG+ (%)aCD27+IgG+ (%)aCD27IgG+ (%)aCD27IgG+/IgG+ (%)b
6.1 4.8 1.4 22.1 
3.7 2.5 1.2 32.8 
14.8 10.6 4.1 28.0 
6.9 5.4 1.5 21.3 
16.5 13.9 2.6 16.0 
11.8 9.0 2.8 23.9 
6.4 4.9 1.6 24.3 
12.4 9.4 2.9 23.6 
11.6 8.3 3.2 27.8 
10 5.3 4.1 1.3 23.6 
11 6.6 4.8 1.8 27.4 
12 11.8 7.8 4.0 34.0 
13 9.7 8.0 1.7 17.4 
14 16.8 12.2 4.7 27.6 
15 5.3 3.7 1.7 31.6 
Mean ± SD 9.7 ± 4.3 7.3 ± 3.3 2.4 ± 1.2 25.4 ± 5.2 
SampleIgG+ (%)aCD27+IgG+ (%)aCD27IgG+ (%)aCD27IgG+/IgG+ (%)b
6.1 4.8 1.4 22.1 
3.7 2.5 1.2 32.8 
14.8 10.6 4.1 28.0 
6.9 5.4 1.5 21.3 
16.5 13.9 2.6 16.0 
11.8 9.0 2.8 23.9 
6.4 4.9 1.6 24.3 
12.4 9.4 2.9 23.6 
11.6 8.3 3.2 27.8 
10 5.3 4.1 1.3 23.6 
11 6.6 4.8 1.8 27.4 
12 11.8 7.8 4.0 34.0 
13 9.7 8.0 1.7 17.4 
14 16.8 12.2 4.7 27.6 
15 5.3 3.7 1.7 31.6 
Mean ± SD 9.7 ± 4.3 7.3 ± 3.3 2.4 ± 1.2 25.4 ± 5.2 
a

PBMC were analyzed for CD19, CD27, and IgG expression, and CD19+ cells were gated and used to determine the proportions of IgG+ and, as shown in Fig. 1 A, CD27+IgG+ and CD27IgG+ cells (R1 and R2 regions, respectively).

b

The percentage of CD19+CD27IgG+ among total CD19+IgG+cells.

FIGURE 1.

Surface IgG expression is not restricted to CD27+ B cells in human blood. CD19, CD27, and IgG expression was evaluated by flow cytometry in PBMC and purified blood CD19+ B cells using triple staining. A, IgG and CD27 expression was evaluated in gated CD19+ cells corresponding to 15% of total PBMC. R1 and R2 regions were used to delineate CD27IgG+ and CD27+IgG+ cells, respectively. These results are representative of 15 independent samples (Table I). B, CD27 expression was evaluated in gated IgG+ cells (∼100,000 events, R1). C, IgG expression was evaluated in CD19+CD27+ (∼50,000 events, R4) and CD19+CD27 (∼50,000 events, R5) cells using purified CD19+ B cells (≥98%). Results presented in B and C are representative of three independent samples.

FIGURE 1.

Surface IgG expression is not restricted to CD27+ B cells in human blood. CD19, CD27, and IgG expression was evaluated by flow cytometry in PBMC and purified blood CD19+ B cells using triple staining. A, IgG and CD27 expression was evaluated in gated CD19+ cells corresponding to 15% of total PBMC. R1 and R2 regions were used to delineate CD27IgG+ and CD27+IgG+ cells, respectively. These results are representative of 15 independent samples (Table I). B, CD27 expression was evaluated in gated IgG+ cells (∼100,000 events, R1). C, IgG expression was evaluated in CD19+CD27+ (∼50,000 events, R4) and CD19+CD27 (∼50,000 events, R5) cells using purified CD19+ B cells (≥98%). Results presented in B and C are representative of three independent samples.

