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The Journal of Immunology, 2007, 178, 5659 -5667
Copyright © 2007 by The American Association of Immunologists, Inc.
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Lymphotoxin-Independent Expression of TNF-Related Activation-Induced Cytokine by Stromal Cells in Cryptopatches, Isolated Lymphoid Follicles, and Peyer’s Patches1

Rebekah T. Taylor*, Seema R. Patel*, Eugene Lin*, Betsy R. Butler*, Jason G. Lake*, Rodney D. Newberry{dagger} and Ifor R. Williams2,*

* Department of Pathology and Laboratory Medicine, Emory University School of Medicine, Atlanta, GA 30322; and {dagger} Department of Internal Medicine, Washington University School of Medicine, St. Louis, MO 63110


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Stromal cells play a crucial role in the organogenesis of lymphoid tissues. We previously identified VCAM-1+ stromal cells in cryptopatches (CP) and isolated lymphoid follicles (ILF) in the small intestine of C57BL/6 mice. Nonhemopoietic stromal cell networks in CP and ILF of adult mice also expressed FDC-M1, CD157 (BP-3), and TNF-related activation-induced cytokine (TRANCE). Individual stromal cells were heterogeneous in their expression of these markers, with not all stromal cells expressing the entire set of stromal cell markers. Expression of VCAM-1, FDC-M1, and CD157 on CP stromal cells was absent in alymphoplasia mice deficient in NF-{kappa}B-inducing kinase (NIK) and NIK knockout mice. Administration of lymphotoxin beta receptor (LTbetaR)-Ig to wild-type mice on day 13 resulted in the absence of CP on day 20; delaying administration of LTbetaR-Ig until day 18 resulted in an 80% decrease in the number of CP on day 22 and diminished expression of VCAM-1, FDC-M1, and CD157 on the remaining CP. In sharp contrast, TRANCE expression by stromal cells was completely independent of NIK and LTbetaR. In addition, expression of TRANCE in ILF was concentrated just beneath the follicle-associated epithelium, a pattern of polarization that was also observed in Peyer’s patches. These findings suggest that TRANCE on stromal cells contributes to the differentiation and maintenance of organized lymphoid aggregates in the small intestine.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Stromal cells have long been appreciated for their role as architectural supports for lymphocyte development in primary lymphoid tissues (1, 2). However, recent studies show that stromal cells do not just form an idle matrix in lymphoid tissues, but also perform key functions in the development of adaptive immune responses. For example, networks of follicular dendritic cells (FDC)3 in B cell follicles support germinal center reactions and the attraction, retention, and activation of B cells (3). Fibroblastic reticular cells, while making contact with lymphocytes, produce an elaborate network of fibrous extracellular matrix which serves to support lymphocyte compartmentalization, movement, and the generation of adaptive immune responses (4). Stromal cells are also active players in the initial development of lymphoid tissues. Current models of lymphoid organogenesis hypothesize that lymphoid tissue development is initiated by stromal "organizer" cells interacting tightly with hemopoietic "inducer" cells, leading to the production of chemokines, cytokines, and adhesion molecules that support cell recruitment and organization into a lymphoid tissue (5). The main molecular mechanism driving the formation and permitting the proper functioning of these stromal networks is the cascade of events following engagement of the lymphotoxin (LT) beta receptor (LTbetaR) on stromal cells by LT{alpha}1beta2 (4, 6, 7, 8, 9, 10).

Cryptopatches (CP) and isolated lymphoid follicles (ILF) are microscopic organized lymphoid structures in the lamina propria of the murine small intestine. Although the better known inductive gut-associated lymphoid tissues (mesenteric lymph node and Peyer’s patches (PP)) develop in embryonic life, CP and ILF development does not begin until after birth (11). The main cellular population within CP is a cell type that lacks lymphoid lineage markers but expresses c-kit, Thy-1, and IL-7R{alpha} (12). Some of these cells also express ROR{gamma}t (13), and thus closely resemble fetal lymphoid tissue inducer cells (LTIC) that are integral to lymphoid organogenesis. ILF are larger lymphoid aggregates that contain B cells organized into a follicle that can support a germinal center reaction and the induction of a specific IgA response (14, 15). ILF may be derived from CP as a result of the recruitment of B lymphocytes and their organization into a follicle (16). This hypothesis is supported by the existence of a range of lymphoid structures with features intermediate between those of classical CP and ILF (17) and the influence the commensal enteric flora has on the extent to which ILF are formed (18). Like other lymphoid tissues, the formation of CP and ILF requires LT{alpha}1beta2 signaling through the LTbetaR. Studies using bone marrow (BM) chimera models have shown that the cellular source of LT{alpha}1beta2 used to generate CP and ILF is within the hemopoietic compartment, whereas nonhemopoietic stromal cells provide the LTbetaR (19, 20).

