The germinal center (GC) is divided into a dark zone (DZ) and a light zone (LZ). GC B cells must cycle between these zones to achieve efficient Ab affinity maturation. Follicular dendritic cells (FDCs) are well characterized for their role in supporting B cell Ag encounter in primary follicles and in the GC LZ. However, the properties of stromal cells supporting B cells in the DZ are relatively unexplored. Recent work identified a novel stromal population of Cxcl12-expressing reticular cells (CRCs) in murine GC DZs. In this article, we report that CRCs have diverse morphologies, appearing in open and closed networks, with variable distribution in lymphoid tissue GCs. CRCs are also present in splenic and peripheral lymph node primary follicles. Real-time two-photon microscopy of Peyer’s patch GCs demonstrates B cells moving in close association with CRC processes. CRCs are gp38+ with low to undetectable expression of FDC markers, but CRC-like cells in the DZ are lineage marked, along with FDCs and fibroblastic reticular cells, by CD21-Cre– and Ccl19-Cre–directed fluorescent reporters. In contrast to FDCs, CRCs do not demonstrate dependence on lymphotoxin or TNF for chemokine expression or network morphology. CRC distribution in the DZ does require CXCR4 signaling, which is necessary for GC B cells to access the DZ and likely to interact with CRC processes. Our findings establish CRCs as a major stromal cell type in the GC DZ and suggest that CRCs support critical activities of GC B cells in the DZ niche through Cxcl12 expression and direct cell–cell interactions.
Lymphoid tissue stromal cells are specialized mesenchymal cells that establish and maintain the distinct niches necessary to support effective adaptive immune responses. Lymphoid follicles, the B cell–rich regions of lymphoid organs, are organized around a complex network of follicular stromal cells (1). Many of the follicular stromal cells in primary (nonreactive) follicles produce the chemokine CXCL13 (BLC) and are involved in attracting B cells into this compartment. Follicular dendritic cells (FDCs) are a subset of these CXCL13-expressing stromal cells situated in the central region of the follicle (1). First defined by their ability to capture opsonized Ags, FDCs are now known to highly express complement receptors-1 and -2 (CD35 and CD21, respectively) and Fcγ receptors to support the process of immune complex capture and display to cognate B cells (1–3). FDCs and the broader CXCL13-producing follicular stromal cell network share a dependence on the cytokines lymphotoxin (LT)–α1β2 and TNF for maintenance and function (4–6).
Although FDCs are one of the stromal cell types supporting B cell follicles, fibroblastic reticular cells (FRCs) are mesenchymal stromal cells that support the structure and function of the T zone. FRCs produce the chemokines CCL19 and CCL21 in an LT-dependent manner to guide CCR7-expressing B and T cells into lymph node (LN) and splenic T zones (4, 7, 8). FRCs also promote T cell homeostasis by producing IL-7 (9). In addition, FRCs form a network of conduits in the T zone that transport Ag and facilitate T cell encounter with Ag-bearing DCs (10).
Following Ag exposure, activated B cells proliferate in B cell follicles and form polarized germinal centers (GCs), each with a light zone (LZ) and a dark zone (DZ). The FDCs within GCs upregulate CD21, CD35, Fc receptors, ICAM1, and VCAM1 as well as show increased staining for activated C4 (FDC-M2) and MFG-E8 (milk fat globule epidermal growth factor 8, FDC-M1) relative to FDCs in primary follicles (2). Ag-bearing FDCs are restricted to the LZ, designating this as the site of B cell Ag recognition and selection (11). GC FDCs are also required for GC B cell confinement and viability (12). CXCL13 is present in the GC LZ and plays a role in positioning GC B cells in this region. In contrast, the DZ has little CXCL13 and instead is a source of CXCL12 (SDF1). GC B cell movement from LZ to DZ as well as GC polarization into zones depends on GC B cell expression of the CXCL12 receptor CXCR4 (13). Once in the DZ, GC B cells express higher amounts of activation-induced cytidine deaminase, undergo somatic hypermutation, and are more likely to proliferate before returning to the LZ (1, 14). Recent work has highlighted the importance of the DZ for affinity maturation and GC participation, as these were impaired in CXCR4 knockout GC B cells that could not access the DZ (15).
In contrast to the extensive study of FDCs since their discovery in the 1960s, little is known about the stromal cells in the GC DZ. Ultrastructural studies revealed the presence of stromal cells in the DZ of human tonsil GCs and referred to them as immature FDCs. However, these cells mostly did not capture or display opsonized Ag, lacked the labyrinth-like structure of LZ FDCs, and their relationship to true Ag-capturing FDCs was unclear (16, 17). Lefevre and coworkers (18) described a mAb, found to bind fibrinogen, that stained DZ stromal cells in bovine and ovine GCs. However, fibrinogen was found not to be made locally by the DZ stroma and was thought to have derived from blood or lymph. In recent work, we followed up on the functional evidence that CXCL12 emanates from the DZ (13) to reveal the existence of Cxcl12-expressing reticular cells (CRCs) in mediastinal LN GC DZs after influenza infection (15). Although CRCs have only minimal overlap with reticular fibers in the mediastinal LN, CRCs were so named to represent their netlike morphology, as the term reticular comes from the Latin word for net (1). This work also provided initial evidence that related cells were present in Peyer’s patches (PPs) and within peripheral LN (pLN) primary follicles. However, whether CRCs are a homogeneous population across tissues, what lineage relationships they have to surrounding stroma, and what distinguishing requirements they have for maturation and maintenance were not established.
