Hematologic disorder–associated Cxcr4 gain-of-function mutation leads to uncontrolled extrafollicular immune response

Following infections, rapid production of neutralizing antibodies (Abs) through the extrafollicular immune response is essential to control infectious agents. Despite the central importance of this pathway in humoral immunity, the molecular mechanisms at play are poorly understood. Early B-cell activation and differentiation initiated through crosslinking of the B-cell receptor (BCR) and Toll-like receptors (TLRs) leads to the production of low-affinity Abs by short-lived extrafollicular plasma cells (PCs), also called plasmablasts (PBs), which provide rapid but transient protection against infection.¹  Upon T-cell help, this early response can evolve through the formation of germinal centers (GCs), to the production of high-affinity long-lived PCs that can persist for an extended period of time in the bone marrow (BM) contributing to immunologic memory.²  The chemokine receptor Cxcr4 and its ligand Cxcl12 are important for the T-dependent immune response through regulation of GC B-cell localization and BM homing of PCs.^3-8  However, the role of this axis in early extrafollicular B-cell response after antigenic challenge has never been addressed. Following Cxcl12 binding, Cxcr4 signals through the G proteins. Signal duration and strength is tightly regulated by receptor desensitization, which involves phosphorylation of the C-terminal tail of Cxcr4 and the subsequent recruitment of β-arrestins, leading to both uncoupling of the receptor from G proteins and receptor internalization.⁹  Our group and others have established that this desensitization step is defective in the rare primary immunodeficiency disease warts, hypogammaglobulinemia, infections, and myelokathexis (WHIM) syndrome (WS),^10,11  which is caused by autosomal dominant mutations in the C-tail of CXCR4, resulting in a gain-of-function in response to CXCL12.^10,12  Strikingly, similar mutations of CXCR4 have been reported in Waldenström macroglobulinemia, an indolent lymphoplasmacytic B-cell lymphoma characterized by an immunoglobulin M (IgM) spike and BM infiltration of malignant clones with an immunophenotype ranging from activated B cells to PCs.^13-15 CXCR4 mutations are observed in ∼30% of Waldenström macroglobulinemia patients in conjunction with a gain of function mutation of the TLR adaptor MYD88 and may be associated with treatment resistance.^16,17  Whether and how CXCR4 and MYD88 mutations synergize in Waldenström macroglobulinemia is still unclear; however, their coincidence in this disease prompted our interest in investigating the general importance of Cxcr4 signaling in the context of TLR-induced MyD88 signaling for B cell and PC biology. Here, using a mouse model¹⁸  bearing an orthologous Cxcr4 mutation observed both in WS and Waldenström macroglobulinemia,^13,14  we demonstrate an unexpected and central role for Cxcr4 desensitization in regulating the early extrafollicular antibody response.

CXCR4^+/1013 mice were generated as previously described.¹⁸  Blimp1^GFP/+ mice were obtained from S. Nutt (The Walter and Eliza Hall Institute of Medical Research, Melbourne, Victoria, Australia)¹⁹  and crossed with CXCR4^+/1013 mice. Mice aged 8 to 16 weeks old or 1 year old and cohoused littermates were used in all experiments that were conducted in compliance with the European Union guide for the care and use of laboratory animals, which have been reviewed and approved by an institutional review committee (C2EA-26, Animal Care and Use Committee, Villejuif, France).

Cryopreserved BM aspirates from 2 WS patients (1 male, age 47 years; 1 female, age >30 years at sampling, both bearing the CXCR4^R334X mutation; collected under National Institutes of Health protocol 09-I-0200) were provided through a Material Transfer Agreement in accordance with National Institutes of Health human subject research policies. BM from hip replacement surgery was obtained from discarded material from 6 patients (4 males and 2 females; 42-58 years old at sampling). All the required ethical and export approvals were obtained from the French Ministries of Health and Agriculture. All individuals provided written informed consent before sampling and data registering. The study was approved by the Comité de Protection des Personnes Ile-de-France VII (protocol 17-030, ID-RCB 2017-A01019-44). Samples were stained with the antibodies indicated in supplemental Table 1 available on the Blood Web site.

Mice were immunized intraperitoneally with 25 µg 4-hydroxy-3-nitrophenylacetyl-lipopolysaccharide (NP-LPS) or 25 µg NP-Ficoll (Biosearch Technologies). Blood, spleen, and BM were harvested and analyzed 3, 6, 12, and 45 days after challenge with NP-LPS and 6 days after NP-Ficoll immunization.

