Extracellular Release of Antigen by Dendritic Cell Regurgitation Promotes B Cell Activation through NF-κB/cRel

Houssam El-Barbry, Marisa Capitao, Sarah Barrin, Samir Amziani, Pascal Pierre Paul, Susanna Borreill, Thomas Guilbert, Emmanuel Donnadieu, Florence Niedergang and Fatah Ouaaz
J Immunol August 1, 2020, 205 (3) 608-618; DOI: https://doi.org/10.4049/jimmunol.1900394

Key Points

  • Dendritic cells transfer Ags to and activate B cells in vivo.

  • Ags released from dendritic cells by regurgitation promote B cell activation.

  • Early B cell activation by dendritic cell regurgitation requires NF-κB/cRel.

Abstract

Dendritic cells (DCs) are professional APCs, which sample Ags in the periphery and migrate to the lymph node where they activate T cells. DCs can also present native Ag to B cells through interactions observed both in vitro and in vivo. However, the mechanisms of Ag transfer and B cell activation by DCs remain incompletely understood. In this study, we report that murine DCs are an important cell transporter of Ag from the periphery to the lymph node B cell zone and also potent inducers of B cell activation both in vivo and in vitro. Importantly, we highlight a novel extracellular mechanism of B cell activation by DCs. In this study, we demonstrate that Ag released upon DC regurgitation is sufficient to efficiently induce early B cell activation, which is BCR driven and mechanistically dependent on the nuclear accumulation of the transcription factor NF-κB/cRel. Thus, our study provides new mechanistic insights into Ag delivery and B cell activation modalities by DCs and a promising approach for targeting NF-κB/cRel pathway to modulate the DC-elicited B cell responses.

This article is featured in In This Issue, p.553

Introduction

Dendritic cells (DCs) are professional APC that sample Ags in the periphery and migrate to lymph nodes (LNs) where they activate T cells and potentially B cells. Activated T cells by DCs could differentiate into follicular helper T cells that provide help to B cells in T-dependent B cell responses in vivo (1, 2). Whereas T cells recognize “degraded” Ags, B cells recognize their cognate Ag in its “native” form. Recent advances in two-photon imaging have provided major insights into Ag encounter by B cells in vivo. Several routes of Ag delivery to the LN B cell compartment have been described, such as the importance of LN follicular DCs (FDC), subcapsular sinus macrophages, follicular reticular cells conduits, or resident DCs (3, 4). The dual ability of DCs to promote Ag presentation to both T and B cells is a major hallmark of the key role of these professional APCs in the host adaptive immunity against infection (5).

Ag presentation to B cells is crucial for B cell activation and the initiation of the humoral response that occurs mainly within the follicular region of secondary lymphoid organs, in which B cells encounter their cognate Ag (6). Activated B cells then undergo clonal expansion, selection, and differentiation into memory or Ab secreting–plasma cells (7). Humoral immunity is essential for protection against cancer and vaccination against infections by generating high affinity Abs that neutralize pathogens in peripheral tissues and at the mucosa sites (8).

We have highlighted, using three-model Ags including the enzymatic activity of HRP, that DCs are able to store Ags captured by macropinocytosis and that they release from late endosomes in their native form in the extracellular medium through a process that was called “regurgitation” (4, 9). It is often and still considered that B cell activation is predominantly promoted by Ags presented on the surface of APCs (6, 10–12). B cells indeed are able to extract Ag displayed on the APC surface either by enzymatic degradation upon local lysosomal secretion at the immunological synapse (13, 14) or by using actomyosin cytoskeleton-driven mechanical forces (15–17). However, extracellular release of native Ag by DCs has not been characterized yet as another mechanism of Ag presentation that could potentially impact direct B cell activation and/or differentiation.

NF-κB is a major transcription factor, crucial for both B cell (18) and DC development and functions as we have previously shown (19). The NF-κB family consists of five Rel subunits: p50 (NF-κB1), p52 (NF-κB2), p65 (RelA), RelB, and cRel, which function as homo- or heterodimers to regulate the expression of genes important for inflammatory and immune responses (20, 21). As a hallmark of an early intracellular event of B cell activation, BCR Ag stimulation triggers the canonical NF-κB mobilization through the recruitment of the two major subunits RelA (p65) and cRel (22, 23). In contrast to transient mobilization of RelA, cRel is considered as the predominant subunit of the MALT-1–derived NF-κB that is activated with sustained nuclear accumulation after BCR stimulation. Furthermore, cRel is highlighted also to be the critical NF-κB member essential for the Ag-dependent B cell activation, germinal center reaction, and optimal T-independent Ab responses in vivo (24, 25). Whether cRel may directly or not drive the transcription of genes involved in B cell activation and/or differentiation induced upon Ag delivery by DCs is not known.

Although Ag presentation and T cell activation by DCs is well established, the mechanisms of Ag encounter and initiation of B cell responses by DCs are still incompletely understood and poorly explored. By using an s.c. delivery of Ag-pulsed DCs in vivo and an in vitro coculture system, we have investigated the potential role of DCs in Ag delivery and B cell activation as well as the underlying mechanisms. We report that DCs are an important cell transporter of Ag from the periphery to the LN B cell zone and also potent B cell activators both in vivo and in vitro. In the present study, we showed that Ag released by DC regurgitation induces early B cell activation, which is BCR driven and mechanistically dependent on the nuclear mobilization of NF-κB/cRel.

This work provide new insights into Ag encounter and B cell activation by DCs and a new strategy for drug inhibition of NF-κB/cRel to modulate the DC-promoted B cell responses.

