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International Immunology Advance Access originally published online on July 18, 2006
International Immunology 2006 18(9):1375-1384; doi:10.1093/intimm/dxl070
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© The Japanese Society for Immunology. 2006. All rights reserved. For permissions, please e-mail: journals.permissions@oxfordjournals.org

The existence of CD11c+ sentinel and F4/80+ interstitial dendritic cells in dental pulp and their dynamics and functional properties

Jian Zhang1, Nobuyuki Kawashima2,3, Hideaki Suda2,3, Yukiko Nakano4, Yoshiro Takano4 and Miyuki Azuma1

1 Department of Molecular Immunology, Graduate School, Tokyo Medical and Dental University, Tokyo 113-8549, Japan
2 Department of Pulp Biology and Endodontics, Graduate School, Tokyo Medical and Dental University, Tokyo 113-8549, Japan
3 Center of Excellence (COE) program for Research on Molecular Destruction and Reconstruction of Tooth and Bone, Graduate School, Tokyo Medical and Dental University, Tokyo 113-8549, Japan
4 Biostructural Science, Graduate School, Tokyo Medical and Dental University, Tokyo 113-8549, Japan

Correspondence to: M. Azuma; E-mail: miyuki.mim{at}tmd.ac.jp


   Abstract
Top
 Abstract
Introduction
 Methods
 Results
 Discussion
 References
 
Dental caries and pulpitis are the most common bacterial infections in humans. However, the immune responses against bacterial stimulation in dental pulp that is bounded by special hard tissues are poorly understood. We examined the initial immune responses in mouse dental pulp after cusp trimming and acid treatment. Using fluorescence immunohistochemistry, two distinct cell populations were identified in the intact pulp; CD11c+F4/80– and CD11c–F4/80+ cells. CD11c+F4/80– cells were localized in the pulp–dentin (P–D) border of the central pulp beneath the dental fissure, whereas CD11c–F4/80+ cells with dendritic morphology were distributed in the perivascular region of the inner pulp and the sub-odontoblastic layer. CD11c+F4/80– cells, but not CD11c–F4/80+ cells, constitutively expressed toll-like receptors 2 and 4 and CD205, and migrated to the P–D border of the treated side within 2 h after the treatment. In parallel, some of the F4/80+ cells migrated to the inner pulp of the treated side, increased in size and enhanced CD86 expression. At 24 h, the CD86+ cells with high fluorescence intensity had disappeared entirely from the pulp. Concurrently, CD86high cells expressing intermediate levels of CD11c and high levels of MHC class II and F4/80, assessed by using flow cytometry, increased significantly in the regional lymph nodes, suggesting migration of these cells from the dental pulp. Our results are the first to demonstrate the existence of at least two types of dendritic cells (DCs) in dental pulp. The CD11c+ sentinel and F4/80+ interstitial DCs might have distinct territories and unique roles in responding to external stimuli via the dentinal tubules.

Keywords: dendritic cells, dental pulp, infection, innate immunity


   Introduction
Top
 Abstract
 Introduction
Methods
 Results
 Discussion
 References
 
Dendritic cells (DCs) are a heterogeneous group of cells that includes different lineages and states of maturation (1–4). DCs are generated from the bone marrow cells and migrate as immature DCs toward the front line of defense, such as the skin and mucosa. Immature DCs express various pattern recognition receptors, such as toll-like receptors (TLRs), for capturing foreign antigens and have high phagocytotic potential. Activation signaling through TLRs induces a release of various cytokines and mediates innate immune responses by regulating phagocytosis and triggering antimicrobial activity (5, 6). After encountering pathogens and/or factors produced by neighboring tissue cells in response to pathogens, DCs in the skin and mucosa mature partially and migrate from the epithelium to the draining lymph nodes, in which they interact with naive T cells. During migration, DCs alter their functional properties from endocytic to non-endocytic cells and from low- to high-capacity antigen-presenting cells altering the expression of cell surface molecules and by secreting cytokines that trigger the adaptive immune response (1, 6). Thus, DCs bridge the innate and adaptive immune systems and the immune responses between peripheral region and secondary lymphoid tissues.