Close modal

To further investigate whether these CD19+CD27IgG+ B cells corresponded to postgerminal center cells or not, the frequency of somatic mutations in their IgG transcripts was evaluated using sorted populations of CD19+CD27IgG+ and CD19+CD27+IgG+ B cells. A total of 51 and 26 IgG transcripts was analyzed for CD19+CD27IgG+ and CD19+CD27+IgG+ B cells, respectively (Table II). Two independent samples were used for each targeted population, CD27 and CD27+IgG+ cells, and showed no difference in their level of mutations (p < 0.01; Student’s t test). CD27+IgG+ and CD27IgG+ cells presented mutation levels significantly different: 22 ± 8 and 11 ± 9 mutated nucleotides per gene, respectively (p < 0,001; Student’s t test, data not shown). All transcripts from CD27+IgG+ B cells showed somatic mutations, with an average frequency of 6.9 ± 2.5%, which is in accordance with our previous observations (25) and the reported 4–12% mutation frequency in peripheral memory B cells (40, 41). Even though CD27IgG+ subset also showed up to 10.6% mutation frequency, a fraction of these transcripts, 13.7% (7 over 51), showed none or only 1 mutated nucleotide per gene. Excluding these seven unmutated transcripts, the average mutation frequency, 4.1 ± 2.6% instead of 3.6 ± 2.8% for all transcripts, was lower than that of CD27+IgG+ cells. The distribution of mutation frequency in these two subsets further highlighted this difference (Fig. 2 A). The proportion of transcripts showing 0–10 mutations was significantly higher in CD27IgG+ cells (57%) than in CD27+IgG+ cells (7%). Conversely, the proportion of transcripts showing 21–30 mutations was significantly lower in CD27IgG+ cells (8%) than in CD27+IgG+ cells (42%).

Table II.

CD19+CD27IgG+ blood B cells express hypermutated IgG

CD19+ CellsaSampleNo. of Transcripts (total nucleotides analyzed)No. of Mutated Nucleotides (total mutated nucleotides)Percentage of Mutations (total percentage of mutated nucleotide)
CD27+IgG+ 10–36 3.2–11.5 
  (2,808) (206) (7.3) 
 17 7–33 2.2–10.6 
  (5,304) (357) (6.7) 
 Total 26 7–36 2.2–11.5 
  (8,112) (563) (6.9 ± 2.5) 
CD27IgG+ 20 0–33 0–10.6 
  (6,240) (223) (3.6) 
 31 0–33 0–10.6 
  (9,672) (344) (3.6) 
 Total 51 0–33 0–10.6 
  (15,912) (567) (3.6 ± 2.8) 
CD19+ CellsaSampleNo. of Transcripts (total nucleotides analyzed)No. of Mutated Nucleotides (total mutated nucleotides)Percentage of Mutations (total percentage of mutated nucleotide)
CD27+IgG+ 10–36 3.2–11.5 
  (2,808) (206) (7.3) 
 17 7–33 2.2–10.6 
  (5,304) (357) (6.7) 
 Total 26 7–36 2.2–11.5 
  (8,112) (563) (6.9 ± 2.5) 
CD27IgG+ 20 0–33 0–10.6 
  (6,240) (223) (3.6) 
 31 0–33 0–10.6 
  (9,672) (344) (3.6) 
 Total 51 0–33 0–10.6 
  (15,912) (567) (3.6 ± 2.8) 
a

Total RNA was prepared, as described in Materials and Methods, from sorted CD19+CD27IgG+ (>95% pure) and CD19+CD27+IgG+ (>95% pure) B cells, respectively; 75,000 and 65,000 cells for sample 1 and 65,000 cells for each subset in sample 2.

FIGURE 2.

CD27IgG+ B cells present a lower mutation load than CD27+IgG+ memory cells with an imprint of Ag selection. A, All IgG transcripts (Table II) were sequenced and classified according to the number of mutated nucleotides per gene for each population. ∗ and ∗∗, Significant difference between the two populations, with p = 0.00005 and p = 0.0006, respectively, using two-tailed Fisher exact tests. B, The mean frequency of S and R mutations, identified in amino acid sequences of FR1, FR2, and FR3, and CDR1 and CDR2, is presented. Each region was delimited according to IMGT standard. Two-tailed unpaired Student’s t tests were used to compare mutation frequency from CDR and FR regions. ∗, ∗∗, and ∗∗∗, Significant difference between CDR1 and FR1 (p < 0.01), CDR2 and FR1, FR2, or FR3 (p < 0.001), and CDR1 and FR1 or FR2 (p < 0.001), respectively.

FIGURE 2.