Isolation of organs was performed, and single cell suspensions were stained as previously described,⁷  using the Abs described in supplemental Table 1. Analyses were carried out on an LSRII Fortessa flow cytometer (BD Biosciences), and data were analyzed with the Flowjo software (TreeStar, Ashland, OR).

For functional assays, total splenocytes or B cells isolated using a naïve mouse B-cell (CD43⁻) isolation kit (Miltenyi Biotec) were cultured using complete medium (RPMI with 10% fetal calf serum, 1% penicillin/streptomycin, 50 µM β-mercaptoethanol, 1 mM sodium pyruvate; Gibco) with 1 U/mL interleukin 4 (IL-4; Miltenyi) and 5 ng/mL IL-5 (Miltenyi) supplemented with 1 µg/mL LPS (Invivogen) or 10 µg/mL CpG oligodeoxynucleotides (Invivogen) and/or 50 nM Cxcl12 (R&D), and/or 10 µM AMD3100 (Sigma Aldrich) for 5 minutes and up to 5 days at 37°C. MK-2206 (Ambeed) and rapamycin (Life Technologies) were used at 1 µM and 50 nM, respectively. Phosphoflow was performed using the PerFix EXPOSE kit (Beckman Coulter). PB differentiation and apoptosis were assessed by flow cytometry (supplemental Table 1). For proliferation, splenocytes were loaded with cell trace violet (CTV) (Thermofisher) according to the manufacturer’s instructions and then stimulated. For cell cycle analysis, cells were permeabilized and fixed with the Foxp3/transcription factor staining buffer set (eBioscience) and labeled with an anti–Ki-67 antibody, and 4′,6-diamidino-2-phenylindole (DAPI) was added just before analysis.

Splenocytes from wild-type (WT) × Blimp1^GFP/+ and Cxcr4^+/1013 × Blimp1^GFP/+ mice were stimulated in vitro with LPS for 4 days. Cells from each genotype were loaded with either CTV or CT yellow (CTY; Thermofisher). A 50:50 mix of cells was made and checked by flow cytometry before injection. A total of 10⁶ cells were transferred intravenously into C57Bl/6J CD45.1 WT recipients. Blood, BM, and spleens were harvested and analyzed at 24 and 48 hours after transfer.

Chemotaxis assays were performed using a Transwell assay as previously described.¹⁰  Splenocytes were incubated for 45 minutes at 37°C before migration for 3 hours through a 5-µm pore Transwell insert (Corning) with medium only or with Cxcl12 (R&D) in the lower chamber. Input cells that migrated to the lower chamber were collected, stained, and enumerated by flow cytometry. The fraction of cells migrating across the membrane was calculated as follows: ([number of cells migrating to the lower chamber in response to chemokine or medium]/[number of cells added to the upper chamber at the start of the assay]) × 100.

Spleens were fixed for 4 hours in phosphate-buffered saline (PBS)-paraformaldehyde 4% and incubated overnight in PBS-sucrose 20% before being embedded in optimum cutting temperature compound (TissueTek). Cryosections were permeabilized with PBS Triton X-100 0.5% for 15 minutes, washed twice with PBS, and blocked with PBS-bovine serum albumin (BSA) 5%. Sections were stained with primary Abs and secondary Ab (supplemental Table 1) and Hoechst 33342. Slides were scanned using a NanoZoomer Digital Pathology system using ×40 objective lenses with numerical aperture of 0.75 (Hamamatsu Photonic). BM embedding, sectioning, and staining were performed as described.²⁰  Sections (30 µm) were permeabilized, saturated, and stained as for the spleen. Images were acquired using a TCS SP8 confocal microscope and processed and quantified with Fiji and Imaris.

Anti-NP and anti-IgM ELISA were performed as previously described.⁷  Enumeration of total IgM and NP-specific IgM Ab-secreting cells (ASCs) were performed by incubating spleen or BM cells overnight at 37°C on plates precoated with NP15-BSA or goat anti-mouse IgM at 5 µg/mL and saturated with culture medium containing 10% fetal calf serum. Plates were then washed and incubated with peroxidase-conjugated goat anti-mouse IgM (Southern Biotech). Spots were revealed with 3-amino-9-ethylcarbazole (Sigma-Aldrich) and analyzed using an ispot Reader (Autoimmun Diagnostika GMBH).