Dental caries and pulpitis have plagued humans from ancient time and are the most common bacterial infections in humans. Oral bacterial agents induce demineralization of enamel that constitutes an impermeable barrier that protects the underlying dentin and dental pulp in the center of the tooth. Once the enamel barrier is disrupted, oral bacterial products diffuse through the dentinal tubule toward the pulp (7). Odontoblasts (OBs) that produce most of the extracellular matrix components found in dentin constitute a densely packed layer at the dentin–pulp interface like epithelial cells in stratified squamous epithelium and send long cytoplasmic processes into dentinal tubules. Therefore, OBs are the first line of cells encountered by bacteria entering dentin from the oral cavity. Dental pulp is a kind of connective tissue that contains various indigenous and recruited immunocompetent cells along with fibroblast-like cells, nerves and capillaries (8, 9). MHC class II-positive cells with dendritic morphology have been found in the dental pulp in humans (8, 10) and rats (9, 11–13). These cells are heterogeneous in localization and morphology, and they consist of at least two distinct types of cells. One cell type has a highly dendritic appearance and is distributed in the para-odontoblastic region of the outer pulp. Another cell type has more rounded, oval, macrophage-like morphology and is located in the perivascular region of the inner pulp (8, 9, 14). MHC class II+ cells were shown to accumulate along the pulp–dentin (P–D) border after cavity preparation in rats (15) and in the human tooth with caries (16, 17). Therefore, a substantial pulpal immune defense may occur in response to microbial penetration via dentinal tubules.

Despite recent progress in DC biology in humans and mice, an understanding of the exact nature of pulpal DCs remains to be established. This is because the number of antibodies that are applicable to decalcified hard tissues or that are available for rats, which are often used as pulpal experimental models, is very limited. To explore the nature of pulpal DCs, we established a murine model of pulpitis and applied a new technique for the preparation of fresh-frozen sections from hard tissues by using Kawamoto's procedure (18) for immunohistochemistry. In this study, we examined the functional phenotypes and dynamics of DCs in the dental pulp.


   Methods
Top
 Abstract
 Introduction
 Methods
Results
 Discussion
 References
 
Mouse model of pulpitis
Female 3- to 4-week-old BALB/c mice were obtained from the Japan SLC (Shizuoka, Japan). All mice procedures were reviewed and approved by the Animal Care and Use Committee of Tokyo Medical and Dental University. The mice were anesthetized by intra-peritoneal injection of sodium pentobarbital and ketamine. The mesial cusp of the lower first molars was trimmed and smoothed to expose dentin using a dental engine with a tungsten carbide bur (diameter, 0.4 mm) under water-spray cooling. After air-drying, the trimmed surface was etched with 37% phosphoric acid gel for 10 s to keep the dentinal tubules open; the surface was washed and dried.

Decalcified sections
Intact and treated mice were anesthetized by intra-peritoneal injection of chloral hydrate and subjected to vascular perfusion for 20 min with 4% PFA in 0.1 M phosphate buffer (pH 7.3). The lower jaws were removed and immersed for 12 h in the same fixative. The samples were decalcified in neutralized 10% EDTA for 2 weeks at 4°C, dehydrated in ethanol and embedded in paraffin, as described previously (13, 19). Five-micrometer-thick sections were processed for H&E staining.

Fresh-frozen sections
At 2, 24 and 72 h after treatment, the mice were sacrificed, and the lower jaws were dissected. Tissue blocks containing the first molar with alveolar bone were immediately frozen in liquid nitrogen for 5 min. The preparation of fresh-frozen sections followed the technique described by Kawamoto (18). The frozen samples were immersed in a cryodish filled with 5% carboxymethyl cellulose and placed in liquid nitrogen. The frozen block was attached to the sample stage of a cryomicrotome (CM 3050; Leica, Nussloch, Germany) in a cryochamber. An adhesive film was prepared by applying a synthetic adhesive (Cryoglue Type I, Finetec, Tokyo, Japan) to a polyvinylidene chloride film (Finetec). The cutting surface of the sample block was covered with a suitably sized piece of adhesive film, and 5-µm sections were cut. The film sections were fixed on precooled glass slides with double-sided adhesive tape for handling, freeze-dried in a cryochamber at –20°C for 12 h and placed in a sealed box at –80°C until further processing.