In this article, we demonstrate that CRC networks have two distinct morphologies and are present in the GC DZs of spleens and pLNs after viral infection. We also find CRCs within the chronic GCs of mesenteric LNs (MLNs) as well as PPs. GC B cells migrate over CRC processes in a similar manner to their migration over FDC processes. Although CRCs are phenotypically distinguishable from FDCs and FRCs, they are likely related in origin based on lineage tracing experiments using CD21-Cre and Ccl19-Cre transgenic mice. Distinct from FDCs, CRCs do not require LT or TNF for short-term maintenance of chemokine expression or morphology. Organization of CRCs into reticular networks, however, depends on CXCR4 function.
Materials and Methods
Mice and chimeras
C57BL/6 (B6) and B6-CD45.1 mice were obtained from The Jackson Laboratory or the National Cancer Institute. B6.Cg-Cxcl12tm2Tng (Cxcl12-GFP) gene-targeted mice were backcrossed to the C57BL/6 background more than seven generations and provided by T. Nagasawa (19). Tg(UBC-GFP)30Scha/J (UBI-GFP) transgenic mice were backcrossed to the C57BL/6 background for more than eight generations and were from The Jackson Laboratory (20). B6.Tg(Cr2-Cre)3Cgn (CD21-Cre) BAC-transgenic mice were fully backcrossed to C57BL/6 and provided by K. Rajewsky (Immune Disease Institute, Boston, MA) (21). B6.Cg-Gt(ROSA)26Sortm6(CAG-Zsgreen1)Hze/J (R26-ZsGreen) mice have a CAG promoter, a floxed stop sequence, and ZsGreen1 knocked into the Gt(ROSA)26Sor locus and were from The Jackson Laboratory. C57BL/6N-Tg(Ccl19-cre)489Biat (Ccl19-Cre) are BAC-transgenic mice (8). Gt(ROSA)26Sortm1(EYFP)Cos (R26-EYFP) transgenic mice express EYFP from the Gt(ROSA)26Sor locus after Cre-mediated deletion of floxed stop cassette (22) and were provided by L. Lanier. Tg(CAG-ECFP)CK6Nagy (CFP) transgenic mice were backcrossed to the C57BL/6 background more than five generations and were from The Jackson Laboratory. Tg(IghelMD4)4Ccg (MD4) transgenic mice were fully backcrossed to C57BL/6 and were from an internal colony.
To make bone marrow (BM) chimeras, UBI-GFP mice were treated i.p. with 500 μg anti-Thy1.2 (clone 30H12) before being lethally irradiated and reconstituted for ≥8 wk with wild-type (WT) CD45.1 BM. CD21-Cre mice were crossed to R26-ZsGreen mice and lethally irradiated and reconstituted for ≥8 wk with WT CD45.1 BM as described previously (15).
Animals were housed in a specific pathogen–free environment in the Laboratory Animal Research Center at the University of California, San Francisco, and all experiments conformed to ethical principles and guidelines approved by the University of California, San Francisco, Institutional Animal Care and Use Committee.
Infections and immunizations
Mice were infected with acute lymphocytic choriomeningitis virus (LCMV)–Armstrong i.v. at 2.5 × 105 PFU and analyzed at day 15 for GCs in pLNs and spleen (23). For induction of spleen GCs, mice were immunized i.p. with 2 × 108 SRBCs (Colorado Serum Company) on day 0 and day 5 and were analyzed on days 10–12. For induction of pLN GCs, animals were immunized s.c. at the shoulders, flanks, and above the tail with SRBCs on day 0 and day 5. Draining pLNs (axillary, brachial, and inguinal) were analyzed on days 10–12.
Treatments and transfers
For LTβR and TNFR signaling blockade, Cxcl12-GFP mice were immunized i.p. or s.c. with SRBCs on day 0 and day 5 and on day 10 treated i.v. with 100 μl each of 1 mg/ml mLTβR-huIgG1 (LTβR-Fc, provided by J. Browning) and 1 mg/ml TNFR55-huIgG1 (TNFR-Fc, provided by J. Browning) or saline. Tissues were analyzed 4 d later.