B cells were isolated and stimulated with LPS. Cells were either imaged using a Biostation (NIKON) in a 37°C, 5% CO₂ supplemented chamber at a concentration of 10⁴ cells/mL or using the Incucyte S3 at a concentration of 10⁴ cells per well. Cells were imaged for phase-contrast and green fluorescent protein (GFP) fluorescence every 15 or 30 minutes, respectively. Images and movies were analyzed with the biostation IM software, the Incucyte S3 software, and Fiji.

Multiplex quantitative polymerase chain reaction (qPCR) analysis was performed using the Biomark system (Fluidigm). Seventy-five cells per well were sorted directly into reverse transcription/preamplification mix, as previously described²¹  (supplemental Table 2). Targeted cDNA preamplification was performed before processing with the dynamic array protocol according to the manufacturer’s instructions (Fluidigm). Mean expression of Actb and Gapdh was used for normalization.

Data are expressed as means ± standard error of the mean (SEM). The statistical significance between groups was evaluated using the 2-tailed unpaired Mann-Whitney nonparametric test (Prism software, GraphPad) unless indicated otherwise in figure legends. Heatmaps were presented using the Z score, and clustering analyses were performed using the Pearson rank correlation test (http://www.heatmapper.ca). Principal component analysis (PCA) was performed using R ggplot packages.

To better understand the pathophysiology of WS, we previously generated a knock-in mouse model bearing a heterozygous nonsense point mutation common to a subset of patients (mutation C1013G in the open reading frame).¹⁸  These mice, referred to as Cxcr4^+/1013, phenocopy the pan-leukopenia, a major hematologic feature of the disease.¹⁸  Moreover, using this model, we recently showed that the gain of function of Cxcr4 affects in a B-cell intrinsic manner the T-dependent humoral immune response recapitulating vaccination failure reported in some WS patients.^7,12,22,23  We took advantage of this model to assess whether Cxcr4 signaling may play a role in the early extrafollicular response and in particular after stimulation with MyD88-dependent TLR ligands. First, we stimulated in vitro splenic B cells from WT and Cxcr4^+/1013 mice with the TLR4 and TLR9 ligands, LPS and CpG, respectively, in the presence or absence of Cxcl12. Under these conditions, PB differentiation, defined as formation of B220^lowCD138⁺ cells, was increased for B cells from Cxcr4^+/1013 mice (Figure 1A-B; supplemental Figure 1A-B) and was associated with a twofold increase in IgM concentration measured in the culture supernatant (Figure 1B). The addition of Cxcl12 to the TLR agonists in the stimulation medium led to a small increase of PB in both WT and Cxcr4^+/1013 B-cell cultures (Figure 1B; supplemental Figure 1A-B). Interestingly, stimulation with Cxcl12 alone promoted some PB differentiation of Cxcr4^+/1013 B cells, a phenomenon significantly less marked in WT cultures (Figure 1B; supplemental Figure 1B). This Cxcl12-mediated PC differentiation was abolished by addition of the Cxcr4 antagonist AMD3100 confirming that Cxcr4 signaling promoted it (supplemental Figure 1B). These observations thus hinted toward a role for Cxcr4 signaling in early B-cell activation and differentiation after stimulation with TLR ligands.

We next determined whether the gain of function of Cxcr4 could affect T-independent B-cell immune responses in vivo. Immunization with the T-independent type 1 antigen NP-LPS did not correct the severe B-cell lymphopenia observed in the spleens of Cxcr4^+/1013 mice¹⁸  (Figure 1C-D). However, compared with WT mice, we observed a significant increase in the frequency of splenic PBs in Cxcr4^+/1013 mice from days 3 through 6 after immunization (Figure 1E-F). Consistent with this, we detected an increased frequency of IgM⁺ ASCs by ELISpot in the spleens of Cxcr4^+/1013 mice compared with WT mice (Figure 1G), and an increased titer of total IgM in the serum of mice bearing the Cxcr4 gain-of-function mutation (supplemental Figure 1C). Moreover, the frequencies of NP-specific IgM⁺ ASCs in the spleen were increased at day 6 in Cxcr4^+/1013 mice compared with WT mice (Figure 1G). Importantly, the T-independent B cell response to NP-LPS immunization was also more sustained in Cxcr4^+/1013 mice compared with WT mice as demonstrated by the still elevated frequency and number of PBs observed at day 6 in the mutant mice (Figure 1F-G). Analysis of the splenic PB population by imaging revealed an increased detection of IgM⁺ PBs in sections of Cxcr4^+/1013 spleens compared with WT tissues (Figure 1H-I). Similarly, immunization with the T-independent type 2 NP-Ficoll led to increased splenic PBs and increased total IgM⁺ serum titers (Figure 1J-K) in the mutant mice compared with controls. Together, these results show that Cxcr4 desensitization is an important negative regulator of extrafollicular PB generation in the spleen.