Immunohistochemistry
The freeze-dried film sections were processed for enzymatic immunohistochemistry and immunofluorescence staining. Antibodies against the following antigens were used: CD11c (N418, hamster IgG, eBioscience, San Diego, CA, USA), F4/80 (rat IgG2a, American Tissue Culture Collection, Manassas, VA, USA), CD86 (PO3, rat IgG2b) (20), CD205/DEC205 (NLDC-145, Serotec, Oxford, UK), TLR2 (polyclonal rabbit IgG, Imenex, San Diego, CA, USA) and TLR4 (MTS510a, rat IgG2a, Imenex). The film sections were fixed in cold absolute acetone for 2 min and treated with 0.3% H2O2 in PBS. After blocking with normal rabbit serum, the sections were incubated with unlabeled, biotinylated or fluorochrome-conjugated antibodies. For the unlabeled antibodies, biotinylated anti-rat IgG (Vector, Burlingame, CA, USA) or anti-rabbit IgG (SouthernBioTech, Birmingham, AL, USA) was used as the secondary antibody. Negative controls were incubated with isotype-matched control Ig. All the incubation steps were performed in a temperature-controlled microwave processor (MI-77, Azumaya, Tokyo, Japan). To detect TLR2 and TLR4, the primary and secondary antibodies were diluted with an immunoreaction enhancer solution (Can Get Signal; Toyobo, Osaka, Japan). For detection using enzymatic immunochemistry, a Vectastain Elite ABC kit (Vector) was used according to the manufacturer's protocol. The sections were visualized with diaminobenzidine (Merck, Darmstadt, Germany) and were counterstained with hematoxylin. For multicolor immunofluorescence, Alexa 488-conjugated streptavidin (Molecular Probes, Eugene, OR, USA), Alexa 555-conjugated F4/80 or Alexa 647-conjugated anti-CD86 mAb was used. The conjugation of Alexa 555 or Alexa 647 to each mAb was performed using the Alexa Fluor protein labeling kit (Molecular Probes) according to the manufacturer's protocols. If needed, the sections were stained with 4',6-diamidino-2-phenylindole (Cappel, Gaithersburg, MD, USA). Staining profiles were obtained using a fluorescence microscope with a charge-coupled device camera system (IX71, U-LH100HGAPO, Pro600ES-D and CoolSNAP HQ/OL, Olympus, Tokyo) and the images were analyzed using MetaMorph software (Universal Imaging, Downingtown, PA, USA). For quantitative analysis, the mean fluorescence intensity (MFI) of CD11c or F4/80 in three fixed selected fields (250 x 150 µm) of each section was measured using MetaMorph image analysis software (Universal Imaging).

Isolation of lymph node cells and flow cytometry
Lymph node cells (LNCs) were obtained from submandibular lymph nodes (LNs) on the treated side (right, regional) and opposite side (left) from mice in which the right lower first molar had been treated before 24 h and from the sub-mandibular LNs bilaterally from intact mice. The collected LNs were minced and digested with Type I collagenase (400 U ml–1, Sigma, St Louis, MO, USA) at 37°C for 30 min to obtain single-cell suspensions. After counting, the cells were stained with an FITC-conjugated mAb cocktail containing mAbs against CD90.2 (30-H12, BD PharMingen, San Diego, CA, USA), CD19 (1D3, BD PharMingen), CD3 (145-2C11, BD PharMingen) and DX5 (eBioscience), PE-conjugated CD11c or CD86 mAb, and biotinylated-F4/80 mAb, followed by allophycocyanin (APC)–streptavidin, or the appropriate fluorochrome-conjugated control Ig. In addition, some cells were stained with FITC-conjugated anti-CD11c, PE-conjugated anti-CD86 or anti-MHC class II and biotinylated anti-F4/80 or anti-CD86 mAbs, followed by APC–streptavidin, or with the appropriate fluorochrome-conjugated control Ig. Immunofluorescence and flow cytometry were performed using FACSCalibur and CellQuest software (BD Biosciences, San Jose, CA, USA).

Statistics
Statistical analysis was performed using the Mann–Whitney U-test. Values of P < 0.05 or P < 0.01 were considered significant.