Materials and Methods

Mice

C57BL6/J female mice, 8–12 wk old, were purchased from Janvier Labs, and BCR-transgenic MD4 mice were kindly provided by F. Batista (The Francis Crick Institute, London, U.K.). Mice were maintained in the pathogen-free animal facility of Institut Cochin in accordance with Institut Cochin guidelines in compliance with European animal welfare regulation.

Abs and reagents

The following fluorescent Abs against CD11c, CD8α, CD11b, CD45RA (B220), CD69, CD86, CD138, allotypic IgMa, and CD3 were purchased from BD Biosciences. Mouse B cell negative isolation kit was from Miltenyi Biotec. Qdot605-coupled F(ab′)2 anti-mouse IgG was purchased from Invitrogen. Anti-IgM F(ab′)2 Ab was from Jackson ImmunoResearch Laboratories, and anti-CD3/anti-CD28 Abs were kindly provided by E. Donnadieu. Anti–cRel (sc-71) was from Santa Cruz Biotechnology, anti–β-actin and anti–H3 histone were from Cell Signaling Technology, and anti–hen egg lysozyme (HEL) (ab391) was from Abcam. HEL, LPS (Escherichia coli), and pentoxifylline (PTXF) were purchased from Sigma. ECL Plus Western Blotting detection reagents were from Thermo Fisher Scientific.

Cells

Bone marrow–derived DCs were differentiated from bone marrow cells flushed from femurs and tibias collected from C57BL/6 mice (26). Cells were seeded in six-well plates at the concentration of 1 × 106/ml in RPMI 1640 medium supplemented with 10% FCS and antibiotics in the presence of supernatant (SpN) (4% v/v) from J558 cells transduced with murine GM-CSF. Cells were cultured for 6 d with medium replacement at day 2, 4, and 6. DC differentiation was controlled at day 6 by flow cytometry on the basis of CD11c marker expression. Primary murine splenic B cells from C57BL/6 mice were isolated using MiniMACS magnetic sorting by negative selection (Miltenyi Biotec) to avoid activation of B cells, according to the manufacturer’s instructions. B cell purity was 90–98% based on flow cytometry analysis following B220 staining.

Footpad injections of Ag-pulsed DCs and in vivo analysis

To assess in vivo Ag transport and trafficking to the LN, DCs harvested at day 6 were incubated with 50 μg/ml of Qdot605-coupled F(ab′)2 anti-mouse IgG as fluorescent Ag for 1 h at 37°C. Cells were then washed twice with cold PBS before footpad injection into recipient C57/Bl6 mice. Two groups of three mice each were injected with sterile PBS as controls, and two others of three mice each were injected in each footpad with 2.5 × 106 fluorescent Ag-pulsed DCs. After 18 and 48 h, draining popliteal LNs (PLNs) were collected and embedded in agarose before vibratome sectioning. Two hundred to three hundred micrometer–thick sections were fixed in 1% paraformaldehyde (PFA) and then washed and blocked with PBS supplemented with 2% BSA. Sections were then stained with FITC–anti-CD11c and Alexa Fluor 647–anti-B220 Abs. After washes, sections were covered with glass coverslips before the imaging of Ag trafficking and distribution visualized by a spinning disc confocal microscope.

For in vivo B cell activation analysis, DCs were incubated with 1 mg/ml of HEL as Ag for 1 h at 37°C. Cells were then washed twice with PBS before footpad injection in vivo. Three groups of four to five mice each were used; one group was injected with sterile PBS as control, a second group was injected in each footpad with 2.5 × 106 nonpulsed DCs, and the third group was injected in each footpad with 2.5 × 106 HEL-pulsed DCs (DCs-HEL). After 3 d, PLNs were collected, and single cell suspensions were prepared for flow cytometry analysis of B cell activation and differentiation.

Stimulations and in vitro cocultures

DCs were left untreated or pulsed with 1 mg/ml of HEL for 3 h at 37°C. Half of both DCs and DCs-HEL were washed twice with cold PBS and then chased for 18 h at 37°C. DCs SpN were collected from 18 h chased nonpulsed and HEL-pulsed DCs. Purified wild-type (WT) and transgenic MD4 B cells were seeded in 12-well plates at the cell density of 106/ml. B cells were left untreated or stimulated for 18 h with 102 μg/ml of soluble HEL, 1 ml of DC SpN, or with 1 ml of DC-HEL SpN. In parallel, 106 B cells were cocultured with 2 × 105 DCs (ratio DCs: 1 − B cells: 5), 106 DCs (ratio DCs: 1 − B cells: 1), 2 × 105 DCs-HEL (ratio DCs: 1 − B cells: 5), or with 106 DCs-HEL (ratio DCs: 1 − B cells: 1) for 18 h.

FACS analysis

B cell activation/differentiation induced in vivo and in coculture in vitro were assessed by using, respectively, LN single cell suspensions and purified splenic B cells. Cells were resuspended in FACS buffer (PBS 0.2% BSA) for immunostaining with Alexa Fluor 488–anti-B220, PE CF594–anti-CD69, Brilliant Violet 421–anti-CD86, Brilliant Violet 605–anti-CD138, and PE-IgMa Abs. Samples were analyzed using the BD LSR II cytometer and FlowJo software (BD Biosciences). Results are expressed in the percentage of numbers of B220+ cells coexpressing CD69, CD86, or CD138 surface markers, found in the LNs or present in cocultures in vitro. DC cell necrosis after 18 h of chase was also analyzed in vitro. DCs were left untreated or pulsed with 1 mg/ml of HEL for 3 h at 37°C and then chased for 18 h at 37°C. Chased DCs were then resuspended in FACS buffer for immunostaining with propidium iodide (PI) or with PE-Cy7–anti-CD11c and analyzed using the BD LSR II cytometer and FlowJo software.