CD27IgG+ B cells present a lower mutation load than CD27+IgG+ memory cells with an imprint of Ag selection. A, All IgG transcripts (Table II) were sequenced and classified according to the number of mutated nucleotides per gene for each population. ∗ and ∗∗, Significant difference between the two populations, with p = 0.00005 and p = 0.0006, respectively, using two-tailed Fisher exact tests. B, The mean frequency of S and R mutations, identified in amino acid sequences of FR1, FR2, and FR3, and CDR1 and CDR2, is presented. Each region was delimited according to IMGT standard. Two-tailed unpaired Student’s t tests were used to compare mutation frequency from CDR and FR regions. ∗, ∗∗, and ∗∗∗, Significant difference between CDR1 and FR1 (p < 0.01), CDR2 and FR1, FR2, or FR3 (p < 0.001), and CDR1 and FR1 or FR2 (p < 0.001), respectively.

Close modal

Because Ag-selected Ig sequences tend to have a higher frequency of mutations leading to the replacement of amino acids in the CDR, the distribution of replacement vs silent mutations among the FR1, FR2, and FR3 and CDR1 and CDR2 regions of IgG variable regions was compared in both subsets (Fig. 2,B). In both populations, the mean frequency of S and R mutations found in FR1, FR2, and FR3 and in CDR1 and CDR2 was similar (Fig. 2 B). As expected, the frequency of R mutations was higher in CDR than FR. The frequency of R mutations in CDR2 was significantly higher than that of FR1, FR2, and FR3 in both subsets (p < 0.001; Student’s t test). The R:S ratios for CD27+IgG+ and CD27IgG+ cells were respectively, 0.8 and 1.8 in FR1, 2.6 and 4.5 in CDR1, 1.9 for both in FR2, 3.9 and 4.5 in CDR2, and 2.0 and 1.8 in FR3. Higher R:S ratios in CDR compared with FR were considered as an indicator of Ag selection, as expected for postgerminal center cells (36, 37, 38, 39). Overall, CD27IgG+ cells conducted lower mutation frequency in their transcripts but showed antigenic selection characteristics similar to that of CD27+IgG+ cells.

Flow cytometry analysis was used to further characterize the blood CD19+CD27IgG+ cells in comparison to CD19+CD27+IgG+ cells. Expression of CD5, CD38, CD40, CD70, CD126, CD138, and B220 was monitored among the B cell populations delineated with CD19, CD27, and IgG using quadruple staining. Analyses were done using PBMC prepared from samples obtained from healthy individuals (Table III). The proportion of CD19+ B cells expressing CD5, CD38, CD40, CD70, CD126, CD138, and B220 corroborated previous reports (42, 43, 44, 45, 46). Both CD19+CD27IgG+ and CD19+CD27+IgG+ cells expressed high levels of CD40 while ∼25% of each subset expressed similar level of CD38. The frequency of CD38- and CD40-positive cells was corroborated using purified CD19+ B cells (data not shown). In addition, the marginal zone B cell marker CD1c (23) was not detected in those two populations (data not shown). Although we cannot exclude the possibility of a rapid CD27 modulation or transient CD27 expression (47), CD27 protein was not detected in CD27IgG+ cells using intracellular staining for flow cytometry analysis (data not shown). As for the CD27+ counterpart, negligible expression of CD5 indicates that the CD19+CD27IgG+ cells do not belong to B1a cell subset (48).

Table III.

CD19+CD27IgG+ and CD19+CD27+IgG+ blood B cells show comparable phenotype

Markers (no. of samples)Percentage of Positive Cells (±SD) (MFI ± SD)a
CD19+CD19+CD27+IgG+CD19+CD27IgG+
CD5 24.0 ± 10 6.7 ± 1.4 5.7 ± 2.5 
(9)b (62 ± 16) (46 ± 14) (33 ± 11) 
CD38 46.2 ± 14.3 23.4 ± 8.3 32.0 ± 11.0 
(10) (39 ± 10) (39 ± 17) (46 ± 18) 
CD40 99.0 ± 0.5 99.2 ± 0.8 99.9 ± 0.1 
(5) (287 ± 61) (348 ± 71) (330 ± 81) 
CD70 1.0 ± 0.5 6.6 ± 3.6 6.1 ± 3.5 
(9) (64 ± 17) (59 ± 27) (43 ± 10) 
CD126 4.3 ± 1.2 39.8 ± 17.0 21.4 ± 11.9 
(9) (29 ± 3) (27 ± 3) (23 ± 3) 
CD138 1.8 ± 1.9 8.6 ± 5.0 9.8 ± 6.8 
(9) (51 ± 17) (38 ± 9) (33 ± 13) 
B220 57.5 ± 15.6 24.1 ± 14.4 50.3 ± 22.7 
(5) (37 ± 8) (39 ± 3) (32 ± 2) 
Markers (no. of samples)Percentage of Positive Cells (±SD) (MFI ± SD)a
CD19+CD19+CD27+IgG+CD19+CD27IgG+
CD5 24.0 ± 10 6.7 ± 1.4 5.7 ± 2.5 
(9)b (62 ± 16) (46 ± 14) (33 ± 11) 
CD38 46.2 ± 14.3 23.4 ± 8.3 32.0 ± 11.0 
(10) (39 ± 10) (39 ± 17) (46 ± 18) 
CD40 99.0 ± 0.5 99.2 ± 0.8 99.9 ± 0.1 
(5) (287 ± 61) (348 ± 71) (330 ± 81) 
CD70 1.0 ± 0.5 6.6 ± 3.6 6.1 ± 3.5 
(9) (64 ± 17) (59 ± 27) (43 ± 10) 
CD126 4.3 ± 1.2 39.8 ± 17.0 21.4 ± 11.9 
(9) (29 ± 3) (27 ± 3) (23 ± 3) 
CD138 1.8 ± 1.9 8.6 ± 5.0 9.8 ± 6.8 
(9) (51 ± 17) (38 ± 9) (33 ± 13) 
B220 57.5 ± 15.6 24.1 ± 14.4 50.3 ± 22.7 
(5) (37 ± 8) (39 ± 3) (32 ± 2) 
a