   Results
Top
 Abstract
 Introduction
 Methods
 Results
Discussion
 References
 
Establishing a murine pulpitis model
To examine the dynamics of immune cells in the dental pulp after stimulation, we established a murine model of pulpitis. The decalcified paraffin and fresh-frozen sections of the intact and treated molars at 2, 24 and 72 h were stained with H&E (Fig. 1A) and immunohistochemically with anti-F4/80 mAb (Fig. 1B), respectively. OBs were distributed regularly in both the P–D borders beneath the dental fissure and the mesial cusp in the intact pulp. Several cells with elongated profiles were scattered in the border space between the front line of the OBs and the dentinal tubules (a, b, and b'). At 2 h, the layers of OBs at the treated side were disorganized, and some cells had a swollen, translucent cytoplasm (d and d'), suggestive of severe damage to these OBs. Furthermore, some elongated cell profiles were seen to be extending into the dentinal tubules on the treated side (d and d') but not in the central fissure area (c). At 24 h, the OBs showed signs of karyorrhexis (f and f'). At 72 h, reparative dentine formation and reorganization of the OBs had occurred on the treated side (h and h'). No clear differences in histological features were observed in the P–D border under the fissure at any of the time points.


Figure 1
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  		Fig. 1 Cusp trimming and acid etching induce changes of pulpal cells. (A) Decalcified sections from intact molars (a, b) and treated molars at 2 (c, d), 24 (e, f) and 72 (g, h) h after treatment were stained with H&E. Two representative areas, the P–D border under the dental fissure (P–D/f) (a, c, e, g) and the trimmed mesial cusp (P–D/c) (b, d, f, h), are shown. Images are representative of three mice per group. The images b', d', f' and h' show the respective selected regions of b, d, f and h at higher magnification. The longer cells (arrows) are scattered in the P–D border of the intact pulp (a, a', b and b'). Some long, thin cell profiles can be seen in the dental tubules (arrows) underneath the trimmed cusp (d). Bars, 50 µm. (B) Fresh-frozen sections at each time point were stained with anti-F4/80 mAb. Representative images of the mesial pulp on the treated side are shown. Bars, 50 µm.

 
In intact molars, F4/80-positive cells with dendritic profiles were abundant throughout the dental pulp, especially in the perivascular region of the inner pulp and in the OB layer (Fig. 1B). Two hours after treatment, most F4/80+ cells had longitudinal axes directed toward the treated cusp. From 2 to 24 h after treatment, F4/80+ cells became larger and accumulated in the inner mesial pulp. At 72 h, the F4/80+ cells in the mesial pulp had decreased in size and number. Cells stained with anti-CD11c mAb were not detected using enzymatic immunohistochemistry. These results demonstrate that cusp trimming and acid etching induced immunological responses as well as reconstitution of the damaged OBs.

Distinct phenotype of DC-like cells and their dynamics after treatment
To identify the lymphocyte subsets of dental pulp in a more sensitive way, we performed multicolor immunofluorescence staining on fresh-frozen sections. In intact molars, CD11c+ cells were localized in the periphery of dental pulp, especially at the P–D border under the fissure (Fig. 2A, a). A larger population of F4/80+ cells was seen in the central portion of the pulp on the inner side of the CD11c+ cells (Figs 2B, a and 3, a). At 2 h after treatment, CD11c+ cells had quickly migrated to the P–D border, which is closest to the trimmed surface, with increased fluorescence intensity (Figs 2A, b and 3, b). By contrast, the number of CD11c+ cells under the fissure was decreased. Quantitative assessment revealed that the MFI of CD11c in the P–D area of the treated side (P–D/c) was significantly increased at 2 h, whereas that under the fissure area (P–D/f) was apparently decreased (Fig. 2C). The MFI of F4/80 in the inner pulp on the treated side (IP/c) was significantly increased at 2 and 24 h, whereas that under the fissure area was clearly reduced. A few F4/80+ cells had reached the P–D border by 24 h (Figs 2B, c and 3, c). These cells had larger profiles and higher fluorescence intensity. CD11c+ and F4/80+ cells had distinct site- and time-related properties of accumulation and migration. There was no obvious overlap in expression between CD11c and F4/80 anywhere in the intact or treated pulp (Fig. 3). These results demonstrate that two clearly distinct subsets of cells, CD11c+F4/80– and CD11c–F4/80+, exist in the dental pulp and show different dynamics after external stimuli via the dentinal tubules.