For CXCR4 inhibitor treatment, Cxcl12-GFP mice were immunized with SRBCs i.p. on day 0 and day 5. On day 8, Alzet osmotic pumps (1-d duration, 8.4 μl/h pumping rate; Model 2001D; Durect Corporation) loaded with saline or 5 mg/ml of the CXCR4 antagonist 4F-benzoyl-TE14011 (24) in saline were implanted dorsally s.c. according to the manufacturer’s instructions. As analgesics, buprenorphine (0.05–0.1 mg/kg; Sigma-Aldrich) was given i.p. before and after surgery, carprofen (5 mg/kg; Pfizer Animal Health) was given i.p. before surgery, and bupivacaine (100 μl of 0.25%; Hospira) was given topically during surgery. Tissues were analyzed 12 or 24 h later.
For two-photon laser scanning microscopy (TPLSM) of intact GCs from pLNs and MLNs, Cxcl1225, 26). On day 9, mice were transferred with 1.5 × 108 CFP transgenic B cells purified from donor spleens using anti-CD43 microbeads (Miltenyi Biotec) i.v., as previously described (26). PLNs and MLNs were mounted and imaged 24 h later.
For TPLSM of PPs, Cxcl12-GFP mice were crossed to MD4 mice and transferred with 20% CFP transgenic B cells and 80% CD45.1 WT B cells. B cells were purified from donor spleens as above, and 1.2 × 107 total B cells were injected i.v. into Cxcl12-GFP MD4 mice. At 2 wk later the mice were injected i.v. with 2 × 10726). The experiment was repeated as above with transfer of 5% CFP transgenic B cells for 4 wk and 10% CFP transgenic B cells for 2 wk, with similar results. PPs were mounted and imaged 24 h later.
Confocal microscopy was performed as described previously, with some modifications (15). Tissues were fixed in 4% paraformaldehyde in PBS for 2 h at 4°C, washed three times for 10 min in PBS, then moved to 30% sucrose in PBS overnight. Tissues were flash frozen in TAK tissue-mounting media the following day, and 30-μM sections were cut and then dried for 1 h prior to staining. Sections were rehydrated in PBS with 1% BSA for 10 min and then blocked for 1 h at room temperature, stained in primary Ab overnight at 4°C, and stained for subsequent steps for 2 h at room temperature, all in PBS with 2% mouse serum, 0.1% BSA, 0.3% Triton X-100, and 0.1% NaN3.
For gp38 staining, LNs were fixed in 4% paraformaldehyde in PBS overnight at 4°C, washed three times in PBS, then moved to 20% sucrose in PBS overnight. Sections were processed as above except for a rinse in PBS and peroxidase quench in PBS with 0.045% H2O2 for 15 min prior to blocking for 30 min. Sections were then stained with primary Ab for 1 h at room temperature and then with streptavidin–HRP (Jackson ImmunoResearch) for 30 min, followed by treatment with the Tyramide Signal Amplification Biotin System tyramide staining kit (PerkinElmer) according to the manufacturer's instructions. Sections were then stained with remaining secondary Abs for 1 h at room temperature. Slides were mounted with Fluoromount-G (Southern Biotech), and images were taken with a Leica SP5 inverted microscope with ×40 and ×63 oil immersion objectives. Images were analyzed and processed with Imaris software, and the statistics reported are average values with variability represented as SEM.
Abs, immunofluorescence, and flow cytometry
For flow cytometry, single-cell suspensions were generated and stained as previously described (26
Explant pLNs, MLNs, and PPs were prepared for TPLSM as previously described for explant pLNs (26), except that PPs were mounted with the serosal side face-up. PPs were stabilized in a customized plastic coverslip window with Vetbond tissue glue (3M) to counter the peristaltic motion of the small intestine. The temperature at the PPs during and at the end of several imaging sessions was measured using a dual-temperature controller (TC-344B; Warner Instruments) equipped with a CC-28 cable containing a bead terminator and was found to remain between 36 and 37°C.
Images were acquired with ZEN2012-Black Edition (Carl Zeiss) using a 7MP two-photon microscope (Carl Zeiss) equipped with a Chameleon laser (Coherent). For video acquisition from MLNs and pLNs, a series of planes of 0.5 μm (MLN) or 1 μm (pLN and primary follicle pLN) Z-spacing spanning a depth of 190–260 μm were collected. Each XY plane spans 283.40 μm × 283.40 μm at a resolution of 0.55 μm per pixel (MLN, primary pLN) or 327.00 μm × 327.00 μm at a resolution of 0.64 μm per pixel (pLN). Some images have been cropped in the XY plane for optimal visualization. The excitation wavelength was 920 nm. For video acquisition from PPs, a series of planes of 3 μm Z-spacing spanning a depth of 50–100 μm were collected every 15–30 s. Each XY plane spans 425.10 μm × 425.10 μm at a resolution of 0.83 μm per pixel or 212.55 μm × 212.55 μm at a resolution of 0.42 μm per pixel. Excitation wavelength was 870 nm. For all TPLSM imaging, emission filters were <452 nm for second harmonic, 460–480 nm for CFP, 500–550 nm for GFP, and 570–640 nm for CMTMR. Videos were made and analyzed with Imaris 7.4.2 364 (Bitplane).