Protein extracts preparation and Western blot

DCs were left untreated or pulsed with 1 mg/ml of HEL for 3 h at 37°C and then washed with PBS and chased for 18 h at 37°C. DC SpN were collected after 18 h of chase. Total lysates were prepared from DCs after 3 h of pulse with HEL or 18 h of chase. Nineteen micrograms of proteins from DC lysates, DC SpN, or serial doses of HEL were resolved on SDS-PAGE gel and subjected to Western blot (WB) analysis of Ag detection with anti-HEL and with anti-GAPDH and anti-BSA (loading control) Abs. Purified splenic WT and MD4 B cells were left untreated or stimulated for 18 h with 10 μg/ml of anti-IgM F(ab′)2 Ab, 102 μg/ml of soluble HEL, 1 ml of DC SpN, or with 1 ml of DC-HEL SpN. Cytosolic and nuclear proteins were extracted according to our previous protocol (19). Nineteen micrograms of nuclear protein extracts were separated on SDS-PAGE gel and subjected to WB analysis with anti–cRel and histone H3 (loading control) Abs. For cRel inhibition by PTXF analysis, purified splenic MD4 B cells were left untreated or stimulated in the absence or the presence of PTXF (500 μg/ml). Whole total lysates were prepared before analysis by immunoblotting with anti–cRel and GAPDH (loading control) Abs.

Mass spectrometry analysis and protein quantification

DCs were left untreated or pulsed in vitro with 1 mg/ml of HEL for 3 h, washed twice with cold PBS, and then chased for 18 h at 37°C. SpN were collected from 18 h chased nonpulsed (DC SpN) and HEL-pulsed (DC-HEL SpN) DCs, filtered, and then subjected to mass spectrometry (M/S) analysis that was performed as described previously (27).

Confocal microscopy and image analysis

Imaging of LN sections was performed by using an upright spinning disc (DM6000 FS; Leica CSU-X1; Yokogawa) and an inverted spinning disc (1Xplore, CSU-W1 T1; Yokogawa) confocal microscope, both equipped with a camera (ORCA-Flash, Hamamatsu). Images were acquired with a 10× objective and a 25× water immersion objective. For image analysis, the paracortical, intermediate (IR), and cortical regions were identified by visual inspection of immunofluorescence images of the whole LNs. The absolute number of Ag-loaded DCs (Ag-DCs) was quantified in these respective regions by using ImageJ software.

Statistical analysis

Results are expressed as mean ± SEM. All statistical analyses were performed in GraphPad Prism 7. Statistical significance was tested with an unpaired Student t test and two-way ANOVA with multiple comparisons and was defined when p < 0.05.

Results

Ag transport, trafficking by DCs, and positioning within the LN B cell zone

We first analyzed the Ag uptake by murine bone marrow–derived DCs generated ex vivo from bone marrow after 6 d of GM-CSF differentiation. DCs were harvested at day 6, and the phenotypic differentiation into myeloid DCs was controlled by flow cytometry on the basis of CD11c marker expression (Fig. 1A). Before exploring in vivo the traffic of Ag transported by DCs from the periphery to the LN, we first wanted to assert that DCs were able to capture Ag after pulse in vitro. For this, we used a fluorescent model Ag that can target both IgM and IgG BCR-bearing B cells (9). DCs were pulsed in vitro with 50 μg/ml of Qdot655-coupled F(ab′)2 anti-mouse IgG for 1 h, washed twice, and stained with anti-CD11c PE-Cy7 to analyze Ag capture. DCs were able to capture efficiently fluorescent Ag as indicated by 90% of double positive CD11c+ Qdot655+ DCs (Fig. 1A).

FIGURE 1.
FIGURE 1.

Ag transport by DCs and positioning within the LN B cell zone. DCs were pulsed with Qdot655-coupled IgG and stained for CD11c to analyze Ag capture in vitro by FACS before in vivo footpad injection. (A) Representative FACS profiles in gated CD11c+ cells. (B) Representative image of PLN section from control recipient mice, showing after PBS injection no presence of Ag (red) within the whole LN. Scale bar, 100 μm. (C) Representative image of PLN section from recipient mice, showing at 18 h after injection the arrival of fluorescent Ag (red) within the paracortex area in the LN and colocalization with CD11c+ DCs (green) as indicated by the white arrows. Scale bar, 100 μm. (D) Representative image of PLN section from recipient mice, showing at 48 h after injection the positioning of Ag-DCs (yellow/orange) within the cortical B cell zone cells (blue) as indicated by the white arrows. Scale bar, 100 μm. (E) Quantification of Ag-DCs within the paracortical, IR, and cortical regions of PLNs from two independent experiments (n = 1, right panel; n = 2, left panel). The absolute number of Ag-DCs was quantified in these respective regions by using ImageJ software.

