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Department of Periodontics, School of Dental Medicine, State University of New York, Stony Brook, NY 11794;
Baylor Institute for Immunology Research, Dallas, TX 75204; and Baylor College of Dentistry, Dallas, TX 75246
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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, PGE2) and Th (IL-10, IL-12)
cytokines. Interestingly, the IL-10:IL-12 ratio elicited from P.
gingivalis-pulsed DCs was 3-fold higher than that from
Escherichia coli-pulsed DCs. This may account for the
significantly (p < 0.05) lower proliferation of
autologous CD4+ T cells and reduced release of IFN-
elicited by P. gingivalis-pulsed DCs. Taken
together, these findings suggest a previously unreported mechanism for
the pathophysiology of CP, involving the activation and in situ
maturation of DCs by the oral pathogen P. gingivalis,
leading to release of counterregulatory cytokines and the formation of
T cell-DC foci. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Of particular significance for initiating the immune response within
respiratory (15) and gastrointestinal mucosa
(16, 17, 18, 19, 20, 21) are DCs. In their immature state, DCs are
efficient Ag capture cells, but as they mature, they undergo phenotypic
changes that facilitate their migration toward lymphoid organs and
their unique ability to prime naive T cells (22, 23, 24). The
presence of the proinflammatory cytokines IL-1 and TNF-
promotes
DC migration and maturation (23, 24, 25) and also enhances the
immune response by promoting clonal expansion and differentiation of
Ag-activated CD4+ T cells (26). In
contrast, the presence of IL-10 is associated with reduced DC migration
(27), immunosuppression (28), conversion of
DCs into macrophages (29), and an unfavorable outcome of
diseases, such as leprosy, caused by intracellular pathogens
(30).
Here we show differential localization of DC subpopulations in gingival
mucosa, with immature CD1a+ localized to the
epithelium in health and disease and mature CD83+
DCs restricted to the T cell-rich lamina propria (LP) in CP. The
presence of elevated levels of cytokines IL-1,
PGE2, and IL-10 in the local milieu of active CP
suggests a counterregulatory microenvironment supportive of DC
maturation, but inhibitory of an effective cell-mediated response. In
vitro evidence indicates that P. gingivalis and its LPS
induce DCs to mature but stimulate the release of counterregulatory
cytokines (as in vivo) and promote an inefficient T cell response. We
propose a novel model for the pathophysiology of CP whereby pathogen-
and cytokine-driven DC maturation in situ results in formation of oral
lymphoid foci.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The Institutional Review Board approved all protocols involving human subjects. Informed consent was obtained from all subjects before commencement of the study. All subjects were in good general health and were not on antibiotic, anti-inflammatory, or anticoagulant therapy during the month preceding the baseline exams. Also excluded from the study were subjects with a history of drug allergies, rheumatic fever or other conditions requiring prophylactic antibiotic treatment, gross caries or other dental pathology, fewer than 20 teeth, and pregnant or nursing females.
Collection and handling of clinical specimens, disease categories, clinical protocols
Gingival tissue specimens from 29 adult subjects, including 8
periodontally healthy, 7 with gingivitis, and 14 with CP, were excised
under nerve block anesthesia by sharp dissection. The clinical criteria
for these disease categories have been described previously (5, 31, 32). Gingival tissue was properly oriented in OCT medium by
inserting a tooth landmark (3-mm strip of filter paper) alongside the
tissue specimen, was flash frozen, and then was stored at -80°C.
Cryostat sections measuring 7 µm thick were cut and fixed in cold
acetone for 10 min. Sections were stained with H&E to confirm the
clinical diagnosis by histological means (not shown). The experimental
design for the induction of active (n = 12) or
"quiescent" (n = 20) gingival inflammation (Table I
) in subjects with preexisting mild CP
(minimum of three probing pocket depths of 
5 mm, clinical
attachment levels of 5 mm, bleeding on probing, and horizontal
bone loss at the same sites), as well as the technique for sampling
gingival crevicular fluid (GCF), have been described previously
(32). All four periopaper strips from each tooth were
placed in one glass screw-capped vial and stored at -80°C.