To track cells, surface seed points were created and tracked over time. Tracks were manually examined and verified. Data from cells that could be tracked for ≥10 min were used for analysis. Tracking data were analyzed in Microsoft Excel with a custom macro written in Microsoft Visual Basic for Applications, as previously described (26). Movies were adjusted for tissue drift in Imaris 7.4.2 364 (Bitplane), and annotation and final compilation of videos were performed with iMovie (Apple). Video files were saved as .mov (Quicktime).
CRCs populate the GC DZ niche with fine, irregular networks
To investigate the distinguishing features of GC DZ stroma across tissues, we infected Cxcl12-GFP reporter mice with LCMV-Armstrong and assessed the properties of the GFP-expressing CRCs. Using confocal microscopy, we identified extensive CRC networks in splenic and pLN (inguinal, axillary, and brachial) GC DZs located opposite the LZ CD35+ FDCs (Fig. 1A, 1B). Chronic GCs in PPs and MLNs also contained CRC networks in the DZ (Fig. 1C), consistent with previous findings in PPs (15). Blood vessels running through the GC were also GFP+, likely indicating blood endothelial cells expressing Cxcl12 (Fig. 1B) (15, 27).
Previous work (15) established that DZ CRC networks are distinguishable from FDCs and FRCs by their location and distinct morphology. Instead of the thick processes and consistent patterning of T zone FRCs, CRCs have fine, disorganized processes more similar to those of FDCs. However, CRC networks are far less dense than FDC networks. CRC processes are so thin and dispersed that 30-μm stack confocal microscopy was required to visualize the networks. We identified surprising variability in CRC network structure when we imaged splenic and pLN GCs of mice responding to LCMV (Fig. 1D). In pLNs, the DZ CRCs formed mostly open mesh structures (Fig. 1B, 1D) similar to those observed in influenza-induced mediastinal LN GCs (15). We define open networks as having processes that extend into the LZ FDC network with no clear boundary and that become continuous with the FRC-like stroma at the T zone proximal edge of the GC at three or greater points. In the spleen, however, one-fifth of the GCs contained CRCs with a closed mesh structure (Fig. 1A, 1D). Closed networks are bounded by a continuous perimeter of Cxcl12-GFP+ processes and connect to the FRC-like stroma at the T zone proximal edge of the GC at fewer than three points. About one-fifth of GCs in both tissues contained both types of network (Fig. 1D). CRCs in MLN GCs and PP GCs formed open networks that frequently extended throughout the DZ (Fig. 1C).
Unlike the continuous FDC networks that fill the LZ, CRC networks frequently had an asymmetric distribution in the DZ. Of >27 single 30-μm z-stack views of GCs from three mice, more than half of spleen GC DZs and almost one-third of pLN GC DZs appeared only partially populated by CRCs (Fig. 1A, 1B). Although tingible body macrophages (TBMs) are present in GCs and do displace stromal processes, these areas were much larger than the size of a TBM and they stained for BCL6+ GC B cells (Fig. 1A, 1B). In the spleen, almost a tenth of the 30-μm image stacks had no detectable CRC network in the GC (Fig. 1D). However, analysis of sequential sections of several splenic GCs revealed they all had CRCs in at least one view.
To further investigate the extent of DZ occupancy by CRCs without the extrapolation required by sections, we used TPLSM to image CRC networks in MLN GCs (Fig. 1E, Supplemental Video 1) and pLN GCs (Supplemental Fig. 1A, Supplemental Video 2) of intact LNs. SRBC-immunized Cxcl12-GFP mice were treated with PE–immune complex to label FDCs (TBMs are also strongly labeled) (25, 26). At 1 d before analysis, mice also received CFP transgenic B cells, which populated the follicle and outlined the GC. MLN GCs were frequently more proximal to the capsule than were pLN GCs, enabling higher resolution imaging of the DZ, which is orientated distal to the capsule in the GC. In agreement with our section data, we observed CRC networks in separate open and closed structures emerging from the outer edges of the DZ and leaving large areas of DZ unoccupied by detectable CRCs. The thicker three-dimensional view also revealed that each distinct network contained one or more CRC cell bodies, depending on the network size, and many of these cell bodies appeared bilobed.
To assess whether the CRC asymmetry in the DZ was a result of CRCs sharing the DZ niche with another stromal cell type or of CRCs having variable Cxcl12 expression, we analyzed pLNs and spleen from UBI-GFP mice reconstituted with WT BM and infected with LCMV. In these mice, all stromal cells express GFP. Surprisingly, we observed that almost nine-tenths of the pLN and splenic GC DZs had CRC-like networks throughout the DZ with no large areas of undetectable stroma (Fig. 1F, Supplemental Fig. 1B). We saw similar results with SRBC-immunized mice (Supplemental Fig. 1C). These data suggest that although CRCs are a major DZ stromal cell type, GCs may contain additional DZ stromal cells that lack detectable Cxcl12-GFP expression.