We then followed the migration of Ag-DCs from the periphery to the draining PLN to assess the spatiotemporal trafficking and distribution of Ag. DCs were pulsed in vitro with 50 μg/ml of Qdot655-coupled F(ab′)2 anti-mouse IgG for 1 h and washed twice before footpad injections with 2.5 × 106 pulsed DCs into C57BL6 recipient animals. PLNs were collected after 18 and 48 h, and thick sections of 200–300 μM were prepared for immunostaining followed by spinning disc confocal microscopy analysis. As expected, no Ag was found in the PLNs collected from the recipient control mice after injection with PBS (Fig. 1B). Eighteen hours following s.c. injection of Ag-DCs, Ag-DCs (Ag colocalized with CD11c+ DCs) were found in the paracortical T cell–rich area but absent in the cortex (Fig. 1C). At 48 h, we still observed some Ag-DCs in the paracortex, but we remarkably also highlight a relocalization of Ag-DCs consisting in an accumulation within the 50-μm IR region between the paracortex and cortex and also within the B cell zone (Fig. 1D). We next quantified this redistribution of fluorescent Ag-DCs by counting the number of Ag-DCs within the paracortical, IR, and cortical regions of PLNs collected from PBS and Ag-DCs–injected mice. Our results indicate in two independent experiments (n = 1, left panel; n = 2, right panel), despite differences in total number of Ag-DCs within the LN (due probably to the differential migratory ability of Ag-DCs in these assays), a dynamic relocalization of Ag-DCs from the paracortex at 18 h to the cortex at 48 h following s.c. injection. This relocalization consists of a dramatic increase and a strategic positioning of Ag-DCs within the LN B cell zone (Fig. 1E).

Taken together, these results indicate that DCs are important cellular carriers for Ag transport and trafficking from the periphery to the LN B cell zone.

In vivo and in vitro B cell activation by Ag-DCs

The repositioning of Ag-DCs within the B cell zone of the LN at 48 h after injection suggests a potential functional interaction of DCs with B cells that may allow Ag transfer and subsequent B cell activation that we decided to investigate. For this, we used the well-studied protein Ag HEL, which is commonly used to assess Ag-specific B cell responses in vitro and in vivo. DCs were left untreated or pulsed in vitro with 1 mg/ml of HEL as used previously (11) for 1 h and washed twice with PBS before footpad injections into C57BL6 recipient animals. Mice were also injected into the footpad with PBS as the control group. PLNs were collected at day 3 after immunization, and single cell suspensions were prepared for immunostaining to assess B cell activation and differentiation by flow cytometry. Seventy-two hours following s.c. injection, DCs-HEL were able to promote in vivo B cell activation as evidenced by the increased CD86 surface expression in contrast to DCs alone (Fig. 2A, 2B). There was a tendency for an induction of plasmablast differentiation (increased CD138 surface expression) following DCs-HEL injection in some experiments, but it did not reach significance (Fig. 2B). The DC-induced activation of B cells observed at day 3 was not visible at day 1 following s.c. injection (data not shown). These results indicate that DCs are not only carriers of Ag from the periphery to the LN B cell zone but also potential inducers of B cell activation in vivo.

FIGURE 2.
FIGURE 2.

In vivo and in vitro B cell activation by Ag-DCs. DCs were left untreated or pulsed in vitro with HEL before in vivo footpad injections. (A) Representative FACS profiles in gated B220+ cells 72 h following mouse immunization with DCs-HEL. n = 4 mice per condition pooled from n = 3 independent experiments. (B) Quantification in percentage of surface expression after FACS analysis. Bar graphs indicate mean ± SEM; *p = 0.0180 (two-way ANOVA with multiple comparisons corrected with the Dunnett method). (C) Representative dot plots in gated B220+ cells 18 h after stimulation of WT and MD4 B cells with HEL or LPS. (D) Quantification in percentage of surface expression after FACS analysis. n = 3 independent experiments. Bar graphs indicate mean ± SEM; *p = 0.0125, ***p = 0.0006 (two-way ANOVA with multiple comparisons corrected with the Sidak method). (E) Representative FACS profiles in gated B220+ cells 18 h after coculture of WT and MD4 B cells with DC, DCs-HEL, or stimulation with HEL. n = 3 independent experiments. (F) Quantification in percentage of surface expression after FACS analysis. n = 5 independent experiments for MD4. Bar graphs indicate mean ± SEM; ****p < 0.0001 (two-way ANOVA with multiple comparisons corrected with the Dunnett method).

To investigate whether Ag-DCs could also induce B cell activation in vitro as it has been analyzed in vivo, we decided to use an in vitro system of coculture of Ag-pulsed DCs (DCs-HEL) with anti-HEL transgenic MD4 B cells (28). We first checked the Ag-specific response of MD4 B cells to HEL in comparison with WT B cells. Both WT and MD4 B cells were left untreated or stimulated for 18 h with 100 μg/ml of soluble HEL or with 1 μg/ml of LPS. After immunostaining, we have analyzed CD69 and CD86 surface expression. Whereas LPS seems to be a potent activator of both WT and MD4 B cells by inducing a significant increase of CD69 and CD86 surface expression, only MD4 B cells that carry anti-HEL BCR respond to HEL Ag by upregulating CD69 and CD86 surface expression (Fig. 2C, 2D).

We then wanted to recapitulate the potential ability of DCs-HEL to activate B cells in coculture in vitro as seen in vivo (Fig. 2E, 2F). DCs were left untreated or pulsed in vitro with 1 mg/ml of HEL for 3 h and washed twice with cold PBS before coculture with B cells. Both WT and MD4 B cells were left untreated, stimulated with 100 μg/ml of soluble HEL, and cocultured with nonpulsed DCs or with DCs-HEL for 18 h. After immunostaining, we analyzed by flow cytometry both B cell activation and plasmablast differentiation. As expected, stimulation with HEL induces a potent activation of anti-HEL MD4 B cells but not WT B cells as indicated by the increased CD69 and CD86 surface expression. In comparison with nonpulsed DCs, DCs-HEL were able to strongly and specifically activate MD4 B cells but not WT B cells. Thus, Ag-carrying DCs are potent inducers of B cell activation both in vivo and in vitro, indicating potential Ag transfer by DCs to B cells.