We next addressed the cellular mechanism underlying enhanced PB generation in Cxcr4^+/1013 mice. We cultured WT × Blimp1-GFP and Cxcr4^+/1013 × Blimp1-GFP splenic B cells with LPS and assessed the kinetics of appearance of GFP as a readout for PB differentiation by video microscopy. As shown in Figure 2A, some GFP⁺ cells were detectable in Cxcr4^+/1013 cultures as early as 46 hours after stimulation, whereas they started appearing in WT cultures 20 hours later (Figure 2A-B). As B-cell and PB survival seemed unaffected (supplemental Figure 2A-B), this accelerated appearance and accumulation of PBs in culture could be caused by an enhanced cycling and/or proliferation of B cells and PBs. B-cell proliferation, measured by CTV dye dilution in vitro after LPS stimulation, was indeed altered in the mutant condition at day 2 both with and without exogenous Cxcl12 addition (Figure 2C-D; supplemental Figure 2C). However, from day 3, B-cell proliferation was equivalent between WT and mutant B cells (Figure 2D), suggesting this was only a transient phenotype. Likewise, proliferation in newly generated PBs was not different between WT and mutant conditions at any of the time points analyzed (Figure 2E-F; supplemental Figure 2D). Previous reports have suggested that B-cell differentiation into PB necessitates at least 3 rounds of cell division.²⁴  Together with our results, this brought us to consider whether it was not the proliferation of B cells per se that was affected, but rather their entry into the cell cycle. Indeed, we observed a significant increase in the percentage of B cells in cycle, especially in the G1 phase, in Cxcr4^+/1013 cultures compared with WT ones (Figure 2G-H). Addition of Cxcl12 to LPS in the culture medium led to a similar result, and treatment with AMD3100 had no effect on the exacerbated entry into cell cycle observed in mutant B cells (supplemental Figure 2E-F). As expected, most PBs were cycling,²⁵  and no difference was observed between our 2 experimental groups for this cell type (Figure 2I). This demonstrates that Cxcr4 desensitization is an essential negative regulator of B cell entry into cycle and that deregulation of this process contributes to exacerbated PB differentiation.

A wealth of literature reports that, on TLR crosslinking, B-cell activation and proliferation are dependent on phosphatidylinositol 3-kinase activation and downstream Akt and mechanistic target of rapamycin (mTOR) signaling.^26-29  Moreover, several works including ours showed that crosslinking of mutant Cxcr4 on B cells promoted aberrant activation of the Akt and mTORC1 pathways.^7,30,31  We confirmed ex vivo that the gain of function of Cxcr4 led to an exacerbated Akt phosphorylation on Cxcl12 stimulation in the presence or absence of TLR crosslinking. However, inhibition of Akt did not impact B cell cycle entry in this system (Figure 2H; supplemental Figure 3C). In parallel, we observed that Cxcl12 stimulation led to a transient increase of the mTORC1 target pS6 in B cells, whereas LPS stimulation promoted strong and sustained S6 activation. Importantly, pS6 signaling was significantly increased in mutant B cells compared with WT controls in all conditions and that was even more marked when both Cxcl12 and LPS were used conjointly (Figure 3A-B). Moreover, AMD3100 preincubation was reducing the frequency of pS6⁺ mutant B cells after Cxcl12 stimulation, hence supporting the role of Cxcr4 gain of function in this process (Figure 3B). These results suggest that TLR4 and the mutant Cxcr4 synergize to promote exacerbated and sustained mTORC1 signaling in Cxcr4^+/1013 B cells. Furthermore, treatment with the mTORC1 inhibitor rapamycin significantly reduced LPS-induced cell cycle entry in WT and Cxcr4^+/1013 B cells (Figure 3C). In line with this, when rapamycin was added, mutant B-cell differentiation was down to the level observed for untreated WT B cells (Figure 3D). Hence, our data suggest that mutant Cxcr4 and TLR4 signaling synergize to promote stronger mTORC1 signaling, leading to increased B cell entry into cycle and subsequently enhanced PB differentiation.