Because CRCs in splenic DZs demonstrated more variability than did CRCs in pLN DZs, we expanded on our previous investigation of CRCs in pLN primary follicles (15) to examine CRCs in splenic primary follicles. In unimmunized Cxcl12-GFP pLNs, CRC networks extended along the T zone proximal side of the follicle with similar morphology to that of pLN DZ CRC networks (Fig. 1G), as previously shown (15). In contrast, CRC networks in splenic primary follicles were less extensive than the networks in splenic GC DZs (Fig. 1H). Although they occasionally appeared to consist of only one cell in a single 30-μm z-stack, splenic primary CRCs still formed small open and closed networks in the T zone proximal region of the follicle. CRCs were detectable with a similar average frequency in pLN primary follicles (96.2% ± 3.8, n = 2), as in LCMV-induced pLN GCs (100% ± 0, n = 3), and in splenic primary follicles (87.1% ± 2.0, n = 3), as in splenic GCs (89.8% ± 3.3, n = 3). We also assessed the three-dimensional organization of pLN primary follicle CRCs using TPSLM analysis of intact pLNs as described above for pLN DZ CRCs (Supplemental Fig. 1D, Supplemental Video 2). On the basis of their shared Cxcl12 expression, morphology, and location, our current work and previous data (15) support the conclusion that both pLN and splenic GC CRCs arise from primary follicle CRCs.
Movement dynamics of GC B cells in association with DZ CRCs
We next investigated whether cell–cell interactions might be important to CRC support of GC B cells in the DZ by determining if GC B cells interact with CRC processes as they move through the DZ. We attempted to visualize the interaction using TPLSM on explanted pLNs (28), but the DZ in LN GCs is orientated distal to the capsule and the fine CRC processes were too deep in the tissue to be imaged using laser intensities that preserved cell viability. In PPs, GCs form with the DZ proximal to the serosal surface and with the LZ embedded deeper in the tissue facing the subepithelial dome (29). To take advantage of this superficial positioning of the DZ, we developed an approach to label a fraction of the B cells within PP GCs. We found that when a mixture of 20% CFP transgenic B cells and 80% WT B cells was transferred into hen egg lysozyme–specific (MD4) Ig-transgenic mice for 2 wk, the host GCs became dominated by the transferred B cells and ∼6% of the GC B cells were CFP+ (Fig. 2A). To visualize CRC–GC B cell interactions, we transferred this mixture of CFP transgenic and WT B cells into Cxcl12-GFP MD4 mice and, 1 d before imaging, injected additional CMTMR-labeled WT B cells to label a portion of the follicular B (FOB) cells and outline the GC (Fig. 2B, Supplemental Video 3). CFP+ GC B cells were observed crawling in and out of the visible DZ CRC networks, sometimes in contact with multiple processes at once, and were seen displacing the fine CRC processes as they moved (Fig. 2C, Supplemental Video 4, Supplemental Video 5). Rotation of the images showing the three-dimensional networks revealed that the GC B cells were often completely surrounded by CRC processes (Supplemental Video 3). GC B cells in CRC networks moved with an average median velocity of 5.44 μm/min ± 0.56 (n = 5) and an average median turning angle of 68.04° ± 2.06 (n = 5). The average median velocity was in the range described for GC B cells in pLN GCs, and the average median turning angle was at the high end of the previously described range (26, 28).
CRCs are phenotypically distinct from FDC and FRC across tissues, but likely lineage related
Because the properties and origin of FDCs in the LZ vary between secondary lymphoid tissues (12, 30, 31), we investigated whether splenic DZ CRCs had a different relationship to FDCs and FRCs than did pLN DZ CRCs. Expanding on the initial characterization of flu-infected mediastinal LN CRCs (15), we stained lymphoid tissues from LCMV-infected and SRBC-immunized Cxcl12-GFP mice for canonical markers of FDCs. In splenic GCs, CRCs expressed low to undetectable levels of CD21/35 and VCAM1 (Fig.1A, 3A) and undetectable levels of FDC-M2, FDC-M1, and CD16/32 (FcγRIII/II) (Fig. 3A). CRCs in pLN, MLN, and PP GCs also expressed low to undetectable levels of CD21/35 and undetectable levels of CD16/32 (Fig. 1B, 1C, Supplemental Fig. 2A), as supported by our previous work (15). PDGFRβ is widely expressed by FRCs, pericytes, and other mesenchymal cells (10, 31), but this marker was undetectable on CRCs in splenic and pLN GCs as well as pLN primary follicles (Fig. 3B). Also unlike FRCs, CRCs in splenic GCs had minimal association with the reticulum and did not stain for anti-laminin (Fig. 3C) and anti–type IV collagen (Supplemental Fig. 2B), except at the T zone proximal edge of the GC where the CRC network meets the FRC network. This observation is consistent with findings in mediastinal LN GCs (15).