Although the BM-derived cells in CP and ILF have been characterized in detail, very little is known about the stromal cell population(s) residing in these structures. We recently discovered a population of VCAM-1+ stromal cells in both CP and ILF, identifying these cells as candidates for LTbetaR-expressing cells involved in the development and maintenance of these structures (20). Because stromal cell populations in CP and ILF had not been previously described, we have characterized the phenotype and origin of these stromal cell populations in detail, focusing on defining what features of these stromal cell networks were dependent on LT signaling. We found networks of stromal cells in CP and ILF that expressed FDC-M1, CD157 (also known as bone marrow stromal cell Ag-1 and BP-3), and TNF-related activation-induced cytokine (TRANCE) in addition to VCAM-1. TRANCE is a member of the TNF superfamily and its ligand is receptor activator of NF-{kappa}B (RANK) (21). TRANCE expression on stromal cells in CP and ILF exhibited several distinctive features including a lack of dependence on LT signaling and a later onset of expression compared with other stromal cell markers examined. A gradient of TRANCE expression was observed in mature ILF with most of the TRANCE-expressing cells concentrated just beneath the follicle-associated epithelium. The same pattern of subepithelial TRANCE expression was also observed in PP. These observations suggest that TRANCE on stromal cells plays a previously unappreciated role in the differentiation and normal function of CP, ILF, and PP.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Animals

CD132null mice (B6.129S4-IL2rgtm1Wjl/J) and C57BL/6 mice were purchased from The Jackson Laboratory. Breeding colonies of the CD132null and C57BL/6 mice were maintained in a conventional mouse facility at Emory University. Mutant aly/aly mice were obtained from a colony maintained by Dr. K. Newell (Emory University, Atlanta, GA). Gene-targeted NF-{kappa}B-inducing kinase (NIK) knockout mice (22) maintained on a 129/SvEv background were provided by Dr. R. Schreiber (Washington University, St. Louis, MO). TRANCE-deficient mice backcrossed to the C57BL/6 background (23) were provided by Dr. Y. Choi (University of Pennsylvania, Philadelphia, PA). Fischer rats were purchased from Charles River Laboratories. Unless otherwise stated, all animals used in these experiments were at least 6 wk of age. All animal studies were reviewed and approved by the Emory University Institutional Animal Care and Use Committee.

Antibodies

Monoclonal and polyclonal Abs were purchased from BD Pharmingen, unless otherwise stated. The mAbs used to stain cell suspensions for flow cytometry included PE-anti-mouse CD3 (145-2C11), allophycocyanin- anti-mouse B220 (RA3-6B2), and FITC-anti-mouse CD45.2 (104; eBioscience). Rat cells were detected with PE-anti-rat CD3 (G4.18), PE-anti-rat CD45R (HIS24), and biotin-anti-rat CD45 (OX-1). Binding of biotinylated mAb was detected with streptavidin conjugated to allophycocyanin (Caltag Laboratories) or a streptavidin-PE conjugate (Beckman Coulter). The mAbs used for immunofluorescence detection of mouse cells on frozen sections included unconjugated FDC-M1, biotin-FDC-M2 (ImmunoKontact), FITC-anti-E-cadherin (36), PE- and FITC-anti-Thy-1.2 (53-2.1), PE-anti-CD157 (BP-3), FITC- and allophycocyanin-anti-B220 (RA3-6B2), biotin- and unconjugated anti-VCAM-1 (429 MVCAM. A), PE-anti-CD11c (HL3), PE-anti-CD45 (30-F11), PE-anti-CD4 (RM4-5), Alexa Fluor 647-anti-c-kit (2B8; Caltag Laboratories), unconjugated anti-platelet-derived growth factor receptor {alpha} (PDGFR{alpha}; APA5; eBioscience), and unconjugated anti-TRANCE (IK22/5 from eBioscience and 88227 from R&D Systems). Anti-mouse IL-7R{alpha} mAb (clone A7R34) was the gift of Dr. S.-I. Nishikawa (Kyoto University, Kyoto, Japan). Anti-rat mAbs used for immunofluorescence included biotin-anti-CD25 (OX-39), PE-anti-CD45R (HIS24), PE-anti-CD3 (G4.18), PE-anti-VCAM-1 (MR106), and FITC-anti-CD45 (OX-1).

Treatment of mice with LTbetaR-Ig

To generate mice with significantly more small intestinal ILF for histological analysis than normal C57BL/6 mice have, timed pregnant C57BL/6 mice were injected i.v. with purified human LTbetaR-Ig fusion protein as described previously (24). Injections of 100 µg of LTbetaR-Ig were given on embryonic days (E) 14 and 16. Mice were sacrificed at 7–8 wk of age and tissue from the distal third of the small intestine was analyzed. In all of the in utero-treated mice used for histological analysis, the absence of PP was verified by gross examination of the small intestine at the time of tissue collection. To interrupt LTbetaR signaling in neonatal mice, 13- or 18-day-old C57BL/6 mice were treated with 50 µg of LTbetaR-Ig i.p. Quantitation of CP in treated and untreated mice was done by counting lymphoid aggregates on H&E-stained sections and dividing by the total crypt area analyzed using Image J version 1.33u image analysis software (http://rsb.info.nih.gov/ij/). A minimum of 1.5 cm2 of intestinal crypt area was analyzed per mouse.