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Sterile saline (400 µl) was added to vials containing strips,
and proteins were eluted by placing the vials on a rotator (Roto-Mix;
Barnstead/Thermolyne, Dubuque, IA) at 250 ± 10 rpm for 1 h
in a 4°C cold room. Initial studies (not shown) established the
dilution of test samples that yielded sufficient volume and sensitivity
for detection of the four cytokines within the linear range of their
respective standard curves (32). Dilutions ranged from
1/2.5 to 1/5 v/v. The diluted samples then were aliquoted into new
vials and frozen at -80°C for subsequent analysis. Commercial ELISA
kits (R&D Systems, Minneapolis, MN.) were used to analyze the levels of
IL-1, IFN-, IL-10, and PGE2 within the GCF
samples. All reagents were brought to room temperature, and assay
diluent (50 µl) was added to each well. The standard or unknown
sample then was added to the wells (200-µl volume), and analysis was
performed as described by manufacturer. Color development and intensity
of the color were measured with an ELISA plate reader (Molecular
Devices, Sunnyvale, CA.). A standard curve was prepared, plotting the
optical density vs the concentration of the cytokine expressed as
pg/30 s.
Single and double immunohistochemistry (enzyme-linked)
Single immunohistochemistry was performed on prefixed frozen
sections by indirect method, as described previously (33).
Sections were stained by the biotin-streptavidin-peroxidase method
(Vectastain ABC Elite kit, Burlingame, CA). The substrate was
3-3'diaminobenzidine tetrahydrochloride (Vector Laboratories,
Burlingame, CA). The primary Abs used are listed (Table II). The specificity was confirmed by
substituting the respective isotype controls for the primary Abs. The
sections were counterstained with hematoxylin. For double staining,
after the first staining, sections were washed and labeled by the
biotin-streptavidin-glucose oxidase method (Vectastain ABC GO
kit). The sections were counterstained with Vector MethylGreen
(Vector Laboratories). The specificity of the secondary Ab was
confirmed by substituting the respective isotype control for the
secondary Abs. A blinded examiner using light microscopy enumerated the
immunoreactive cells. The area of the grid was calculated to obtain the
number of cells per square millimeter. Two different areas were
randomly selected by a blinded examiner in the epithelium and two in
the LP to determine the number of cells per square millimeter. The
number of positively stained cells in the epithelium and LP were also
compared with each other. Statistical analysis is described below.
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Monocyte-derived DCs (MDDCs) were generated as described by
Palucka et al. (34). Briefly, monocytes were isolated from
mononuclear fractions of peripheral blood by negative selection and
seeded in the presence of GM-CSF and IL-4 (1–2 x
105 cells/ml) for 6–8 days, after which flow
cytometry (see Fig. 3) was performed to confirm the immature DC
phenotype
(CD14-CD83-CD1a+).
Cell surface markers of DCs were evaluated by four-color
immunofluorescence staining with the following mAbs: CD1a-FITC
(BioSource International, Camarillo, CA); CD40-PE (Coulter, Seattle, WA
and Coulter-Immunotech, Westbrook, ME); CD80-PE (BD Biosciences,
Mountain View, CA); CD83-PE (Immunotech); CD86-PE (BD PharMingen, San
Diego, CA); HLA-DR-PerCP (BD Biosciences); and CD14 APC (Caltag
Laboratories, Burlingame, CA). After 30 min at 4°C and washing with
staining buffer (PBS, pH 7.2, 2 mM EDTA, and 2% FBS), cells were fixed
in 1% paraformaldehyde. Analysis was performed with FACSCalibur (BD
Biosciences). Marker expression was analyzed as the percentage of
positive cells in the relevant population defined by forward scatter
and side scatter characteristics. Expression levels were evaluated by
assessing mean fluorescence intensity indices calculated by relating
mean fluorescence intensity noted with the relevant mAb to that with
the isotype control mAb for samples labeled in parallel and acquired by
using the same setting.
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CD4+ cells were purified from PBMC of the same donor that were used for DC generation (autologous), as described previously (34). Cells bearing CD4 Ag were isolated from mononuclear fraction through positive selection with anti-CD4 mAb and goat anti-mouse IgG-coated microbeads (Miltenyi Biotec, Gladbach, Germany). Isolation of CD4+ cells was achieved with Minimacs separation columns (Miltenyi Biotec) as described by the manufacturer. In all of the experiments, the isolated cells were 80–90% CD4+ as determined by staining with FITC-conjugated anti- CD4 mAb followed by flow cytometry analysis (not shown).
LPS purification
The methodology for isolation and purification of LPS from P. gingivalis 381 and E. coli American Type Culture Collection (ATCC, Manassas, VA) type strain 25922 was as described previously in our laboratory (35). Briefly, whole cell pellets were subjected to hot-phenol water extraction; the aqueous phase was subjected to extensive dialysis against distilled water, followed by lyophilization and then isopyknic density gradient centrifugation. The LPS-containing fractions were dialyzed extensively against distilled water, lyophilized, and subjected to biochemical analysis for purity (34).