DZ CRCs in pLN GCs did express gp38 (podoplanin), a defining marker of FRCs, as did LZ FDCs (Fig. 3D, Supplemental Fig. 2C) (32). Of interest, we also observed gp38+ stromal processes in the pLN GC DZ that were undetectable for Cxcl12-GFP and low to undetectable for CD35 (Fig. 3D, Supplemental Fig. 2C). This finding is consistent with the data in Fig. 1F, Supplemental Fig. 1B, and Supplemental Fig. 1C and further supports the presence of an additional stromal cell type in the DZ. By comparing the areas occupied by gp38+ CD35− Cxcl12-GFP+ CRC processes and gp38+ CD35− Cxcl12-GFP− DZ stromal processes in each of several GC DZs, we found that non-CRC DZ stroma accounted for half of the DZ stromal network (50.06% ± 26.56, n = 7). However, this proportion varied widely between individual GCs.
Although we found CRCs to be phenotypically distinct from FDCs and FRCs, we inquired whether they might still share a precursor with these cells. To test for a relationship with FDCs, CD21-Cre mice (21) were crossed with R26-ZsGreen reporter mice. Analysis of the resulting mice revealed CD21/35lo/− ZsGreen+ CRC-like stromal cells in spleen, MLN, and pLN GC DZs and pLN primary follicles, along with the expected CD21/35hi ZsGreen+ FDCs in these tissues (Fig. 4A). We also observed the recently reported CD21/35− ZsGreen+ versatile stromal cells (VSCs) in the T zone in pLNs (33) and similar CD21/35− ZsGreen+ FRC-like cells, possibly also VSCs, in the GC-proximal T zone in spleen and MLNs (Fig. 4A). To mark Ccl19-expressing FRC and lineage-related cells, we crossed Ccl19-Cre mice (8) to R26-EYFP reporter mice. We found CD21/35lo/− EYFP+ CRC-like stromal cells in spleen, pLN, MLN, and PP GC DZs of Ccl19-Cre R26-EYFP mice, along with the expected EYFP+ T zone FRCs (Fig. 4B) (8). The majority of CD35hi FDCs in splenic, pLN, and MLN LZs were also Ccl19-Cre lineage marked in agreement with previous work (8, 34). However, in PPs the extent of lineage marking of FDCs was variable (Fig. 4B). With both reporters, stroma throughout the GC DZ was lineage marked, providing strong support for the conclusion that DZ CRCs were labeled as well as any non-CRC DZ stroma. In summary, CRCs do not express most of the distinguishing markers of FDCs or FRCs but may share a lineage precursor with both stromal cell types.
CRCs do not require LT or TNF for maintenance of Cxcl12 expression or network morphology
FDCs require LT and TNF signaling for their maintenance (4–6). To study whether CRCs similarly required intact LT or TNF signaling, we immunized Cxcl12-GFP mice with SRBCs and, on day 10, treated them with LTβR-Fc and TNFR-Fc to block signaling from the respective receptors. At 4 d post treatment, we observed CRC networks still visible in, and predominantly confined to, the T zone proximal side of GCs in spleen, MLNs, and PPs (Fig. 5A, 5B). FDCs, however, no longer expressed CD35, indicating the initial decline of this population, as expected with effective blockade (Fig. 5A, 5B) (5). CRC networks retained morphologies similar to those in the saline-treated mice, although some splenic GCs additionally contained GFP+ stromal cells in the T zone distal region of the GC (Fig. 5A), and CRC networks in PPs receded slightly toward the serosal surface as a proportion of the GC diameter (Fig. 5A). FDCs in MLN GCs were less affected at this time point. Therefore, across the tissues, we quantified CRC presence only in GCs with complete loss of CD35 expression on FDCs (Fig. 5B).
To better relate the observed CRC independence from LT and TNF signaling to the well-reported FDC requirement, we attempted to test whether FDCs are lost from the LZ after this short blockade or have just lost expression of essential surface markers. With the same treatment as before, we analyzed the GC stroma in UBI-GFP mice previously reconstituted with WT BM, as FDCs would still be detectably GFP+ in these mice even if they lost CD35 expression. At 4 d post treatment, stromal cells were still present throughout the GC in spleen, MLNs, and PPs (Fig. 5C, Supplemental Fig. 3), suggesting FDCs were not completely lost. However, the T zone distal GC stroma no longer had the dense, mesh structure of LZ FDCs and instead consisted of dispersed processes with a morphology similar to that of CRCs (Fig. 5C, Supplemental Fig. 3). We propose that these dispersed networks are former FDCs that have lost CD35 expression and adopted a CRC-like morphology in the absence of LT and TNF signaling. Despite the morphological similarity, we suggest these cells are not expanded CRCs based on our observation that CRCs in LTβR-Fc– and TNFR-Fc–treated Cxcl12-GFP tissues did not expand into the LZ. Thus, in a timeframe when FDCs rely on LT and TNF signaling for functional and structural maintenance, CRCs do not require LT and TNF signaling to maintain Cxcl12 expression and network morphology.