Preparation of radiation BM chimeras and flow cytometry

BM was eluted from the tibia and femur of 11-wk-old donor Fischer rats. A single-cell suspension was prepared in complete medium and depleted of mature T cells using OX-52 (rat T cell marker) MACS beads and an LD depletion column (Miltenyi Biotec). T cell depletion was verified by flow cytometry. CD132null recipient mice were given 10 Gy of gamma irradiation from a cesium source. Donor BM cells were resuspended in PBS and 2 x 107 cells were injected i.v. into recipient mice. Mice were given drinking water supplemented with neomycin sulfate (2 mg/ml) for 2 wk after transfer. The extent of chimerism was monitored at several time points following BM transfer by flow cytometric analysis of PBMC isolated through centrifugation over Histopaque-1077 (Sigma Diagnostics). To block nonspecific FcR-mediated binding, the isolated PBMC were incubated with supernatant from the 2.4G2 hybridoma line (American Type Culture Collection) for 10 min before addition of mAb. After staining with mAbs, the cells were analyzed on a FACSCalibur cytometer (BD Biosciences) using CellQuest software (version 3.3). By 5 wk after transfer, over 95% of the blood mononuclear cells in each chimera were positive for rat CD45.

Immunofluorescence staining of frozen sections

Small intestines were excised, placed in cold PBS, and opened longitudinally. For horizontal sections, small sheets of tissue (~15 x 20 mm) were stacked (three at a time) and covered with OCT freezing medium (Sakura Finetek). Swiss rolls of intestine were prepared for vertical sections. For E17.5 and day of birth PP, whole intestines were dissected from embryos and covered in OCT. Blocks were quick-frozen in cold 2-methylbutane on dry ice. Frozen sections of 6-µm thickness were cut with a cryostat, air dried for at least 1 h, and fixed for 10 min in acetone at –20°C. Endogenous peroxidase activity was quenched with 0.3% H2O2 in PBS for 30 min at 37°C. All sections were washed in PBS and blocked in TNB buffer (PerkinElmer Life Sciences). Biotinylated primary mAb diluted in TNB buffer were applied for 1 h at room temperature or overnight at 4°C and detected using streptavidin-HRP followed by FITC- or Cy3-tyramide from a tyramide signal amplification (TSA) kit (PerkinElmer Life Sciences). In other cases, purified primary mAbs were detected using a biotinylated polyclonal goat-anti-rat Ig, followed by streptavidin-HRP and FITC- or Cy3-tyramide. Binding of the PE anti-rat VCAM-1 mAb was amplified using a biotinylated polyclonal goat-anti-mouse Ig followed by streptavidin-HRP and Cy3-tyramide. TSA amplification was followed by application of rat serum for blocking and direct detection of surface markers using mAb to Thy-1.2, B220, CD11c, CD45, c-kit, rat CD45R, and/or rat CD3. Although not needed for detection, PE-conjugated anti-CD157 was amplified with biotin anti-PE (PE001; BioLegend) followed by streptavidin-HRP and Cy3-tyramide for consistency in amplification when performing comparisons to parallel sections stained with other markers using TSA amplification. To avoid possible cross-reaction of anti-rat secondary Abs with endogenous rat Ig when staining intestine sections from rat into mouse BM chimeras with the unconjugated rat FDC-M1 mAb, the FDC-M1 mAb was preincubated with biotin-anti-rat IgG2c (A92-1) before application to the section followed by detection of the biotin label with streptavidin-HRP and Cy3-tyramide. For experiments where TSA amplification was not used, sections were stained with directly labeled mAbs or by biotinylated Abs followed by detection with streptavidin-Alexa Fluor 488 or 546 (Molecular Probes). FITC-anti-rat CD45 and FITC-anti-E-cadherin were amplified with Alexa Fluor 488-conjugated goat anti-fluorescein IgG (Molecular Probes). All slides were mounted in ProLong antifade medium (Molecular Probes). Images were acquired using a Zeiss LSM510 confocal microscope or a Nikon Eclipse E400 fluorescent microscope and image editing was done with Photoshop software (Adobe) and/or Spot Advanced software (Diagnostic Instruments). Single-color images of allophycocyanin and Alexa Fluor 647 staining were pseudocolored in red to enhance signal clarity in printed images. All staining patterns described for CP, ILF, and PP were reproducibly observed in sections obtained from two to five mice.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Stromal cell networks in CP and ILF express VCAM-1, FDC-M1, CD157, and TRANCE

We previously identified VCAM-1+ stromal cells in both CP and ILF (20). Given the importance of stromal cells in the organogenesis and architecture of other lymphoid tissues, we sought to further characterize the phenotype of stromal cells in CP and ILF and the dependence of specific stromal cell Ags on LT signaling. We evaluated CP and ILF for the expression of Ags previously localized to stromal cells in other types of lymphoid aggregates. CP were evaluated using horizontal frozen sections of C57BL/6 small intestine. Because ILF are present in small numbers in normal C57BL/6 mice (15), the studies of ILF were done using C57BL/6 mice treated with LTbetaR-Ig in utero on E14 and E16 to generate mice with a much higher frequency of intestinal ILF (19). Fig. 1A demonstrates that a reticular network of stromal cells expressing FDC-M1, CD157, and TRANCE was identified in both CP (identified by Thy-1 staining) and ILF (identified by B220 staining). All of these markers were found on stromal cells distributed throughout CP. In contrast, the distribution of TRANCE staining (detected by the IK22/5 mAb) in ILF was distinct from the other stromal cell markers, with preferential expression of TRANCE on stromal cells located in the subepithelial dome area just beneath the follicle-associated epithelium (FAE).