Extracellular release of Ag by DCs regurgitation induces early B cell activation associated with NF-κB/cRel nuclear mobilization

The observed B cell activation induced in coculture in vitro could result from Ag transfer either by cell-to-cell contact or through an extracellular release by DC regurgitation as described previously (9). To test this hypothesis, DCs were left untreated or pulsed in vitro with 1 mg/ml of HEL (DCs-HEL) for 1–3 h, washed twice with cold PBS, and then chased for 18 h at 37°C. SpN were collected from nonpulsed (DC SpN) and HEL-pulsed (DC-HEL SpN) DCs, filtered, and then applied to negatively purified splenic MD4 and WT B cells for 18 h. B cell activation and plasmablast differentiation were then analyzed by flow cytometry. Interestingly, in comparison with the treatment with DC SpN, DC-HEL SpN was able to induce a significant increase of CD69 surface expression on MD4 B cells carrying anti-HEL BCR (Fig. 3A, 3B, right panel) but not on WT B cells (Fig. 3A, 3B, left panel). These data indicate that Ag-specific B cells carrying anti-HEL BCR undergo early activation upon contact with the extracellular medium of chased DCs-HEL.

FIGURE 3.
FIGURE 3.

Extracellular release of Ag by DC regurgitation induces early B cell activation associated with NF-κB/cRel nuclear mobilization. DCs were left untreated or pulsed in vitro with HEL and then chased for 18 h before collection of the extracellular regurgitation medium to assess B cell activation. (A) Representative dot plots in gated B220+ cells 18 h after stimulation of WT and MD4 B cells with HEL, DC SpN, or with DC-HEL SpN. (B) Quantification in percentage of surface expression after FACS analysis. n = 3 independent experiments. Bar graphs indicate mean ± SEM; *p = 0.0230, **p = 0.0064 (two-way ANOVA with multiple comparisons corrected with the Sidak method). (C) Representative WB on DC total lysates made after 3 h of capture or 18 of chase (left panel) and on DC SpNs collected after 18 of chase (left panel) for HEL expression across three independent experiments. (D) Representative WB on nuclear protein extracts made from both WT and MD4 B cells for cumulative nuclear expression of cRel across three experiments. (E) Quantification of cRel after normalization to loading control H3 across three experiments. Bar graphs indicate mean ± SEM; *p < 0.020 (unpaired Student t test with equal SD).

We then aimed to characterize the presence of HEL Ag in the DC-HEL SpN, which may support the specific activation of anti-HEL MD4 B cells by the DC regurgitation medium in vitro. First, DC SpN were collected after 18 h of chase of nonpulsed (DC SpN) and HEL-pulsed (DC-HEL SpN) DCs, filtered, and then subjected to M/S analysis. Protein quantification by Bradford assay in both DC and DC-HEL SpNs revealed no difference between these two conditions in total protein levels (data not shown). The M/S analysis revealed the presence of HEL exclusively in DC-HEL SpN (data not shown). Second, using an anti-HEL Ab, we analyzed by WB the distribution of HEL from DC capture, chase, and until its extracellular release in the medium. Our results indicate the presence of HEL (with two bands at 14 and 27 kDa) in DC total lysates following 3 h of pulse (Fig. 3C, left panel), which decreases dramatically after 18 h of chase (Fig. 3C, left panel) due most likely to degradation/processing and/or potential extracellular release by regurgitation. To test this hypothesis, we have then analyzed also by WB the presence or not of HEL in DC SpNs collected after 18 h of chase. Interestingly, WB analysis revealed the presence of native HEL with a band visible at 27 kDa in DC-HEL SpN and not in DC SpN (Fig. 3C, right panel). These results indicate that DCs-HEL are able to release part of the internalized Ag after capture in native form by regurgitation in the medium, supporting the observed Ag-specific B cell stimulatory effect.

To assess whether the Ag released in the SpN could result from damaged DCs undergoing necrosis after 18 of chase, we collected the 18-h chased DCs to analyze cell death after PI staining by flow cytometry. Our analysis indicates that the majority of the 18-h chased HEL prepulsed DCs were within the “living cells” gate (Supplemental Fig. 1A) of CD11c+ DCs (Supplemental Fig. 1B) and did not show any significant cell death by necrosis after PI staining (Supplemental Fig. 1C). These results indicate that the Ag released in the SpN by regurgitation does not result from damaged or disrupted cells dying by necrosis.

To estimate the concentration of HEL released in the DC SpN, we analyzed by WB in parallel to serial doses of HEL (0.01, 0.05, 0.10, and 0.5 μg), DC, and DC-HEL SpN. Our WB results indicate the presence of HEL (band at 14 kDa) detected in DC-HEL SpN corresponding to an amount ranging from 0.01 up to 0.05 μg (Supplemental Fig. 2A). We then established a standard line with different doses of HEL (Supplemental Fig. 2B, Supplemental Table I) that allowed us to estimate the HEL concentration in DC-HEL SpN to be around 6 μg/ml (Supplemental Table II). These results indicate that DCs-HEL are able to release a subfraction of the internalized Ag after capture by macropinocytosis in a nondegraded form by regurgitation in the extracellular medium.