The proportion of positive cells and MFI for each marker are presented as a mean ± SD.

b

Total CD19+, CD19+CD27IgG+, and CD19+CD27+IgG+ B lymphocytes were gated within PBMC using 150,000–200,000 events as shown in Fig. 1 A, and expression of CD5, CD38, CD40, CD70, CD126, CD138, and B220 was evaluated in each subset. Anayses were done on 5, 9, or 10 independent samples, as indicated.

To confirm that the CD27IgG+ B cells were not CD27IgD+IgM+ naive B cells that have bound IgG via FcγR (49, 50, 51), IgD and IgM expression was verified on purified B cells using quadruple staining (Fig. 3 A). Both CD19+CD27IgG+ and CD19+CD27+IgG+ cells were negative at ≥95% for IgD and IgM expression.

FIGURE 3.

Blood CD27IgG+ and CD27+IgG+ B cells are similarly negative for IgD, IgM, and CD138 expression while differently expressing B220. In each assay, a total of 150,000–200,000 events was analyzed by flow cytometry. A, Purified blood CD19+ B cells were used to delineate CD19+CD27IgG+ and CD19+CD27+IgG+ cells, as done in Fig. 1,C, and IgD and IgM expression was evaluated using quadruple staining. These results are representative of three independent samples. B, PBMC were used to delineate CD19+CD27IgG+ and CD19+CD27+IgG+ cells, as done in Fig. 1,A, and CD138 and B220 expression was evaluated using quadruple staining. These profiles are representative of nine and five independent samples for CD138 and B220 analysis, respectively (Table III).

FIGURE 3.

Blood CD27IgG+ and CD27+IgG+ B cells are similarly negative for IgD, IgM, and CD138 expression while differently expressing B220. In each assay, a total of 150,000–200,000 events was analyzed by flow cytometry. A, Purified blood CD19+ B cells were used to delineate CD19+CD27IgG+ and CD19+CD27+IgG+ cells, as done in Fig. 1,C, and IgD and IgM expression was evaluated using quadruple staining. These results are representative of three independent samples. B, PBMC were used to delineate CD19+CD27IgG+ and CD19+CD27+IgG+ cells, as done in Fig. 1,A, and CD138 and B220 expression was evaluated using quadruple staining. These profiles are representative of nine and five independent samples for CD138 and B220 analysis, respectively (Table III).

Close modal

A fraction of peripheral CD19+CD27+IgG+ and CD19+CD27IgG+ cells, 8.6 ± 5.0 and 9.8 ± 6.8%, respectively, expressed CD138 (Table III). Taken together, these cells represented <1% of the blood CD19+ cells and expressed a low level of CD138 (MFI < 40; Fig. 3 B), in comparison to that expected for terminally differentiated blood plasma cells (MFI > 300) (52). These CD138low cells may thus represent IgG+ cells that are in transition toward plasma cell differentiation.