Figure 1
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  		FIGURE 1. Stromal cell networks expressing FDC-M1, CD157, and TRANCE are present in CP and ILF. A, CP from horizontal sections of small intestine from C57BL/6 mice (n = 7) and ILF from vertical sections of small intestine from C57BL/6 mice treated in utero with LTbetaR-Ig (n = 3) were stained with mAbs against Thy-1.2 or c-kit to identify CP, B220 to identify ILF, and various stromal markers. Note the expression of TRANCE is polarized toward the epithelial side (identified by E-cadherin) of ILF. B, PP from 22-day-old TRANCE-deficient (n = 3) and wild-type littermates (n = 4) were stained with mAbs against TRANCE and CD45 and counterstained with 4',6'-diamidino-2-phenylindole (DAPI). No TRANCE staining is observed in PP from TRANCE-deficient mice and the aggregates of hemopoietic cells in this structure are not normally organized. Scale bar, 50 µm.

 
TRANCE is also expressed in the subepithelial dome area of PP

The concentration of TRANCE-expressing cells immediately beneath the epithelium overlying ILF (Fig. 1A) led us to examine whether TRANCE was also expressed in the corresponding region within PP. TRANCE was found in the subepithelial dome area of PP from C57BL/6 mice in a pattern similar to ILF, and the same distribution of TRANCE staining was also found in colonic and cecal patches (data not shown). The subepithelial staining pattern observed with anti-TRANCE was absent in PP from TRANCE-deficient mice (Fig. 1B). Costaining with CD11c revealed that TRANCE was not expressed on dendritic cells (Fig. 2A), which are known to be a major cell population in the subepithelial dome of PP and ILF. In addition, TRANCE expression did not colocalize with CD4 or CD45 (data not shown). All the staining patterns observed in CP, ILF, and PP with the IK22/5 mAb to mouse TRANCE were reproduced with the 88227 mAb to mouse TRANCE (data not shown).


Figure 2
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  		FIGURE 2. Heterogeneity among stromal cell populations in CP, ILF, and PP is revealed in dual immunofluorescence studies. A, An ILF from a C57BL/6 mouse treated in utero with LTbetaR-Ig was stained with mAbs against TRANCE and CD11c (n = 3). TRANCE is not expressed by the CD11c+ DCs in the subepithelial dome. B, A CP from a C57BL/6 mouse stained with FDC-M1 mAb and anti-VCAM-1 mAb simultaneously (n = 3). Arrowhead points to a cell expressing FDC-M1 and VCAM-1 at equal levels, while the arrow points to a cell expressing only FDC-M1. Nearly all other cells express both markers, but at unequal levels. C, An ILF stained with mAbs against TRANCE and VCAM-1 (n = 3). Although colocalization is observed in a few cells, the stromal cells expressing TRANCE and VCAM-1 are predominantly nonoverlapping populations. D, A PP follicle stained with mAbs against TRANCE and PDGFR{alpha}. Many of the PDGFR{alpha}+ cells located just beneath the basement membrane in the subepithelial dome of the PP also express TRANCE (n = 2). Scale bar, 50 µm.

 
Stromal cells in CP, ILF, and PP are heterogeneous

The discovery of the expression of multiple stromal markers in CP and ILF led us to question whether the stromal cells expressing these Ags were a homogenous population. Double staining of CP and ILF with combinations of two mAbs to stromal cell markers revealed some heterogeneity in the stromal cells. For example, Fig. 2B consists of high magnification images of a CP costained for FDC-M1 and VCAM-1. Some of the stromal cells express equal levels of these markers, resulting in a yellow signal in the merged image, but most of the cells express a higher level of one marker or the other. In addition, a few cells express one marker but appear to be devoid of the other. The same observations were made in costaining experiments for VCAM-1 and CD157 in CP (data not shown). In ILF, the distribution of stromal cells expressing TRANCE was distinct from those expressing other stromal cell markers. Fig. 2C illustrates an ILF costained with TRANCE and VCAM-1. Although a few cells express both markers, the cells expressing TRANCE in the subepithelial dome area are largely separate from those expressing VCAM-1. To further investigate the stromal expression of TRANCE, PP were costained with TRANCE and Abs to PDGFR{alpha}, a receptor associated with cells of mesenchymal origin (9). Although the highest levels of PDGFR{alpha} expression were seen on subepithelial fibroblasts in villi adjacent to the PP, most of the cells expressing PDGFR{alpha} in the subepithelial area of PP also expressed TRANCE (Fig. 2D).