CXCR4 blockade disrupts CRC distribution in the GC DZ
Because GC B cells must upregulate CXCR4 to travel to the CXCL12-rich DZ (13), we investigated whether CXCR4 signaling plays a role in CRC network maintenance. Blockade of CXCR4 with genetic knockout mice or inhibitors causes GC depolarization marked by appearance of FDCs throughout the GC instead of predominantly in the LZ (13). We hypothesized that the depolarization of FDCs indicated that either CRCs were converting into FDCs or CRCs were being displaced from the DZ in the absence of CXCR4 signaling. To investigate these possibilities, we treated SRBC-immunized Cxcl12-GFP mice with the CXCR4 inhibitor 4F-benzoyl-TE14011 or saline for 12 h via osmotic pumps. Because the half-life of GFP in vivo has been estimated to be ≥26 h (35), this time point allowed us to assess the conversion of CRCs to FDCs even if the CRCs no longer expressed Cxcl12. As expected with an effective CXCR4 blockade, CD35+ FDCs were detectable throughout GCs in spleen and PPs (Fig. 6). However, the dispersed stroma did not include Cxcl12-GFP+ stroma expressing FDC markers, as would have been expected if CRCs were converting to FDCs. Instead, Cxcl12-GFP+ CRC networks were found collapsed against the DZ boundary with the T zone in splenic GCs or against the serosal-proximal lymphatics in PP GCs. Collapse of CRC networks was not observed in the GCs of saline-treated mice (Fig. 6). We suggest that CRCs require CXCR4 signaling, likely in GC B cells, for structural maintenance, but not for sustaining phenotypic distinction from FDCs.
The above findings build from recent work (15) to establish the morphology, distribution, lineage, and maintenance requirements of CRCs in splenic, pLN, MLN, and PP GC DZs. Our observations that DZ CRCs have low network density and variable morphology are in accord with ultrastructural studies of human tonsil GC stroma, which described FDC-like cells in the DZ as half as dense as FDCs in the LZ and lacking in the LZ FDC labyrinth-like structure (16, 17). Despite structural differences between CRCs and FDCs, we show that GC B cells crawl in and around CRC networks with a motion similar to their activity in FDC networks (12, 26). These data suggest CRCs could likewise support GC B cells in the DZ niche through structural maintenance of the compartment and direct cell–cell interactions.
Although previous studies of human tonsil GCs proposed that DZ stromal cells were a less differentiated form of LZ FDCs (16, 17), we find that CRCs and FDCs are already established as distinct populations in both pLNs (15) and splenic primary follicles prior to GC formation. The role of CRCs in primary follicles is not yet clear. Although naive B cells have been shown to respond to CXCL12 for entry into pLNs and entry and exit from PPs (36, 37), further study is needed to characterize the effect of CRC-derived CXCL12 on naive B cell dynamics in the primary follicle and during GC formation.
Our findings in this and previous work (15) that DZ CRCs share almost no canonical surface markers with LZ FDCs or FRCs distinguish CRCs as a distinct cell type across different lymphoid tissues. Lacking these FDC mediators of Ag capture and integrin interaction with GC B cells (38), DZ CRCs likely make distinct contributions to the GC that meet the specialized requirements for the DZ niche (1–3). In human tonsil and LN, DZ stromal cells have similarly been reported to not express FDC markers but have been found to express members of the S100a family of intracellular calcium-binding proteins (39–41). Future studies will be needed to discern whether murine CRCs express S100a family members uniquely among lymphoid stroma and if they contribute to CRC function in the GC.
We did find that DZ CRCs express the FRC-associated surface glycoprotein, gp38. Recently, gp38 has been described as playing a role in regulating the contraction of the LN FRC network and, as a result, the expansion of the LN after immunization (42, 43). Whether gp38 has a similar role in regulating FDC and CRC network expansion during GC formation will require future study. In addition, our combined phenotypic characterization of CRCs as Cxcl12-GFP+ gp38+ CD31− PDGFRβ− from this and previous work (15) suggests a strategy for flow cytometric separation of CRCs from the other known Cxcl12-GFP+ lymphoid stroma. This strategy will likely be useful for investigating CRC-specific expression patterns and functions in the future.
CRCs are likely not the only stromal cells in the GC DZ. We observed that CRCs often only partially occupy GC DZs, that ubiquitous labeling of stroma shows more extensive DZ networks, and that gp38+ CD35− Cxcl12-GFP− reticular networks are visible in pLN DZs. These data support either the presence of an additional stromal cell type or variable Cxcl12 expression by CRCs. GC stromal heterogeneity has precedent in an ultrastructural study of human tonsil that described three morphological types of DZ FDCs and four morphological types of LZ FDCs (16). Future study of the prevalence and properties of Cxcl12-GFP− DZ stroma will be needed to fully understand their relationship to CRCs and their role in the GC response.