CD27 naive B cells are considered to express B220 (>95%), whereas ∼40 ± 10% of isotype-switched CD27+ cells can be B220+ (42, 53). We observed that 24 ± 14% of CD19+CD27+IgG+ and 50 ± 23% of CD19+CD27IgG+ cells expressed B220 (Table III and Fig. 3 B). However, this 2-fold difference in B220 expression was not statistically significant (p < 0.05, Student’s t test), thus reinforcing the similarity between IgG+CD27 and bona fide IgG+CD27+ memory cells.

Significant differences (p < 0.05, Student’s t test) were observed only for the CD126 expression, the α-subunit of IL-6R (54). Two-fold less CD19+CD27IgG+ cells were expressing CD126 compared with CD19+CD27+IgG+ cells. These results were confirmed using purified B cells prepared from five independent samples (Fig. 4 A). This lower CD126 expression suggests that IL-6 stimulation could be distinct in these populations (55).

FIGURE 4.

Expression of CD126 is higher in CD27+IgG+ compared with CD27IgG+ B cells but both populations are negative for ABC B1 transporter. A, CD126 expression was evaluated using quadruple staining in CD27+, CD27, CD27IgG+, and CD27+IgG+ populations using purified B cells (>98% CD19+). B, After MTG incubation, PBMC were stained for CD19, CD27, MTG, and IgG expression, and CD19+IgG+ cells were gated (R3) and analyzed for CD27 expression and MTG content. Gated events (200,000) were analyzed in A and B. Representative profiles of four and three independent samples are shown for A and B, respectively.

FIGURE 4.

Expression of CD126 is higher in CD27+IgG+ compared with CD27IgG+ B cells but both populations are negative for ABC B1 transporter. A, CD126 expression was evaluated using quadruple staining in CD27+, CD27, CD27IgG+, and CD27+IgG+ populations using purified B cells (>98% CD19+). B, After MTG incubation, PBMC were stained for CD19, CD27, MTG, and IgG expression, and CD19+IgG+ cells were gated (R3) and analyzed for CD27 expression and MTG content. Gated events (200,000) were analyzed in A and B. Representative profiles of four and three independent samples are shown for A and B, respectively.

Close modal

Additionally, based on a recent report (29), we also observed that both CD27+IgG+ and CD27IgG+ cells were negative for ABC transporter activity (Fig. 4 B). As reported, ∼75% of CD27 cells were able to expel MTG (MFI 156) while >90% of CD27+ cells were not (MFI 1230). This new physiological marker is considered to delineate more accurately naive vs memory B cells (29) and therefore indicates that CD27IgG+ cells are memory B cells.

Finally, we investigated the morphology of sorted CD19+CD27IgG+, CD19+CD27+IgG+, and CD19+CD27IgG B cells using Wright-Giemsa staining (Fig. 5). Memory B cells are expected to be larger than CD27 naive cells, with more abundant cytoplasm (7, 56, 57). Indeed, the CD19+CD27IgG+ cells showed morphology similar to that of CD27+ memory B cells, according to cellular size and cytoplasm abundance. Overall, phenotype and morphology of CD19+CD27IgG+ B cells indicate that these cells belong to the memory compartment.

FIGURE 5.

CD27IgG+ and CD27+IgG+ B cells are morphologically similar. Blood B cells were sorted according to CD27 and IgG expression and stained using Wright-Giemsa stain. A, CD19+CD27IgG naive cells, CD19+CD27+IgG+ memory cells (B), and CD19+CD27IgG+ cells (C) are presented, next to a 25-μm bar reference (magnification, ×400). These results are representative of two independent samples. Cell purity was >95% in all samples.

FIGURE 5.

CD27IgG+ and CD27+IgG+ B cells are morphologically similar. Blood B cells were sorted according to CD27 and IgG expression and stained using Wright-Giemsa stain. A, CD19+CD27IgG naive cells, CD19+CD27+IgG+ memory cells (B), and CD19+CD27IgG+ cells (C) are presented, next to a 25-μm bar reference (magnification, ×400). These results are representative of two independent samples. Cell purity was >95% in all samples.