Full expression of TRANCE is delayed in developing PP

TRANCE expression on stromal cells that also express VCAM-1 has been previously reported in the E16.5 peripheral lymph node anlage (25). Given its presence in the developing lymph node, we also investigated TRANCE expression in developing PP. Strong VCAM-1 expression on stromal cells throughout the PP anlage was observed on E17.5 as described previously (26), but TRANCE expression was minimal at this stage in PP development (Fig. 3). The characteristic subepithelial expression pattern of TRANCE was much more evident in PP harvested on the day of birth (Fig. 3), indicating that the onset of TRANCE expression on stromal cells in PP occurs a few days later than expression of VCAM-1, which is first detected on E15.5 (27).


Figure 3
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  		FIGURE 3. Full expression of TRANCE in developing PP is not complete on day E17.5. Sections of PP anlage at day E17.5 (n = 3) and birth (n = 2) were immunostained with Abs to IL-7R{alpha} (to identify LTIC clusters), VCAM-1, and TRANCE. Consecutive sections of the same PP anlage are shown. Note that the expression of TRANCE at E17.5 in this image is less intense and organized than that at time of birth. Other PP anlagen at E17.5 have no visible TRANCE expression. Dashed lines identify the epithelial side of the PP. Scale bar, 50 µm.

 
Clusters of stromal cells are present in the earliest recognizable CP of neonatal mice

CP develop postnatally and can be first appreciated in C57BL/6 small intestine at day 14 by staining for clusters of cells with c-kit and Thy-1 expression (12). Because the earliest steps of lymphoid organogenesis typically require interactions between hemopoietic "inducer" cells and nonhemopoietic "organizer" stromal cells, we asked whether newly formed CP already contained a population of stromal cells in addition to the lymphoid precursor cells. Staining of small intestine sections from 16-day-old C57BL/6 mice identified Thy-1+ CP containing FDC-M1+, VCAM-1+, and CD157+ cells (Fig. 4A). Interestingly, TRANCE was expressed in only a fraction of CP at day 16 (CP in 1 of 3 mice studied), and at variable levels of intensity (Fig. 4B), suggesting that stromal cells in CP may undergo sequential stages of differentiation with onset of TRANCE expression occurring later than the other stromal cell markers we have analyzed.


Figure 4
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  		FIGURE 4. Stromal cell networks are present in the earliest recognizable CP in C57BL/6 small intestine. A, CP from horizontal sections of small intestine from 16-day-old C57BL/6 mice were stained with mAbs against Thy-1.2 in combination with FDC-M1, VCAM-1, or CD157 (n = 3). Consecutive sections of the same lymphoid aggregates stained for different markers are shown. B, Stromal cells in a different CP from a 16-day-old C57BL/6 mouse express TRANCE. Scale bar, 50 µm.

 
Expression of FDC-M1, VCAM-1, and CD157, but not TRANCE, by stromal cells is dependent on the functional presence of NIK

Engagement of the LTbetaR on stromal cells by membrane-bound LT{alpha}1beta2 on hemopoietic cells is required for the organogenesis of most lymphoid tissues, including CP and ILF (7, 19, 20). In aly/aly mice with a mutant form of NF-{kappa}B-inducing kinase (NIK), LTbetaR signaling is impaired and ILF development does not take place (14). However, development of lymphoid aggregates with many features of CP occurs in aly/aly mice despite the defect in LTbetaR signaling (12). We analyzed these CP from aly/aly mice to determine whether stromal cells retained expression of FDC-M1, VCAM-1, CD157, and TRANCE. FDC-M1, VCAM-1, and CD157 expression could not be demonstrated on a stromal cell population in the CP of aly/aly mice, indicating that expression of these markers by stromal cells was dependent on intact LT signaling through the noncanonical NF-{kappa}B activation pathway (Fig. 5). However, stromal cells expressing TRANCE were still present in aly/aly CP. Because it has been suggested that the mutant form of NIK in the aly/aly mutant may retain some kinase activity (28, 29), we also examined CP from gene-targeted NIK-deficient animals. These samples also showed the presence of TRANCE and the absence of FDC-M1, VCAM-1, and CD157 on stromal cells (Fig. 5). The absence of VCAM-1+ and CD157+ cells in CP of aly/aly and NIK-deficient mice was not unexpected because induction of VCAM-1 and expression of CD157 have been shown to depend on LTbetaR signaling (30, 31), but similar data for FDC-M1 expression on stromal cells has not been previously reported. The presence of TRANCE in CP from both aly/aly and NIK–/– mice indicates that the formation of stromal cell networks and some degree of intestinal lymphoid organogenesis can still proceed in the absence of an intact noncanonical NF-{kappa}B signaling pathway downstream of the LTbetaR. Recent studies of mice deficient in the NF-{kappa}B family members p50 and p52 demonstrate the existence of cross-talk between the canonical and noncanonical NF-{kappa}B signaling pathways, thereby suggesting a pathway by which a limited degree of lymphoid organogenesis might be preserved in the absence of NIK (32).


Figure 5
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  		FIGURE 5. Expression of FDC-M1, VCAM-1, and CD157, but not TRANCE, in CP is dependent on a functional NIK. CP from horizontal sections of small intestine from aly/aly (n = 4) and NIK–/– (n = 2) mice were stained with mAbs against Thy-1.2 or c-kit and FDC-M1, VCAM-1, CD157, or TRANCE. The images from the NIK–/– mice represent consecutive sections of the same lymphoid aggregate stained for different stromal cell markers. Scale bar, 50 µm.