Our lineage-tracing experiments conservatively suggest that DZ stromal cells, with morphology, location, and surface marker expression similar to that of CRCs, share a lineage relationship with FDCs, VSCs, and FRCs (33). Recent studies have provided evidence that splenic FDCs develop from perivascular cells (31), whereas LN FDCs develop from marginal reticular cells in the outer region of the follicle (30). These cell types may similarly function as precursors for CRCs. Previous studies have shown that FDCs and FRCs are both marked by the Ccl19-Cre lineage reporter in pLN GCs (8, 34). Our findings suggest that DZ CRC-like stroma share this lineage across tissues. The likely common progenitor for these lymphoid stromal cells is the Ccl19-expressing lymphoid tissue organizer that functions early in LN development (44). However, our finding of heterogeneity in PP FDC labeling may indicate an additional progenitor cell of distinct lineage for this population.
Our finding that DZ CRCs do not require LT and TNF signaling for short-term maintenance of Cxcl12 expression or network morphology further distinguishes them from LZ FDCs that require these factors for their maturation and maintenance (4–6). Previous work supports the independence of Cxcl12-expressing stromal cells from LT and TNF signaling, as Cxcl12 expression was unaffected in spleens of LT- and TNF-deficient mice, whereas Cxcl13 expression was significantly diminished (4). These data do not exclude that DZ CRCs could depend on LT and TNF signaling for other functions.
We demonstrate that DZ CRCs depend on CXCR4, likely in GC B cells, for network distribution. We propose that the collapse of CRC networks after CXCR4 inhibition is due to GC B cells losing attraction to the CXCL12-rich DZ and collecting in the LZ, thus removing structural support from the CRC networks. The substantial hourly exchange of cells between DZ and LZ under normal conditions (45, 46) supports the possibility that the majority of DZ B cells could relocate to the LZ during the 12-h inhibition. Although DZ CRCs do not acquire an FDC phenotype with CXCR4 inhibition, CRC functions may still be impacted. Although it was not possible to investigate the impact of inhibition on non-CRC DZ stroma, the complete occupancy of the GC by FDCs suggests that expansion of non-CRC DZ stroma is unlikely to be the cause of CRC collapse.
Production of CXCL12 is likely one of the essential functions of CRCs in the GC DZ. However, most other secondary lymphoid stromal cells, including FRCs, blood endothelial cells, lymphatic endothelial cells, and red pulp fibroblasts, express Cxcl12 outside the GC (15). Within secondary lymphoid organs, CXCL12 has established roles in promoting B and T cell entry to LNs and PPs (36), in supporting DC entry to spleen (47), in regulating egress from PPs and retention in LNs (37, 48), in aiding vascular development (49), and in guiding plasma cells to the splenic red pulp or LN medulla (50). The requirement for different stromal cell sources of CXCL12 in adjacent niches suggests there is tight control over chemokine protein distribution to create the appropriate gradients. For example, in this and previous work (15), we observed Cxcl12 expression on blood vessels traversing both the LZ and DZ that did not seem to affect GC polarity. This observation is possibly due to vascular endothelial cells expressing CXCR7, a sink receptor for CXCL12 (51). How CRCs maintain the CXCL12 gradient in the DZ when Cxcl12 is being expressed in all bordering niches remains an important question for future investigation.
The authors have no financial conflicts of interest.
We thank Y. Xu and J. An for technical assistance, A. Reboldi and H. Chen for help with TPLSM and confocal microscopy, A. Reboldi for helpful discussion, M. Matloubian for the LCMV-Armstrong, T. Nagasawa for the Cxcl12-GFP mice, C. Lowell for the Ccl19-Cre mice, L. Lanier for the R26-EYFP mice, K. Rajewsky for the CD21-Cre mice, A. Lechner and J. Rock for the PDGFRβ Ab, A. Mujal and A. Gerard for the Alexa 488–conjugated anti-GFP Ab, and J. Browning for LTβR-Fc and TNFR-Fc.
This work and L.B.R. were supported by National Institutes of Health Grant AI45073. O.B. is supported by a Sir Henry Dale Fellowship of the Wellcome Trust/Royal Society. J.G.C. is an investigator of the Howard Hughes Medical Institute.
The online version of this article contains supplemental material.
Abbreviations used in this article:
- bone marrow
- 5-(and -6)-(((4-chloromethyl)benzoyl)amino)tetramethylrhodamine
- Cxcl12-expressing reticular cell
- dark zone
- follicular dendritic cell
- follicular B
- fibroblastic reticular cell
- germinal center
- lymphocytic choriomeningitis virus
- lymph node
- light zone
- mesenteric LN
- peripheral LN
- Peyer patch
- tingible body macrophage
- two-photon laser scanning microscopy
- versatile stromal cell
- Received May 28, 2015.
- Accepted September 9, 2015.
- Copyright © 2015 by The American Association of Immunologists, Inc.