Figure 2
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  		Fig. 2 Changes of CD11c+ and F4/80+ cells in the pulp after cusp treatment. Fresh-frozen molar sections at each time point were double stained with mAbs against CD11c (green) and F4/80 (red). Nuclei were stained with 4',6-diamidino-2-phenylindole (DAPI) (blue). Representative images of CD11c (A) and F4/80 (B) with DAPI staining are shown. The selected fields in the light microscopic images are shown in (A). All bars, 100 µm. (C) The left panels show light microscopic images of fresh-frozen sections from intact (upper image) and treated (lower image) molars. The plain line shows the outline of the intact first molar; the dotted line shows the trimmed surface. Bars, 100 µm. The MFI of CD11c and F4/80 staining in the fixed regions, the P–D borders under the fissure (P–D/f), the P–D borders under the mesial cusp (P–D/c) and the inner pulp under the mesial cusp (IP/c) were measured. Values are the mean MFI ± standard deviation from seven to nine mice at each time point. *Statistically different from the intact pulp (P < 0.05).

 

Figure 3
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  		Fig. 3 No overlapped expression between CD11c+ and F4/80+ cells in dental pulp. Fresh-frozen sections from intact and treated molars were stained as described in Fig. 2. Representative merged images of CD11c (green) and F4/80 (red) with 4',6-diamidino-2-phenylindole (blue) staining are shown. The upper panels show light microscopic images. The field of mesial/treated cusp side is selected. Bars, 100 µm.

 
Differential expression of functional receptors between CD11c+ and F4/80+ cells
To further investigate the functional phenotypes of both subsets, we examined their functional molecules for antigen uptake and presentation. The immunostaining for TLR2, TLR4 and CD205 revealed that the localization of positive cells was similar to that of CD11c+ cells in the intact pulp (compare Figs 2A and 4A). After treatment, the changes in the distribution and fluorescence intensity of these immunostained molecules coincided with those of CD11c+ cells. The fluorescence intensities of the three molecules seemed to have increased after 2 h, and the sites of positive signals had moved from the central pulp to the treated side. These results imply that the CD11c+ cells co-expressed TLR2, TLR4 and CD205, whereas F4/80+ cells failed to express these molecules.


Figure 4
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  		Fig. 4 Differential expression of functional receptors between CD11c+ and F4/80+ cells. Fresh-frozen sections at each time point were stained. (A) Antibodies against TLR2, TLR4 and CD205 (all green) were used. Bars, 50 µm. (B) Sections were double stained with anti-F4/80 (red) and anti-CD86 (green) mAbs. Representative images from three mice are shown with 4',6-diamidino-2-phenylindole staining (blue). Bars, 100 µm.

 
Next, we examined the functional molecules required for antigen presentation. Slight expression of CD86 was seen on F4/80+ cells in the center of the intact pulp (Fig. 4B). By contrast, at 2 h after treatment, abundant CD86 expression was observed on F4/80+ cells, especially on the cells that had migrated to the inner pulp of the treated side. The cells that acquired CD86 expression had clearly increased in size and fluorescence intensity. At 24 h, the CD86+ cells with high fluorescence intensity had disappeared entirely from the pulp despite the continued existence of F4/80+ cells. A few MHC class II+ cells were observed in the inner pulp at all time points, and these cells rarely co-expressed F4/80 (data not shown).