Close modal

To further compare CD19+CD27IgG+ and CD19+CD27+IgG+ B cell populations, we have analyzed the distribution of IgG1, IgG2, IgG3, and IgG4 mRNA in two independent samples (Fig. 6). RT-PCR amplification was done on sorted cells from two samples using β-actin as control (Fig. 6,A). Both CD19+CD27IgG+ and CD19+CD27+IgG+ B cells expressed all IgG subclasses (Fig. 6,B). Densitometry was used to estimate the relative expression level of each IgG subclass. The relative expression levels of IgG1 and IgG4 were similar in both populations, whereas IgG2 gene expression was ∼4-fold higher in CD27+IgG+ B cells, and IgG3 gene expression was ∼10-fold higher in CD27IgG+ B cells. The relative expressions of IgG1, IgG2, and IgG3 in these two populations were corroborated by the analysis of the cloned transcripts presented in Table II and in Fig. 2 (Fig. 6 C). No IgG4 transcripts have been cloned, probably due to their low expression level (58), but a comparable fraction of IgG1 transcripts was observed in both populations. A significant predominance of IgG2 transcripts was observed in CD27+IgG+ over CD27IgG+ B cells, representing 30.8 vs 3.9%, respectively (p = 0.002, Fisher exact test). IgG3 transcripts were predominant in CD27IgG+ over CD27+IgG+ B cells, representing, respectively, 31.4 vs 7.7% (p = 0.02, Fisher exact test). Finally, the distribution of cells positive for surface expression of IgG2 and IgG3 was reverse in CD27IgG+ vs CD27+IgG+ when analyzing purified B cells by flow cytometry. Indeed, a proportion of 28.4 ± 10.7% of the CD27IgG+ cells expressed IgG3 compared with 16.4 ± 5.8% within the CD27+IgG+ cells (n = 6; p < 0.05, Student’s t test) (data not shown). Conversely, 12.3 ± 1.2% of CD27+IgG+ cells expressed IgG2 compared with 2.8 ± 1.0% within the CD27IgG+ cells (n = 3; p < 0.05, Student’s t test) (data not shown). In both subsets, the proportion of IgG3+ cells was higher than IgG2+ cells. Overall, the significant difference in IgG subclass distribution, in accordance to a recent report (29), suggests that CD27+IgG+ and CD27IgG+ B cells could play distinct roles during secondary immune responses.

FIGURE 6.

IgG2 and IgG3 are differentially expressed in CD27IgG+ and CD27+IgG+ cells. A, IgG1, IgG2, IgG3, IgG4, and β-actin mRNA were amplified by RT-PCR using total RNA from sorted CD27+IgG+ and CD27IgG+ B cells, prepared from two independent samples (1 and 2), and from CD40-activated B cells as control. Amplicons of 327, 312, 327, 312, and 540 bp were obtained for IgG1, IgG2, IgG3, IgG4, and β-actin, respectively. One kilobase Plus DNA ladder m.w. markers were used in lanes 1 and 7. B, Relative intensity of gene expression was determined by densitometry for γ 1, 2, 3, and 4 using β-actin as a reference, as described in Materials and Methods. C, Subclasses of each IgG transcripts from CD27IgG+ and CD27+IgG+ cells, respectively, 51 and 26 (Table II), were determined as described in Materials and Methods. ∗ and ∗∗, Significant difference between paired observations, with p = 0.002 and p = 0.02, respectively, using two-tailed Fisher exact tests.

FIGURE 6.

IgG2 and IgG3 are differentially expressed in CD27IgG+ and CD27+IgG+ cells. A, IgG1, IgG2, IgG3, IgG4, and β-actin mRNA were amplified by RT-PCR using total RNA from sorted CD27+IgG+ and CD27IgG+ B cells, prepared from two independent samples (1 and 2), and from CD40-activated B cells as control. Amplicons of 327, 312, 327, 312, and 540 bp were obtained for IgG1, IgG2, IgG3, IgG4, and β-actin, respectively. One kilobase Plus DNA ladder m.w. markers were used in lanes 1 and 7. B, Relative intensity of gene expression was determined by densitometry for γ 1, 2, 3, and 4 using β-actin as a reference, as described in Materials and Methods. C, Subclasses of each IgG transcripts from CD27IgG+ and CD27+IgG+ cells, respectively, 51 and 26 (Table II), were determined as described in Materials and Methods. ∗ and ∗∗, Significant difference between paired observations, with p = 0.002 and p = 0.02, respectively, using two-tailed Fisher exact tests.