We next aimed to investigate the potential intracellular mechanism underlying the B cell activation induced by DCs through extracellular release of Ag. We decided to analyze NF-κB, in particular cRel nuclear mobilization, as it is described to be an important early B cell event following BCR stimulation (22, 23). For this, both MD4 and WT B cells were left untreated or stimulated with 10 μg/ml of anti-IgM F(ab′)2 Ab as a positive control, 100 μg/ml of soluble HEL, 1 ml of DC SpN, or with 1 ml of DC-HEL SpN for 18 h at 37°C. After stimulation, cells were harvested, and cytosolic and nuclear extracts were prepared as previously described (19). Nuclear extracts were analyzed by WB to assess nuclear mobilization of NF-κB/cRel (Fig. 3D, 3E). As expected, in both WT and MD4 B cells, BCR stimulation with an anti-IgM F(ab′)2 Ab for 18 h led to a significant nuclear accumulation of NF-κB/cRel, as previously reported (22). Interestingly, stimulation with HEL as a protein Ag induced also a strong nuclear accumulation of NF-κB/cRel in MD4 B cells but not in WT cells. Strikingly, DC-HEL SpN but not DC SpN induced a potent nuclear accumulation of NF-κB/cRel specifically and exclusively in anti-HEL MD4 B cells (Fig. 3D, 3E). These results indicate that Ag release by DC regurgitation induces an early B cell activation that is BCR driven and associated at the intracellular level with sustained nuclear accumulation of NF-κB/cRel.

Drug inhibition of cRel affects both Ag-dependent early B cell and T cell activation

We then wanted to assess the potential involvement of NF-κB/cRel in the early B cell activation induced by Ag delivered by DCs in vitro. For this, we decided to block cRel expression and activation by chemical inhibition. We have used PTXF, which is a xanthine derivative that has been approved by the U.S. Food and Drug Administration for clinical use and recently reported to be able to cause selective degradation of cRel without affecting p65 in T cells (29). First, we wanted to assess whether cell treatment with PTXF could also cause specific inhibition or degradation of cRel in B cells. MD4 B cells were left untreated or pretreated with 500 μg/ml of PTXF for 15 min before being stimulated with 100 μg/ml of HEL or with 1 μg/ml of LPS for 18 h at 37°C. After stimulation, whole extracts were prepared to be analyzed by WB by using anti–cRel Ab. Stimulation of both BCR with HEL and TLR4 with LPS for 18 h led to a significant nuclear accumulation of NF-κB/cRel. Interestingly, pretreatment of B cells with PTXF strongly inhibited cRel nuclear expression induced specifically upon BCR stimulation with HEL. Strikingly, pretreatment with PTXF did not affect cRel induction upon TLR4 stimulation with LPS (Fig. 4A, 4B). These results indicate that PTXF is a potent and highly specific inhibitor of NF-κB/cRel induction triggered upon BCR stimulation in B cells.

FIGURE 4.
FIGURE 4.

Drug inhibition of cRel affects both Ag-dependent early B cell and T cell activation. MD4 B cells were purified and stimulated for 18 h as indicated for the different conditions in the absence or presence of 500 μg/ml PTXF. (A) Representative WB on nuclear protein extracts made from MD4 B cells for cumulative nuclear expression of cRel across three experiments. (B) Quantification of cRel after normalization to loading control GAPDH across three experiments. Bar graphs indicate mean ± SEM; *p = 0.0163 (unpaired Student t test with equal SD). (C) Representative dot plots in gated B220+ cells 18 h after stimulation of MD4 B cells with LPS, HEL, or with DC-HEL in the absence or the presence of PTXF. (D) Quantification in percentage of surface expression after FACS analysis. n = 3 independent experiments. Bar graphs indicate mean ± SEM; **p < 0.010 (unpaired Student t test with equal SD). (E) Purified spleen CD3+ T cells were stimulated for 18 h with anti-CD3/CD28 in the presence of 500 μg/ml PTXF or H2O and analyzed by FACS. Data are shown as dot plots in the gated B220+ cells. (F) Quantification in percentage of surface expression after FACS analysis. n = 3 independent experiments. Bar graphs indicate mean ± SEM; **p = 0.0013 (unpaired Student t test with equal SD).

We next evaluated the implication of NF-κB/cRel in the early B cell activation induced upon DCs Ag release by blocking cRel with PTXF. MD4 B cells were left untreated or pretreated with 500 μg/ml of PTXF for 15 min before being stimulated with 100 μg/ml of HEL, 1 μg/ml of LPS, or with 1 ml of DC-HEL SpN for 18 h at 37°C. After stimulation, cells were collected for immunostaining followed by flow cytometry analysis of CD69 surface expression, a marker of early B cell activation. As expected and in line with the results of Figs. 2 and 3, stimulation with HEL, LPS, or with DC-HEL SpN induced early B cell activation, as indicated by increased CD69 surface expression. Although cell treatment with PTXF blocked significantly and specifically the increased CD69 surface expression induced by HEL or DC-HEL SpN, no impact of such treatment was observed on LPS-induced early B cell activation (Fig. 4C, 4D). These results suggest that inhibition of NF-κB/cRel blocks the early B cell activation specifically induced by soluble Ag and Ag delivered by DCs but not through TLR4 stimulation with LPS.

To extend the specificity of the blocking action of PTXF on CD69 surface expression seen in B cells, we then examined the effect on this drug on T cell CD69 surface expression. Purified splenic CD3+ T cells were left untreated or stimulated with 5 μg/ml of anti-CD3/CD28 Abs in the absence or presence of 500 μg/ml of PTXF (for 15 min before) for 18 h at 37°C. After stimulation, cells were collected for immunostaining followed by flow cytometry analysis of CD69 surface expression. As expected, costimulation of T cells through TCR/CD28 led to a significant increase of CD69 surface expression. Interestingly, treatment with PTXF also inhibited strongly the T cell CD69 surface expression induced upon TCR/CD28 costimulation (Fig. 4E, 4F). These results highlight that inhibition of NF-κB/cRel blocks both the early B cell and T cell activation specifically induced upon Ag receptor stimulation.