Increment of F4/80highCD86highCD11cmed cells in regional LNs
The disappearance of F4/80+CD86+ cells at 24 h implies the migration of CD86+ cells into the regional LNs. To determine whether F4/80+CD86+ cells were increased in the regional LNs, we examined submandibular LNCs from intact and treated mice 24 h after treatment. The total number of LNCs in regional LNs on the treated side was increased significantly compared with the number in LNs on the opposite side and from intact mice (Table 1). The percentage of non-T, non-B and non-NK cells (referred as non-T/B/NK cells), which were identified by negative staining for CD3, CD90, CD19 and DX5 in the regional LNs, was increased significantly compared with LNCs on the opposite side. In all the groups of LNCs, non-T/B/NK cells contained three major populations: CD11c–F4/80+, CD11c+F4/80+ and CD11c–F4/80– cells. A representative regional LN is shown in Fig. 5(A). The percentages of F4/80+, CD11c–F4/80+ and CD11c+F4/80+ within non-T/B/NK cells of the LNCs from the LNs on both the treated and opposite sides were higher than those from intact LNs, although the differences were not significant. The total number of F4/80+ cells in regional LNs was clearly increased compared with the number in both intact and opposite side LNs. Approximately 90% of the F4/80+ cells co-expressed CD11c, and no difference was observed between the three LNC groups (Fig. 5A, b). Consistently, most of F4/80+ cells in the T cell area co-expressed CD11c in the fluorescence histostaining of regional LNs, and the number of F4/80+, CD11c+ or CD86+ cells were clearly increased compared with intact LNs (data not shown). However, in the medullary sinus regions, more abundant F4/80 single-positive cells were observed in both intact and treated LNs. The dominant percentages of F4/80+CD11c+ double-positive cells in flow cytometric analysis may due to the selective loss of F4/80 single-positive cells by our gate setting and the preparation of single-cell suspensions. The FACS profiles with CD11c and CD86 staining (Fig. 5A, c) showed that CD11c-positive cells could be divided into three regions based on the differences in the expression of CD11c and CD86: CD86highCD11cmed (R1), CD86medCD11chigh (R2) and CD86negCD11cmed (R3). All three groups of LNCs contained cells from three regions. The percentage of CD86highCD11cmed (R1) cells in regional LNs increased significantly compared with both that in LNs from intact mice and the opposite side (Table 1). CD86 expression was not detected on CD11c– F4/80+ (R4) cells (data not shown). The FACS profiles with F4/80 and CD86 staining showed that CD86high cells expressed F4/80 (Fig. 5A, d). These results indicate that CD86highCD11cmedF4/80+ cells were increased in the regional LNs 24 h after cusp trimming. To further characterize the CD11cmedCD86high cells, we performed three-color staining with combinations of either CD11c, CD86 and F4/80 or CD11c, MHC class II, and CD86, and then compared the levels of MHC class II and F4/80 within the above three regions. The levels of both F4/80 and MHC class II expression were highest in the R1 cells, followed by R2 cells and then R3 cells (Fig. 5B). Combined, we observed the following four populations of F4/80+ LNCs: (R1) CD86highCD11cmedMHC class IIhighF4/80high cells, (R2) CD86medCD11chighMHC class IImedF4/80high-med cells, (R3) CD86negaCD11cmedMHC class IIlowF4/80med cells and (R4) CD86negaCD11cnegaF4/80med cells. The R1 cells increased significantly in the regional LNs.


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  		Table 1 Comparative analyses of LNCs between intact, regional and opposite side LNs

 

Figure 5
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  		Fig. 5 Profiles of F4/80, CD11c, CD86 and MHC class II expression in LNCs. (A) LNCs were stained with FITC–anti-CD90/CD3/CD19/DX5 mAb cocktail, PE–anti-CD11c or -CD86 mAb and biotinylated anti-F4/80 mAb, followed by APC–streptavidin or with appropriate fluorochrome-conjugated control Ig. Samples were analyzed by flow cytometry. An electronic gate was placed to include lymphocytes and a major population of macrophages with characteristic forward scatter (FSC) and side scatter profiles as shown in (a), and then the logical gate including FITC-negative (non-T/B/NK) cells was set. Note that this gate did not include large macrophages >600 in FSC. The data are displayed as dotted plots (4-decade log scale) with quadrant markers positioned to include >98% of control Ig-stained cells in the lower left. In c, regions R1, R2 and R3 were placed based on the levels of CD86 and CD11c expression. The numbers indicate percentages for each quadrant or region within non-T/B/NK cells. Data are representative LNC samples from six regional LNs. (B) LNCs were stained with FITC–anti-CD11c, PE–anti-CD86 or -MHC class II and biotinylated anti-F4/80 or -CD86 mAbs, followed by APC–streptavidin or with appropriate fluorochrome-conjugated control Ig. Samples were analyzed by flow cytometry. An electronic gate was set on the lymphocyte/macrophage gate, and the expressions of CD86 and CD11c with each staining were displayed. Regions R1, R2 and R3 were placed as described above, and CD86, F4/80 or MHC class II expression was displayed with FSC as dotted plot profiles. The MFI for CD86, F4/80 and MHC class II within each region is shown. Data are representative from six regional LNs.