The T-independent extrafollicular B-cell response mainly leads to the generation of short-lived PBs, although it has been reported that some may be able to home to and persist in the BM.^32-34  We therefore investigated whether the enhanced extrafollicular response observed in the spleen in Cxcr4^+/1013 mice was mirrored by changes in BM homing. After NP-LPS immunization, we observed an increased detection of total IgM⁺ ASCs from day 3 and of NP-specific IgM⁺ ASCs from day 6 in the BM of Cxcr4^+/1013 X Blimp1-GFP mice compared with WT mice (Figure 4A-D). This was associated with a fivefold increase in PBs (Blimp1-GFP^hi CD138⁺) in the BM of mutant mice compared with WT mice from day 3 after immunization (Figure 4E). Moreover, fully differentiated PCs (Blimp1-GFP^low CD138⁺) were significantly increased from day 6 in the BM of the Cxcr4^+/1013 mice compared with their WT littermates (Figure 4E). Consistently, we enumerated in situ significantly more IgM⁺ PB/PCs per mm² at day 6 after NP-LPS immunization in the BM sections of Cxcr4^+/1013 mice compared with WT mice (Figure 4F-G). Thus, Cxcr4 desensitization is essential to limit BM accumulation of PBs generated through the extrafollicular response.

Previous reports have suggested that PB/PCs increase with age in mice.³⁵  Accordingly, 1-year-old middle-aged Cxcr4^+/1013 mice display a very significant increase in the frequency and number of PBs compared with WT mice at steady state that was particularly marked for BM PBs (Figure 5A-D). Thus, the spontaneous and ongoing immune response observed in mice as they age appears exacerbated by the Cxcr4 gain-of-function mutation, and this translates into an abberant accumulation of PBs within the BM.

We next investigated whether CXCR4 gain-of-function mutations might also distort the basal distribution of PCs in BM from WS patients. To this end, we compared BM samples from 2 WS patients carrying the heterozygous CXCR4^R334X mutation with aged-matched control BMs. When we compared WS and control BMs, we observed that the frequencies of PBs (CD19⁺CD138⁻) were significantly increased in patients compared with control BM (Figure 5E-F), with the 2 less mature fractions representing most of the BM PB/PCs in the patient samples (Figure 5G). Altogether, these results suggest that gain-of-function mutations of CXCR4 can be associated with spontaneous aberrant accumulation of PBs within the BM in middle-aged mice and humans.

We then explored whether the exacerbated PB accumulation within the BM observed in absence of Cxcr4 desensitization could be accounted for by a cell intrinsic advantage in terms of BM homing. Mutant splenic PBs express slightly but significantly more surface Cxcr4 than their WT counterpart (Figure 6A-B). Moreover, in a transwell-based migration assay, mutant splenic PBs migrated more toward Cxcl12 in vitro than their WT counterparts, and this was abrogated by the addition of AMD3100 (Figure 6C). Moreover, by cotransfer experiments of in vitro–generated PBs loaded with either CTV or CTY and mixed at a 1:1 ratio (Figure 6D-E), we showed that Cxcr4^+/1013 mutant PBs had an intrinsic advantage for homing to the BM compared with WT PBs at all time points considered, whereas the reverse was true for the spleen (Figure 6F). This experiment demonstrates that, on top of their enhanced generation in secondary lymphoid organs, mutant PBs also have an intrinsic advantage in terms of BM homing.

Transcriptional profiling of WT and Cxcr4^+/1013 BM PBs revealed significant differences with a notable alteration in the expression of several genes involved in cell adhesion and migration (Figure 6G). We thus further assessed the impact of Cxcr4 signaling on the migratory and adhesion properties of BM PBs and PCs. We showed that mutant PBs and PCs had a unique transcriptional profile characterized by increased expression of Cxcr4, Cxcr3, Itgal, Cd44, and Itgb1 and decreased expression of Ccr7, Cxcr5, Sell, and Klf2 compared with WT PBs and PCs (Figure 6H). These differences were sufficient to differentially cluster WT and Cxcr4^+/1013 BM PB/PCs in a PCA (Figure 6I). Moreover, these altered expressions were confirmed by classical reverse transcriptase-qPCR (supplemental Figure 4A-C) and flow cytometry (Figure 6J; supplemental Figure 4D-E). In summary, our data support that Cxcr4 desensitization is a key control mechanism that intrinsically limits the homing of PBs to the BM, and that once there, this gain of function may confer them with specific localization and/or cell-cell interaction potential.