 
CP development and stromal cell differentiation is disrupted by treatment of 18-day-old mice with LTbetaR-Ig

The previous experiments indicated that expression of FDC-M1, VCAM-1, and CD157 on stromal cell networks in CP was dependent on NIK, which is a kinase downstream of the LTbetaR in the noncanonical NF-{kappa}B-signaling pathway. To further evaluate the importance of LTbetaR signaling in the maintenance and development of stromal cell networks in CP, we injected 13-day-old C57BL/6 mice with 50 µg of LTbetaR-Ig or 50 µl of PBS i.p. and analyzed CP development in small intestinal tissue 7 days later. A loss of FDC-M1+ cells in germinal centers and a disruption of T/B cell compartmentalization were observed in the spleens of treated mice (data not shown), confirming the efficacy of the LTbetaR-Ig treatment (6, 33). Examination of the small intestine of these animals revealed a complete block in CP development in LTbetaR-Ig-treated animals compared with those that received PBS (Fig. 6B). This experiment confirmed that signaling through the LTbetaR is required for CP development, because LTbetaR-Ig was given at a time point before recognizable CP are present in the intestine. When the same dose of 50 µg of LTbetaR-Ig was given to 18-day-old mice sacrificed 4 days later, histologic analysis revealed a dramatic reduction in the number of CP in LTbetaR-Ig-treated animals (0.84/cm2 of intestinal crypt area) compared with those that received PBS (4.38/cm2) (Fig. 6B). Stromal cells expressing VCAM-1 and CD157 were absent from all CP retained in the LTbetaR-Ig-treated animals, while trace FDC-M1 expression was found on a few of the remaining CP (Fig. 6C). In contrast, TRANCE expression in the remaining CP from these LTbetaR-Ig-treated mice was unaffected. TRANCE expression in PP was also unaffected (data not shown). We propose that a single LTbetaR-Ig treatment in mice at day 18 can block the formation of additional CP and can concurrently disrupt the expression of LTbetaR-dependent stromal cell markers on the CP that remain. Furthermore, the expression of TRANCE on the PP and remaining CP indicates that stromal cell networks in CP can remain intact after inhibition of LT signaling at the level of the LTbetaR.


Figure 6
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  		FIGURE 6. CP are reduced in number and have altered stromal cell networks after treatment with LTbetaR-Ig. A, Two litters of 18-day-old C57BL/6 mice were treated with PBS (n = 5) or 50 µg of LTbetaR-Ig (n = 5) and sacrificed 4 days later. In a separate experiment, 13-day-old mice were injected with PBS (n = 2) or 50 µg of LTbetaR-Ig (n = 2) and sacrificed 7 days later. B, Frequency of CP in untreated and treated mice. A minimum of 1.5 cm2 of intestinal crypt area was analyzed per mouse. C, CP from 22-day-old mice treated with PBS or 50 µg of LTbetaR-Ig on day 18 were stained with mAbs against Thy-1.2 and either FDC-M1, VCAM-1, CD157, or TRANCE. Consecutive sections of the same lymphoid aggregate stained for different markers are shown. Scale bar, 50 µm.

 
Host-derived stromal cells cluster at the site of intestinal lymphoid aggregates developing in CD132null mice 2 wk after reconstitution with rat BM

Stromal organizer cells participating in the development of lymphoid structures including PP and lymph nodes are generally considered to be cells of nonhemopoietic origin. To confirm that the stromal cells involved in the formation of CP and ILF in mouse intestine were also of nonhemopoietic origin, we used a previously established BM reconstitution model (20, 34) in which the recipient mice were CD132-deficient and consequently lacked any of their own CP and ILF. Because stromal cell markers with Ab-defined allelic variants expressed by congenic mouse strains do not exist, we instead used an established method based on transfer of rat BM into mice to generate stable, fully xenogeneic BM chimeras (35). These BM chimeras were made by injecting donor BM cells from Fischer rats (T cell-depleted to prevent the possibility of graft vs host disease) into lethally irradiated CD132-deficient recipient mice. The mAb we use to identify mouse stromal cells in CP are rat anti-mouse mAb that do not react with the corresponding Ags in rat tissue (data not shown) and therefore only recognize host-derived stromal cells. Before preparing the rat-mouse BM chimeras, we examined frozen sections of small intestine from 10-wk-old Fischer rats to verify that CP-like structures were present in rats. In addition to the previously described lymphocyte-filled villus (36, 37), we found structures morphologically identical with mouse CP that contained CD3–, CD45R–, CD25+, CD45+ cells (data not shown). Stromal cells expressing rat VCAM-1 were also present in these aggregates (data not shown). From these studies, we anticipated that rat BM would harbor cells that could induce lymphoid organogenesis in the small intestine of irradiated CD132null mice. Immunostaining of horizontal frozen sections from the chimeric mice revealed the presence of clusters of stromal cells expressing mouse VCAM-1, CD157, and FDC-M1 beginning 2 wk after transfer (data not shown). The frequency, size, and cellularity of these CP-like aggregates increased at later time points after transfer (Fig. 7), with formation of structures with a diameter of up to 200 µm. CP with stromal expression of mouse TRANCE were present in the chimeric animals, but were first detected at 5 wk (in 5 of 12 aggregates) and were still present on a minority of CP (3 of 11) at 11 wk posttransfer (Fig. 7). These CP also contained clusters of rat CD45– cells. Importantly, these CP in the small intestine of chimeras did not show any positive signal when stained with anti-mouse CD45 or anti-rat VCAM-1 mAbs (data not shown), indicating that residual recipient mouse hemopoietic cells and donor rat stromal cells were not present. These results indicate that host-derived mouse stromal cells are capable of rapidly organizing into clusters and nucleating the development of CP-containing rat lymphocytes.