 

   Discussion
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
References
 
In this study, we established a murine model of pulpitis using cusp trimming and acid etching. Multicolor immunofluorescence staining using undecalcified cryosections of fresh-frozen teeth enabled us to sensitively and accurately examine multiple antigens expressed on the pulpal cells. With this method, we analyzed the initial response of the pulpal immune cells to stimulation via the dentinal tubules. We made several novel findings. (i) Two distinct subsets of pulpal immune cells, CD11c+F4/80– and CD11c–F4/80+ cells, were resident in the intact pulp. (ii) A limited number of CD11c+F4/80– cells were localized in the P–D border of the central pulp beneath the dental fissure, whereas abundant CD11c–F4/80+ cells were distributed in the perivascular region of the inner pulp and the sub-odontoblastic layer. (iii) Most CD11c+F4/80– cells co-expressed TLR2, TLR4 and CD205, and migrated to the P–D border beneath the treated cusp within 2 h. (iv) Some CD11c–F4/80+ cells also migrated to the inner pulp after 2 h, increased in size and showed enhanced CD86 expression. (v) F4/80highCD86highCD11cmedMHC class IIhigh cells assessed by using flow cytometry increased significantly in the regional LNs 24 h after cusp trimming and etching.

In the oral mucosa and skin, two types of DCs have been described: Langerhans cells (LCs) in the epithelial layers and dermal/interstitial DCs in the dermis and lamina propria. LCs act as sentinel cells at the front line of defense in cooperation with epithelial cells (4). Our finding that CD11c+F4/80– cells were located at the P–D border and the OB layer, which is the front line against foreign pathogens, and that these cells migrated rapidly into the potential site of microbial penetration strongly suggests that the CD11c+F4/80– cells are the equivalent of LCs in the dental pulp. Similar to the relationship between the stratified squamous epithelial cells and LCs in the oral mucosa and skin, the OBs and CD11c+F4/80– cells might cooperate in sensing microbial invasion. Note that cusp trimming alone failed to elicit the migration of CD11c+F4/80– cells and that acid etching was required to induce migration (data not shown). It has been accepted that acid etching widens the openings of the dentinal tubules, increases dentin permeability and enhances bacterial penetration of the dentin (21). Durand et al. (22) demonstrated that the stimulation of in vitro differentiated human OBs with lipoteichoic acid, which is a TLR2 ligand, induced the production of the chemokines CCL2 and CXCL10 and enhanced the migration of immature DCs in vitro. It is likely that bacterial invasion induces an initial change in the OBs and that putative cytokines/chemokines secreted from the OBs attract the resident CD11c+F4/80– cells. Gram-positive bacteria, such as streptococci, are frequently identified in dentinal tubules and are associated with the pathogenesis of pulpitis (7). The frequency of Gram-negative bacteria, such as Porphyromonas, is clearly lower, but these bacteria are present in dentinal tubules. TLR2 recognizes peptidoglycan and lipoteichoic acid, which are present in the cell membrane of Gram-positive bacteria, whereas TLR4 recognizes LPS, which constitutes the cell wall of Gram-negative bacteria (5). CD205/DEC205 is a multi-lectin receptor and functions as an endocytic receptor for the uptake of extracellular antigens and their processing (23, 24). The constitutive expression of TLR2, TLR4 and CD205 on CD11c+F4/80– cells suggests that these cells play a crucial role in sensing microbial invasion and in the uptake of pathogens. CD11c+F4/80– cells may function as sentinel DCs in the first line of defense in the pulp, just as LCs do in the skin and oral mucosa. It is reasonable that, in the steady state, the localization of these sentinel DCs is limited to beneath the dental fissure because this site is anatomically and physiologically at high risk of exposure to microbial invasion.