We last questioned the fate of these aberrant PBs within the BM. The number of PBs in the BM of Cxcr4^+/1013 mice peaked at day 6 and then rapidly declined and was back to the baseline WT level 12 days after NP-LPS immunization (Figure 7A). In parallel, the number of more mature B220⁻ PCs progressively and markedly increased in the BM of Cxcr4^+/1013 mice, reaching a peak between 8 and 12 days after immunization, and then slowly declined (Figure 7A). The distinct kinetics for PBs and PCs suggest that PBs reaching the BM of Cxcr4^+/1013 mice may be able to differentiate into PCs. We exploited our transfer experiments (Figure 6D) and observed that at day 1 in the BM, most of the transferred cells were found in the PB gate (CD138^hiB220⁺), and very few were detected within the PC gate (CD138^hiB220⁻; Figure 7B). At day 2, however, most of the transferred cells were detected among the CD138^hiB220⁻ PCs (Figure 7B), and approximately 80% of them came from the Cxcr4^+/1013 cells, whereas the reverse was true for PBs (Figure 7C). In addition, we analyzed the expression of key developmental factors of PCs and observed that Cxcr4^+/1013 BM PBs had a transcriptional profile corresponding to more mature cells than their WT counterparts, albeit less mature than fully differentiated PCs, and characterized by decreased expression of the B cell transcription factors Pax5 and Bach2 and enhanced expression of the PC transcription factors Prdm1, Xbp1, and Irf4 (Figure 7D). Finally, we observed that total and NP-specific IgM⁺ ASCs were increased in the BM of Cxcr4^+/1013 mice compared with the WT ones, and this increase was still observed 45 days after immunization (Figure 7E). Thus, Cxcr4^+/1013 PBs may differentiate into PCs and persist for more than 1 month in the BM. Altogether, our results demonstrate that Cxcr4 desensitization limits both PB maturation into PCs and their persistence within the BM.

Despite its essential role in host protection, how the extrafollicular immune response is regulated remains mysterious. Here we demonstrate that Cxcr4 desensitization is an essential regulatory mechanism controlling the duration and the strength of Ab production after induction of a T-independent immune response by finely modulating the pool of extrafollicular-derived PCs.

Our data support a critical role for Cxcr4 desensitization during this process at multiple checkpoints of B-cell activation and PC differentiation. First, we observed that impaired Cxcr4 desensitization led to increased TLR-driven B-cell differentiation into PB with, as consequences, more PBs generated and increased IgM production in the course of an extrafollicular response. This is associated with enhanced entry into cycle and is consistent with previous studies that have linked B-cell division to their capacity to differentiate into PCs.^25,36,37  In addition, we showed that Cxcr4 gain of function following Cxcl12 stimulation and in synergy with TLR signaling promoted induction of the mTORC1 pathway that drives cell cycle entry and PB differentiation.

Second, PBs and PCs bearing the Cxcr4 gain-of-function mutation display an enhanced BM tropism that may not only be caused by the gain of function of Cxcr4 per se as the expression of a whole range of migration/adhesion factors are modified in PBs in Cxcr4^+/1013 mice, including integrins that can be activated by Cxcr4 signaling.^38,39 

Third, differentiation of PBs into PCs and their persistence are enhanced in the BM of Cxcr4^+/1013 mice compared with WT mice. This could be through direct effects of the enhanced Cxcr4 signaling on the PC maturation transcriptional program. Alternatively or conjointly, different environmental cues specific to mutant PB/PCs and linked with their altered adhesion/migration potential might contribute to this phenomenon. Importantly, some insights gained from the study of a Cxcr4 gain-of-function mouse model were validated in the BM of WS patients in whom we observed an over-representation of PBs.

Conceptually our results suggest an important role for Cxcr4 signaling in limiting both the number of PBs generated and the lapse of time during which Abs generated in the course of a TLR-dependent extrafollicular response are produced. Indeed, Cxcr4 desensitization not only regulates the early generation of PBs in secondary lymphoid organs after immunization but also their homing to and maturation within the BM, hence ensuring that only a few PCs generated during this response persist in the BM. As such, Cxcr4 desensitization seems to act as a gatekeeper, ensuring that the Abs produced by PBs during the extrafollicular response are restricted to a transient wave of protection. Limiting PB homing to the BM in this way may also advert the filling of BM niches normally dedicated to other cell types, including GC-derived high affinity PCs that start arising around day 9 after immunization.²  Thus, our results, taken together with our previously published data,⁷  suggest that defective long-term vaccine protection, previously reported in WS patients,^12,22,23  may be the consequence of defective PC homeostasis with an over-representation of low-affinity extrafollicular PBs and reduced contribution to the humoral response of GC-derived high affinity PCs.