Figure 7
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  		FIGURE 7. Nonhemopoietic stromal networks are present in lymphoid aggregates formed in the small intestine after transfer of rat BM into irradiated CD132null recipient mice. Lymphoid aggregates from horizontal frozen sections of small intestine from rat-mouse BM chimeras (11 wk posttransfer, n = 2) were stained with mAbs against rat CD45 and mouse FDC-M1, VCAM-1, CD157, and TRANCE. Consecutive sections of the same lymphoid aggregates stained for different markers are shown. Scale bar, 50 µm.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Stromal cells of nonhemopoietic origin are known to be a critical component of many types of primary and secondary lymphoid tissues, but comparatively little is known about the stromal cells that populate intestinal CP and ILF. Building on our previous identification of VCAM-1+ cells in CP and ILF (20), we sought to determine when these stromal cells first associated with CP and ILF and determine to what extent the presence of these stromal cells depended on signaling through the LTbetaR. We showed that stromal cells are already present at the earliest stages of CP development and that the stromal cell markers VCAM-1, FDC-M1, and CD157 found on these cells are dependent on signaling through the LTbetaR for their expression. The most surprising findings emerging from this study concerned stromal cell expression of TRANCE in the CP, ILF, and PP. TRANCE differed from the other stromal cell markers examined in the time course of its induction in CP, its pattern of expression in ILF and PP, and its lack of dependence on LT. These findings in combination with the previously demonstrated requirement for TRANCE in the initial stages of lymph node development (38) suggest that TRANCE-RANK signaling plays a greater role than previously appreciated in the development and normal functioning of these intestinal lymphoid aggregates.

Reticular networks of stromal cells expressing VCAM-1, FDC-M1, and CD157 were found in both CP and ILF. The pattern of distribution of these markers in CP, ILF, and PP is summarized in Table I. The absence of each of these markers on stromal cells in CP from mice with a mutant NIK gene (both aly/aly mice and gene-targeted NIK-deficient mice) demonstrated that induction of these markers is dependent on the noncanonical pathway of NF-{kappa}B signaling. This noncanonical NF-{kappa}B pathway is the primary mechanism by which activation of the LTbetaR initiates lymphoid organogenesis (39). The ability of a single injection of LTbetaR-Ig on day 18 to cause loss of VCAM-1, FDC-M1, and CD157 from the stromal cells in the CP remaining 4 days later demonstrated that signals specifically initiated through the LTbetaR are also responsible for maintenance of these stromal cell markers.


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  		Table I. Distribution patterns of cells expressing specific stromal cell markers in CP, ILF, and PP of adult mice

 
Immunostaining of stromal cells from CP and ILF with sets of two mAb recognizing individual stromal cell markers revealed heterogeneity in the expression of the stromal cell markers analyzed in this study. Although most stromal cells expressed several of the stromal cell markers, the relative levels of expression of these markers varied. Examples of stromal cells expressing one stromal cell marker and lacking a second stromal marker were found. TRANCE-expressing stromal cells in ILF and PP were those most likely to not overlap with the distribution of stromal cells expressing the other stromal cell markers. In aggregate, these findings indicate that the there may be distinct stages of differentiation in stromal cells in CP and ILF and possibly more than one type of stromal cell.

The expression of FDC-M1 on stromal cells located throughout CP and ILF demonstrates that FDC-M1 expression is not restricted to classical FDC, which are typically located in germinal centers. The FDC-M1 mAb was originally selected on the basis of its binding to FDC in the germinal centers of the mouse spleen (40), and the specific Ag recognized by the Ab has not yet been identified. In contrast, the FDC-M2 mAb that detects a form of mouse complement component C4 restricted to FDC (41) was only detected on scattered cells located within the germinal centers of larger ILF (data not shown). Consistent with our findings in CP and ILF, extensive networks of FDC-M1+ stromal cells have also been reported in several other types of tertiary lymphoid structures (42, 43). Dependence of FDC-M1 expression on LT has been shown previously in experiments in which LTbetaR-Ig administration to mice resulted in loss of splenic FDC expressing FDC-M1 (6). There is also prior evidence for LT dependence of VCAM-1 and CD157 expression. In vitro experiments have directly demonstrated that VCAM-1 expression on fibroblasts can be induced by LTbetaR stimulation (30). CD157 expression on splenic stromal cells found within splenic lymphoid follicles was inhibited by in vivo LTbetaR-Ig treatment and was nearly absent in LT-{alpha}- and LT-beta-deficient mice (31).