Close modal

Until recently, the presence of CD27IgG+ B cells showing mutated Ig genes was supported only by observations done within spleen (59, 60) and tonsils (7, 50, 61). For instance in humans, CD27IgG+ B cells are found in the spleen (59, 60), interfollicular CD27 B cells with somatically mutated Ig genes are detected in lymph nodes (61), and CD27IgG+ B cells expressing mutated genes and the FcR homolog 4 (FcRH4) are reported in tonsils (50). More recently, a minor population of IgG+ B cells, representing ∼4% of CD27 B cells, was observed in human blood (29, 62). We observed a similar frequency of CD27IgG+ cells within the peripheral B cells (1–4%) and the CD19+CD27 cells (∼5%) (29, 62). The fact that these CD27IgG+ cells were not reported before by many exhaustive studies on human B cell repertoire could be related to their low frequency in PBMC or to the use of IgD to delineate B cell populations (4, 15, 56). Besides, even though the data are not shown in the two recent studies (29, 62), the frequency of CD27IgG+ cells over IgG+ blood cells can be estimated to a range of 10–16%. We observed that 16–33% of IgG+ cells were CD27 in 15 samples of PBMC (Table I) and also in 6 purified B cell samples (Figs. 1 and 4) following the analysis of ≥150,000 events in both cases. This variation between their observations and ours could result from the parameters used to analyze the IgG+ cells or from the methods used to purify B cells, which was negative selection in our case and positive selection of CD19+ cells in the two others. Despite that, we have all consistently observed a significant population of CD27IgG+ B cells in human blood. Moreover, we corroborated that the CD27IgG+ B cells were negative for ABC transporter activity and showed a bias for IgG3 expression at both mRNA and protein levels, as reported by Wirths and Lanzavecchia (29).

The FcRH4+CD27 B cells represent a distinctive tissue-based memory population that is rarely observed in blood (50). These cells are mainly IgG+ and show a mutation frequency of 4.5 ± 3.3% in their VH3 Ig gene regions. Similarities between these FcRH4+CD27 cells and the CD27IgG+ cells described herein suggest a possible relation between these two and thus justify further investigations.

Furthermore, in reference to the aforementioned studies (29, 50, 62), we have also investigated in more detail the phenotype, morphology and VH repertoire of this blood CD27IgG+ human B cell population. Overall, the CD27IgG+ cells, expressing mutated Ig and showing imprints of antigenic selection, were morphologically and phenotypically similar to the blood CD27+IgG+ memory cells, which originated from germinal center (11, 63). However, differences were observed in mutation frequency, IgG subclasses distribution, proportion of CD126+ cells and, obviously, CD27 expression, suggesting a distinct origin or role for these two populations during immune responses.

First, the mutation frequency in IgG VH genes expressed by the peripheral CD27IgG+ cells when only mutated transcripts are analyzed (4.1 ± 2.6%) appears closer to that of CD27+IgD+IgM+ memory B cells from periphery (2–5%) (1, 16, 18, 41, 57) or splenic marginal zone (2–4%) (16, 64, 65) than to germinal center IgG+ cells (4–12.6%) (40, 41, 61). However, marginal zone B cells express CD27 and CD1c (23), which are not expressed by the CD27IgG+ cells. On the other hand, many studies have investigated the gene expression profiles of germinal center B cell subsets (66, 67, 68, 69). In mice, CD27 is not considered as a memory B cell marker (11) being expressed for short period of time on a limited proportion of cells only (69). Rather, CD27 appears following recent B cell activation and is not absolutely necessary for secondary responses (26). In humans, the requirement for CD27 expression on germinal center memory cells is not yet well characterized, mainly because the genes regulated by its stimulation remain to be defined and because no human disease related to CD27 dysfunction has been reported (24, 69). However, in vitro studies suggested that hypermutation, isotype switching, and CD27 expression can occur independently in germinal center (25, 70). Therefore, blood CD27IgG+ B cells could reasonably be related to germinal center because of their switched Ig showing mutated VH genes with antigenic selection characteristics.

Approximately 14% of the IgG+CD27 B cells showed unmutated IgG VH gene (7 over 51 transcripts) with a distribution of IgG1 (3 of 7), IgG2 (1 of 7), and IgG3 (3 of 7) similar to that observed for the mutated IgG+CD27 B cells (data not shown and Fig. 6). Interestingly, peripheral IgD+IgM+CD27, as well as IgD+IgM+CD27+ cells driven to switch in vitro, showed a similar pattern of IgG subclass expression, with higher proportion of IgG3 over IgG2 (71). Such switched human B cells carrying unmutated VH genes could originate from a rapid T cell-independent response as reported elsewhere (72, 73, 74). Additionally, the low frequency of somatic mutation in the remaining CD27IgG+ cells suggests that this population could emerge independently from T cell help or from CD40-CD154 interaction in humans (17). In fact, the recent finding that IgG+ memory B cells can be generated following T cell independent responses in mice (75) suggests the existence of such cells in humans.