Somatic CXCR4 mutations are observed in ∼30% of Waldenström macroglobulinemia patients, most of the time in conjunction with a gain-of-function mutation of the TLR adaptor MYD88,^16,17,40,41  and the presence of the 2 mutations is associated with shorter time to treatment and earlier progression with ibrutinib.^13,15,31,42  Our data thus support that CXCR4 nonsense mutations leading to premature stop codon may promote BM homing of malignant cells and niche-mediated therapy resistance previously reported in this disease.^43,44  Moreover, they suggest the paradigm-shifting hypothesis of a potential CXCR4-dependent extrafollicular origin for Waldenström macroglobulinemia malignant clones. Whether CXCR4 frameshift mutations, also observed in some Waldenström macroglobulinemia patients, may have similar impact remains to be evaluated.^45,46 

The authors thank M.-L. Aknin, H. Gary, F. Gaudin, V. Nicolas, B. Lecomte (Ingénierie et Plateformes au Service de l’Innovation Thérapeutique, technical facilities Plateforme de Cytométrie en Flux [PLAIMMO], Plateforme d’Histologie Expérimentale [PHIC], Plateforme d'Imagerie Cellulaire [MIPSIT] and AnimEx, Clamart), J. Megret, C. Cordier (Flow Cytometry Core Facility, Structure Fédérative de Recherche Necker, Paris, France), P. Chappert, A. Vandenberghe (Institut Necker-Enfants Malades, Paris, France), C. Doliger, and S. Duchez (Flow Cytometry Core Facility, Institut de Recherche Saint Louis, Paris, France) for technical assistance and M.A. Linterman, S. Luther, and S. Peebles for comments on our manuscript.

The study was supported by the Laboratory of Excellence in Research on Medication and Innovative Therapeutics (LabEx LERMIT) (M.E. and K.B.); an Agence Nationale de la Recherche (ANR) Jeunes Chercheurs/Jeunes Chercheuses grant (ANR-19-CE15-0019-01), an ANR @RAction grant (ANR-14-ACHN-0008), a Fondation Arthritis grant, a grant from IdEx Université de Paris (ANR-18-IDEX-0001), and a “Fondation ARC pour la recherche sur le cancer” grant (P JA20181208173) (M.E.); and an ANR Projets de Recherche Collaborative grant (ANR-17-CE14-0019), an Institut National du Cancer grant (PRT-K 2017), and the Association Saint Louis pour la Recherche sur les Leucémies (K.B.). D.H.M and P.M.M. are supported by the Division of Intramural Research of the National Institute of Allergy and Infectious Diseases, National Institutes of Health. N.A. and M.K. were supported by a fellowship from the French Ministry for education and by a fourth-year fellowship from the “Fondation ARC pour la recherche sur le cancer” (N.A.). V.R. was supported by a fellowship from the Fondation pour la Recherche Médicale and a fourth-year fellowship from the Ligue Nationale Contre le Cancer. R.H.-A. was supported by a master fellowship from the “Fondation ARC pour la recherche sur le cancer.” The EMiLy U1160 INSERM unit is a member of the OPALE Carnot Institute, The Organization for Partnerships in Leukemia.

Contribution: N.A. designed and performed experiments, analyzed data, and wrote the manuscript; A.B. performed experiments, analyzed data, and wrote the manuscript; V.R. and R.H.-A. performed experiments and analyzed data; J.N., V.B., M.K., E.C., N.S., N.D., and M.M. helped with experiments; P.M.M. and D.H.M. provided essential human reagents and reviewed the manuscript; K.B. contributed to the project design and to the manuscript redaction; and M.E. designed the project, analyzed data, and wrote the manuscript.

Conflict-of-interest disclosure: The authors declare no competing financial interests.

Correspondence: Marion Espéli, Institut de Recherche Saint Louis, INSERM U1160, 1 Av Claude Vellefaux, 75010 Paris, France; e-mail: marion.espeli@inserm.fr.