The earliest time point at which CP are recognizable by routine histology and c-kit immunostaining in C57BL/6 mice is 14 days after birth (12). We found that day 16 CP contained stromal cell networks, indicating that stromal cells are active participants in the early events of CP formation. Stromal cell networks were also present in the rare CP detected at day 14 (data not shown), but we did not identify any clusters of stromal cells before day 14. Because the induction of VCAM-1, FDC-M1, and CD157 on stromal cells is dependent on signaling through the LTbetaR, we propose that a critical threshold of LT signaling must take place in the organizing lymphoid aggregates before the stromal cells begin to express these markers. In C57BL/6 mice, this does not occur before 14 days of age.

The stromal cells involved in lymphoid organogenesis are typically considered to be of nonhemopoietic origin. However, FDC-M1+ cells derived from the donor cells in BM chimeras have been described (44). Such donor-derived FDC-M1+ cells might represent the progeny of mesenchymal stem cells included in preparations of BM cells used to generate radiation BM chimeras (45). We used BM chimeras in which donor marrow from rats was transferred to irradiated CD132null recipient mice to determine whether the stromal cells present in developing CP were of host (nonhemopoietic) or donor (hemopoietic) origin. The stromal cells associated with the CP that developed in these chimeric animals by 5 wk after BM transfer were uniformly positive for mouse VCAM-1, FDC-M1, and CD157, but did not include any cells positive for rat VCAM-1. From these observations, we conclude that the stromal cells involved in the development of intestinal CP are exclusively of nonhemopoietic origin.

The absence of all lymph nodes in TRANCE-deficient mice indicates that TRANCE is an important mediator in lymphoid organogenesis (38). Engagement of the TRANCE receptor RANK on LTIC by TRANCE on stromal cells is known to induce LT{alpha}1beta2 expression by the LTIC (46). TRANCE is not required for PP development (38) and the number of TRANCE-expressing cells is much less in the PP anlage than in the peripheral lymph node anlage (25). However, the reduced size of PP in TRANCE-deficient mice (23, 38) indicates that TRANCE signaling does contribute to normal PP development. Similar changes in PP are described in RANK-deficient mice that duplicate almost all the phenotypic features of TRANCE-deficient mice (47). We have identified polarized expression of TRANCE by stromal cells found in the region beneath the FAE of both PP and ILF. In contrast, TRANCE was expressed diffusely by stromal cells in CP. If one fate of CP is subsequent conversion into ILF, polarization of TRANCE expression in the evolving structure may be an important part of the differentiation process associated with the development of a FAE covering an ILF. Determining whether structures with features intermediate between CP and ILF express TRANCE polarized to the subepithelial dome region may be one criterion that could be used to decide whether specific aggregates merit designation as ILF. The polarized pattern of TRANCE expression in PP and ILF beneath the FAE also suggests a possible function for TRANCE in regulating the induction of mucosal immune responses following uptake of particulate luminal Ags through the FAE including bacteria and viruses. In support of this concept, TRANCE-RANK signaling was previously implicated in the stimulation of IL-10 expression by PP-associated DC and the induction of oral tolerance (48). Furthermore, treatment with soluble TRANCE was recently found to dampen proinflammatory cytokine responses to LPS and other bacterial components in both in vitro and in vivo experiments (49). The elucidation of the mechanisms responsible for the specialized pattern of TRANCE expression in PP and ILF and its potential role in tolerance induction at these sites will be an important area for future studies.


   Acknowledgments
 
We thank Dr. Kenneth Newell for supplying us with alymphoplasia mice and for his valuable comments throughout the course of this project. We thank Dr. Robert Schreiber for providing the NIK knockout mice and Dr. Yongwon Choi for the TRANCE knockout mice.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


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

1 This work was supported by grants from the National Institutes of Health (DK64730 (to I.R.W.), DK64798 (to R.D.N.), and DK64399 supporting the Imaging Core Facility of the Emory Digestive Diseases Research Development Center). Back

2 Address correspondence and reprint requests to Dr. Ifor R. Williams, Department of Pathology and Laboratory Medicine, Emory University School of Medicine, Whitehead Building 105D, 615 Michael Street, Atlanta, GA 30322. E-mail address: irwilli{at}emory.edu Back

3 Abbreviations used in this paper: FDC, follicular dendritic cell; LT, lymphotoxin; LTbetaR, LT beta receptor; CP, cryptopatch; ILF, isolated lymphoid follicle; PP, Peyer’s patch; LTIC, lymphoid tissue inducer cell; BM, bone marrow; TRANCE, TNF-related activation-induced cytokine; RANK, receptor activator of NF-{kappa}B; FAE, follicle-associated epithelium; NIK, NF-{kappa}B-inducing kinase; TSA, tyramide signal amplification; PDGFR{alpha}, platelet-derived growth factor receptor {alpha}; E, embryonic day. Back

Received for publication June 7, 2006. Accepted for publication February 5, 2007.


   References
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 Introduction
 Materials and Methods
 Results
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 Disclosures
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