In addition, regardless of CD27 expression, the low frequency of somatic mutation in CD27IgG+ cells suggests that these cells could represent a first wave of memory B cells as recently proposed (76, 77). Based on mouse models, these studies report the rapid emergence of isotype-switched B cells early after primary immunization. Interestingly, the population of blood (4-hydroxy-3-nitrophenyl)acetyl specific B cells expressing VH genes with low mutation frequency seems tightly regulated (76), suggesting that the CD27IgG+ human blood B cells could represent, as proposed elsewhere (78), a pool of short-lived memory B cells, while the CD27+IgG+ B cells could be more related to long-lived memory B cells.

In humans, IgG2 are mainly reactive to polysaccharides while IgG3 react with proteins and are more potent than IgG2 in classical complement activation and Ab-dependent cell-mediated cytotoxicity (79). Therefore, the differences between IgG2 and IgG3 expression within CD27+IgG+ and CD27IgG+ cells further suggest that these cells could be distinct in relation to the nature of the Ags and the outcome of the immunological response, as already proposed for the rotavirus-specific B cell response (29, 80).

CD27 can be triggered by its ligand, CD70, which is expressed on activated B and T cells (69, 81, 82). In human B cells, CD27 triggering promotes mainly differentiation into plasma cells (69). However, interaction between CD27 and CD70 generates negative feedback signal to leukocytes during immune activation (83) and is also involved in T cell homeostasis (84, 85). Disrupted interaction through CD27 and CD70 also correlates with the progression of multiple myeloma and the loss of CD27 expression (86). Therefore, CD27IgG+ cells, in opposition to CD27+IgG+ cells, are essentially disconnected from any further regulation through CD70+ T or B cells (69, 81, 82), supporting once more their distinct roles in immune response.

We have noted that CD27IgG+ blood cell characteristics were close to that of hairy cell leukemia cells. These cells are indeed characterized by the lack of CD27 expression (27, 28, 87), the presence of B220 (53), and expression of switched Ig with a dominance in the IgG3 subclass and somatically mutated Ig genes (27, 28, 88, 89, 90, 91), with frequency of mutations ranging from 0 to 5% (28, 88, 89, 90). Interestingly, these cells show heterogeneous mutation frequency, with 6 cases on 32 without mutation (90), which also characterizes CD27IgG+ cells. Approximately 40% of hairy cell leukemia cases coexpress multiple, clonally related Ig isotypes (reviewed in Ref. 92), which was not observed in the CD27IgG+ population. Nevertheless, as suggested by Forconi’s work (27, 28), the malignant change could occur during the generation of CD27IgG+ cells before the deletional recombination events characterizing normal isotype switching, which would allow the expression of several Ig isotypes by a single cell.

Although the exact origin of hairy cell leukemia cells, whether derived from marginal zone, germinal center (28, 89, 90, 92, 93, 94), or T cell-independent reaction (95), is still unclear, this report of a new CD27IgG+ blood B cell subset showing similar characteristics could help in future investigations. Furthermore, the possible link with a first wave of early memory B cells could also be of interest in a better understanding of hairy cell leukemia

Overall, the characterization of a CD19+CD27IgG+ population brings a new look in the organization and homeostasis of peripheral B cells in humans, not only by extending the memory B cell reservoir beyond the CD27+ compartment, but also by questioning the role of the lack of interaction with CD70 in B cell homeostasis. We are currently investigating whether this minor CD27IgG+ B cell population can be stimulated in vitro to express CD27 and to respond to variable level of CD40 stimulation.

We thank Drs. André Darveau, Daniel Jung, and Marguerite Massinga-Loembé for helpful discussions and suggestions and Gerry Boucher for his help in sequence analysis. We are thankful to Annie Roy and Claudia Racine for skillful assistance in cell culture. We are also grateful to Jean-François Leblanc for editing the manuscript. Finally, we are thankful to all participants to this study and to Claudine Côté for the coordination of blood sample collection.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

J.F.F. was supported by a Ph.D. fellowship from le Fond Québécois de la Recherche sur la Nature et les Technologies and from Héma-Québec.

3

Abbreviations used in this paper: ABC, ATP-binding cassette; MTG, MitoTracker Green; IMGT, ImMunoGeneTics; FR, framework region; R, replacement; S, silent; MFI, mean fluorescence intensity; FcRH4, FcR homolog 4.

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