Discussion

In this study, we report that DCs are an important cell transporter of Ag from the periphery to the LN B cell zone and also potent inducers of B cell activation and differentiation both in vivo and in vitro. We demonstrated that extracellular release of Ag by DC regurgitation induces early B cell activation, which is BCR driven and mechanistically dependent on the nuclear mobilization of the transcription factor NF-κB/cRel.

B cells can acquire small soluble Ags diffusing directly from the lymph into the follicle, whereas large size particulate Ags can be delivered to B cells by the local LN B cell Ag cell carriers, such as FDCs, subcapsular sinus macrophages, follicular reticular cells conduits, or resident DCs (3, 4). However, not all Ags can be opsonized by LN-resident macrophages and FDCs to be presented to B cells. Many pathogen-associated Ags are likely captured from peripheral tissues in which conventional DCs are the “best option” as they are potent migratory capturing cells, in contrast to other cells, for trafficking Ag from the periphery into draining LNs to present it to both T and B cells. To access the B cell compartment for particulate Ags accumulated in peripheral tissues during infection, based on our data, we propose DCs as a major cell transporter of Ag from the periphery to the LN B cell zone.

We initially focused on the analysis by imaging of the Ag transport and trafficking by DCs from the periphery to the LN. Previous studies have shown by in vivo imaging the ability of DCs to transport Ags from the periphery to the paracortical zone of the LN, leading to the Ag presentation to T cells (30–32). However, concerning the Ag presentation to B cells in vivo, the kinetics and the geographic LN distribution of Ag potentially transported by DCs from the periphery has been incompletely visualized. Eighteen hours following s.c. injection of DCs pulsed in vitro with fluorescent Ag, we could visualize the presence of the Ag captured by DCs in vitro in the LN within the paracortical T cell–rich area and colocalized with CD11c+ DCs. At 48 h, we then observed a dynamic relocalization of Ag and Ag-DCs illustrated by an accumulation within the IR region followed by repositioning within the LN cortical B cell zone. Although low levels of nonspecific staining to circulating IgGs cannot be excluded, most of the Ag fluorescence was cell associated as indicated by the white arrows at higher magnification in Fig. 1C and 1D. Quantification of our image data indicates more Ag-DCs in total at 48 h. The observed accumulation and the strategic positioning of Ag-DCs within the cortex at 48 h suggests a relocalization from the paracortex at 18 h to the cortex at 48 h after injection. As we still see Ag-DCs within the paracortex at 48 h, this may suggest also de novo infiltration of Ag-DCs migrating from the periphery to the LN, suggesting that both possibilities remain open.

In LNs, the majority of DCs reside in the T cell–rich area (33), raising the question of how naive B cells might access DC-presented Ags in vivo. In this study, our results might suggest that some T cell–derived signals such as CD40L could, upon interaction with CD40 on DCs, trigger overexpression of CXCR5 (34), the receptor for CXCL13, allowing their homing from the T cell–rich paracortex to the B cell zone. The migration of Ag-carrying DCs to the B cell zone could most likely be driven by the potent B cell chemoattractant CXCL-13 (35). Previous studies described an established interaction between DCs and B cells observed in vivo within the extrafollicular paracortical T cell area (11, 36, 37). In this study, we reveal a strategic redistribution of Ag-DCs within the cortical B cell region, suggesting a functional interaction of DCs with follicular B cells in vivo.

The strategic repositioning of Ag-DCs within the follicular B cell zone observed at 48 h after injection suggests a potential functional interaction of DCs with B cells, leading to Ag transfer and B cell activation by DCs. Indeed, 72 h after s.c. delivery of Ag-pulsed DCs, we observed in vivo B cell activation indicated by increased CD86 surface expression. In a previous study using transgenic B cell transfer and s.c. delivery of DCs-HEL, it has been shown by in vivo imaging that Ag-carrying DCs migrating from the periphery can activate the B cells entering the LN via high endothelial venules (HEV) 12 h after B cell transfer (11). This landmark study highlighted an extrafollicular activation of immigrant B cells by peripheral Ag-bearing DCs within the peri-HEV paracortex T cell region. In this study, we observed a potential activation in vivo of LN-resident follicular B cells by peripheral DCs 72 h after s.c. injection, resulting from a strategic positioning of Ag-DCs within the cortical B cell zone at 48 h. Taken together, our results may suggest that Ag-DCs migrating from the periphery can activate early the immigrant extrafollicular LN B cells entering the LN via HEV within the paracortex and later the resident follicular B cells within the cortical B cell zone.

By using an in vitro coculture system of DCs-HEL with anti-HEL transgenic MD4 B cells, we were able to recapitulate in vitro the ability of DCs-HEL to activate B cells as seen in vivo. This result suggests that the B cell activation induced in vivo following s.c. injection of DCs-HEL could result from the observed potential interaction of DCs with B cells in the cortical follicular region without excluding the involvement of other cell IRs within the LN such as T cells, resident DCs, or FDCs.

In vivo imaging of B cells in intact secondary lymphoid organs had provided major insights about B cell contacts with APCs (11), but how these interactions promote Ag transfer and acquisition by B cells were still incompletely understood. This transfer can be mediated by cell–cell contact, allowing B cells to extract surface-tethered Ags through two nonexclusive mechanisms. The first one involves enzymatic degradation through local secretion of lysosomes recruited at the immunological synapse (13, 14). The second one relies on the use of myosin IIA–mediated mechanical forces that trigger invagination of Ag-containing membranes and internalization (15–17). In this study, we report another potential mode of Ag acquisition by B cells at the immunological synapse and highlight a novel extracellular mechanism of B cell activation by DCs through regurgitation of Ag. This Ag delivery from late endosomal compartments was named regurgitation in our previous study to distinguish it from the classical endosomal recycling pathway (9).