F4/80+ cells are another major subset of immune cells residing in the steady state of the dental pulp. Most F4/80+ cells were located in the interstitial central pulp and within the OB layer under the dental fissure. Unlike CD11c+ DCs, these cells were rare in the front line of the P–D border and did not express TLR2, TLR4 or CD205. The lack of TLRs and CD205 and the location of F4/80+ cells imply that these cells are not primary defense cells that directly react with pathogens, unlike CD11c+ sentinel DCs, but may respond indirectly to cytokines secreted from pathogen-captured sentinel DCs. Most pulpal F4/80+ cells exhibited obvious DC-like morphology, with long processes and spindles. Tsuruga et al. also reported the similar features of F4/80+ cells that were seen in and around the OB layer of mouse molars during postnatal development (25). These F4/80+ cells did not express Mac-1, Mac-2, MOMA-2, CD205, 33D1, MIDC-8 and MHC class II, but co-expressed Fc{gamma}R III/II (CD16/32). Within the F4/80+ cells in the interstitial central pulp, only selected F4/80+ cells accumulated into the treated side within 2 h, increased in size and induced a potent costimulator CD86, suggesting their maturation to antigen-presenting cells. Interestingly, these F4/80+CD86+ double-positive cells observed at 2 h in the inner pulp had disappeared almost entirely by 24 h, and at the same time the cell number of F4/80+ cells and the percentages of F4/80highCD86highCD11cmed cells increased significantly in the regional LNs. The time- and space-dependent distribution of F4/80+CD86+ cells suggests that the increased F4/80+CD86+ cells in the regional LNs include the migrating F4/80+CD86+ cells from the pulp. Although CD11c on the F4/80+CD86+ cells was not detected in the dental pulp, in flow cytometric analysis of LNs, F4/80+CD86+ cells expressed intermediate CD11c. This discrepancy may due to the limited detection sensitivity of immunohistochemistry in addition to the less tissue compatibility of anti-CD11c mAb (N418). Unfortunately, we could not directly examine isolated pulpal cells using flow cytometry because of the limited cell availability and the lack of an established isolation method for pulpal cells. The increased F4/80highCD86highCD11cmed cells in the regional LNs expressed high levels of MHC class II. Although we could not directly assess the antigen-presenting capacity of these cells, they might possess potent antigen-presenting ability because of the high CD86 and MHC class II expression. In our preliminary studies, T cells first appeared in the pulp at 7 days after cusp trimming. It is likely that a sub-population of pulpal F4/80+ cells migrates to LNs and induces T cell priming, and then the induced effector T cells home to the infected pulpal site. Our results suggest that the pulpal F4/80+ CD11c– cells assessed by fluorescence histostaining are interstitial DCs in the pulp that migrate to regional LNs to present antigens.

In general, F4/80 was expressed on tissue macrophages and some interstitial DCs in the peripheral tissues. Both cells are differentiated from monocytes emigrating across the endothelial lining of the bloodstream (26–28). Furthermore, it seems that additional F4/80+ cells that were supplied from the bloodstream accumulated in the IP/c area at 24 h after treatment. Therefore, these F4/80+ cells are heterogeneous cells that include resident macrophage, recruited monocytes and differentiated interstitial DCs. Currently, we cannot discriminate resident F4/80+ cells and migrating F4/80+ cells from interstitial DCs.

It is also possible that F4/80–CD11c+ sentinel DCs in the pulp may migrate to regional LNs, as CD11chigh (R2) cells were also increased in number. In addition to CD11c+ sentinel DCs, the migration of dermal macrophages or inflammatory monocytes to LNs and differentiation into DCs in the LNs have been suggested (26, 29). Therefore, we cannot eliminate the possibility that pulpal F4/80+ cells lacking CD86 also migrate to regional LNs. A trafficking experiment using fluorescence labeling should help to clarify this. Further studies are required to identify the origin and trafficking of pulpal cells.

In conclusion, we demonstrated for the first time that there are at least two distinct subsets of resident DCs in the dental pulp, CD11c+ sentinel and F4/80+ interstitial DCs. The sentinel pulpal DCs might play an initial role in sensing pathogens and inducing TLR-mediated signaling. The interstitial DCs might phagocytose infected OBs and migrate to regional LNs, where they induce T cell priming. Each DC probably plays a unique role in the initial defense against pathogens in the dental pulp. Our findings provide important information for estimating the healing properties of dental pulp and for developing possible treatments of dental restoration.


   Acknowledgements
 
The authors thank T. Okiji (Niigata University, Japan) for helpful discussions. This work was supported by Grants-in-Aid from the ministry of Education, Culture, Sports, Science and Technology of Japan.


   Abbreviations
 
APC, allophycocyanin
DC, dendritic cell
LC, Langerhans cell
LNC, lymph node cell
MFI, mean fluorescence intensity
OB, Odontoblast
P–D, pulp–dentin
TLR, toll-like receptor

   Notes
 
Transmitting editor: K. Okumura

Received 24 March 2006, accepted 22 June 2006.


   References
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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