DC regurgitation of Ag may play a role in the B cell activation induced in vivo within the LN B cell zone. Interestingly, we have previously uncovered that Ag released by DCs after macropinocytosis in vitro was unexpectedly enhanced by CXCL-13 (9). Taken together, we can then propose that CXCL-13 can both drive the homing of Ag-carrying DCs to the B cell zone as reported previously (38, 39) and promote subsequent Ag regurgitation by DCs to target B cells.

Our findings indicate that the B cell stimulatory activity elicited in the pulsed DCs SpN is due to the presence of HEL. Our results indicate that the release of Ag would not result from damaged 18-h chased DCs undergoing cell necrosis (Supplemental Fig. 1). Ag can be released by DCs upon regurgitation in a soluble form that was able to induce BCR cross-linking, leading to the transduction of activation signal for B cells. Otherwise, DCs may also deliver native Ag within extracellular vesicles (EVs), such as microvesicles or exosomes, resulting in a potent stimulatory action on B cells. In our previous study, it was reported that regurgitation was regulated by the small GTPase Rab27 (9), which is known to also control exosome delivery (40). The regurgitated Ag [F(ab′)2 anti-mouse IgG] used in that study was not associated with the exosomal fraction (9). However, concerning the HEL model Ag used in this current study and the in vivo situation, we cannot exclude that Ag released by DC regurgitation could be delivered into DC EVs, and this remains to be investigated.

Despite the small amounts of native Ag released in the medium by DC regurgitation (Supplemental Fig. 2A, Supplemental Table II), this low concentration is sufficient to efficiently induce early B cell activation at the level of both CD69 surface expression and NF-κB/cRel nuclear mobilization (Fig. 3A, 3B, 3D). This would therefore imply that in addition to being delivered potentially within DC EVs, the released Ag could also be associated probably with a costimulatory factor that remains to be explored.

At the intracellular level, canonical NF-κB activation, in particular cRel nuclear mobilization, has been described as an important early B cell event following BCR stimulation (22, 23). In these previous studies, BCR stimulation was triggered by using an anti-IgM F(ab′)2 Ab but not yet reported with a relevant protein Ag. In this study, we show a strong NF-κB/cRel nuclear translocation following B cell stimulation with soluble HEL but also with HEL released by DCs in a BCR-specific manner because this translocation was not seen in WT B cells.

To probe the possible involvement of NF-κB/cRel in the early B cell activation, we have blocked cRel by chemical inhibition with PTXF. We have analyzed the effect of PTXF on cRel inhibition and B cell activation following BCR stimulation with Ag for 18 h as an optimal time point. In this study, we show that cRel inhibition affects specifically the Ag-dependent early B cell activation induced following BCR stimulation with HEL and DC-HEL SpN but not through TLR4 stimulation with microbial LPS. Furtheremore, treatment with PTXF prior to TCR/CD28 stimulation also strongly prevents CD69 surface expression. This could be explained also by cRel inhibition by PTXF as recently described (29). Together, our findings indicate that PTXF is a highly specific inhibitor of the NF-κB/cRel pathway, which is activated in common downstream of BCR and TCR through the CARMA–BCL10–MALT-1 complex (22, 23). Loss of cRel was recently shown to lead to defective generation of the germinal center reaction and impaired T-independent Ab responses in vivo by using B cell–specific cRel-deficient mice (24, 25). As cRel is a transcription factor, our results suggest that CD69 is a potential κB target gene controlled specifically by cRel. Interestingly, cRel was previously reported to control also the gene transcription of IL-21 in T cells (41), a key “T cell help” cytokine produced by activated (CD69+) follicular helper T cells for supporting B cell responses (1, 2). Thus, the NF-κB/cRel member seems to play an important dual role in the control of both Ag-dependent B cell early activation and also T cell help for B cells.

We have previously demonstrated that distinct NF-κB subunits control DC survival, functions, and DC-induced T cell responses (19, 42). Whether cRel or other NF-κB members also play a role in the Ag regurgitation by DCs that impact the direct early B cell activation is not known and remains to be investigated.

DCs are highly crucial for adaptive immunity in infection as they are equipped with both nondegradative and degradative Ag uptake pathways to perform Ag presentation to both B and T cells. The role of cRel in the Ag-dependent early B and T cell activation suggests that drug inhibition of this NF-κB member would represent a promising approach to limit the excessive DC-elicited B/T cell responses in autoimmunity without altering the host humoral defense against Gram-negative bacterial infection.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank the Proteomics Core Facility at Institut Jacques-Monod (notably T. Léger, C. Garcia, and B. Morlet) for the M/S analysis and the Flow Cytometry–Immuno-Biology, Animal, and Photonic Imaging facilities at Institut Cochin for help and advice.

Footnotes

  • ↵2 F.N. and F.O. are co-last authors.

  • This work was supported by the Fondation ARC pour la Recherche sur le Cancer, CNRS, INSERM, and Université Paris-Descartes.

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    Ag-DC
    Ag-loaded DC
    DC
    dendritic cell
    DC-HEL
    HEL-pulsed DC
    EV
    extracellular vesicle
    FDC
    follicular DC
    HEL
    hen egg lysozyme
    HEV
    high endothelial venule
    IR
    intermediate
    LN
    lymph node
    M/S
    mass spectrometry
    PI
    propidium iodide
    PLN
    popliteal LN
    PTXF
    pentoxifylline
    SpN
    supernatant
    WB
    Western blot
    WT
    wild-type.

  • Received April 4, 2019.
  • Accepted May 3, 2020.

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