Cytokines from DCs and T cells
To study cytokines produced by MDDCs, culture supernatants were
collected at 24 h from DC pulsed with P. gingivalis
381, E. coli ATCC type strain 25922 (25:1 bacteria-to-DC
ratio), or 100 ng/ml of P. gingivalis or E. coli
LPS. For T cell cytokines (IFN-), culture supernatants were
collected after DCs were cocultured with autologous
CD4+ T cells for 5 days. The production of
cytokines (IL-10, IL-12 p70, IL-1, PGE2, and
IFN-) was determined in triplicate by using commercial ELISA Kits
(R&D Systems) and by following the manufacturer’s instructions. The
values shown in Table III represent the
mean of 10 separate analyses.
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The ability of bacteria-pulsed DCs to stimulate autologous T
cell proliferation was performed as described previously
(36). Day 6 MDDCs were pulsed with either P.
gingivalis (25:1 multiplicity of infection (MOI)); E.
coli (25:1 MOI); 100 ng/ml P. gingivalis LPS; or 100
ng/ml E. coli LPS, for 24 h at 37°C. DCs in complete
RPMI with no bacteria or LPS were used as controls. DCs were washed
extensively and cultured at graded doses (5000 DC/200 µl, 1000
DC/200 µl, and 300 DC/200 µl) in complete RPMI 1640 medium
with 10% human AB serum with autologous CD4+ T
cells (50,000 cells/200 µl). Peak proliferative response was achieved
with 1000 DC, so this number was used for subsequent assays.
Proliferation was determined after 5 days by uptake of tritiated
thymidine (1 µCi/well for the last 16 h). Assay was repeated 10
times on separate days, and the mean results are shown in Table III
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Statistical analyses
Descriptive statistics, means, and SE for the numbers of
immunoreactive cells with each cell surface marker in healthy,
gingivitis, and CP tissues (see Fig. 2) were calculated and analyzed
for statistical significance by Tukey’s multiple comparisons test
(p < 0.05; Minitab, State College, PA).
Differences within epithelium and LP for each individual cell marker
were analyzed by Student’s t test
(p < 0.05). Descriptive statistics, means, and
SE of the clinical indices (not shown) and GCF cytokine levels (Table I) were calculated by using SAS 6.12 (Cary, NC) and Proc Mixed
with experimental group and genotype as the factors and the baseline
levels as the covariate, as described previously (31).
Proc Mixed covariate analysis was used to determine which means were
statistically significant, with the output being least squares means
and least significant difference. Differences in IL-10, IL-1,
PGE2, and IFN- between-group means were
declared only if the p value for the F statistic in the
analysis of covariance was 
0.05, and the least significant difference
also was significant at 0.05. Results of in vitro cytokine levels and
T cell proliferation (Table III) were analyzed by Kruskall-Wallis test
(p < 0.05; Minitab).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We have enumerated immature CD1a+ Langerhans
cells (LCs) and mature CD83+ DCs by using
immunohistochemistry (Fig. 1) within
epithelium and LP of the gingiva in health, gingivitis, and CP (Fig. 2A). Macrophage/myeloid cells
(CD14+) and lymphocyte subsets also were
enumerated within these same tissue compartments for comparison (Fig. 2B).
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In healthy epithelium, lymphocytes also comprise a large number of
cells (Fig. 2B) and, relative to the epithelium, the LP
contains a denser resident population of lymphocytes. Transition from
health to CP was marked by a significant increase
(p < 0.05) in numbers of
CD4+ T cells and CD45RO+ T
cells, both of which were predominantly present in LP, the zone of
mature DC infiltration. CD8+ T cells also
increased in CP, but this increase was only significant
(p < 0.05) in the epithelium, where LCs
reside. B cells were present in large numbers (
150
cells/mm2) at all times and always localized to
the LP, similar to the pattern observed by CD14+
myeloid cells.
Increased levels of DC-mobilizing/maturing cytokines are released in active periodontitis in vivo
We next analyzed the levels of cytokines that might influence DC
redistribution/maturation and T cell activation within this tissue. We
designed a clinical study in human subjects with preexisting mild CP to
determine the level of these cytokines in the GCF in active vs
quiescent disease. The results (Table I) indicate that disease
exacerbation is accompanied by significant changes in the local
cytokine microenvironment, including significant
(p < 0.05) increases in the levels of IL-1,
PGE2, and IL-10. IFN- was present in the GCF
of all CP subjects but did not increase significantly in active
disease.
The oral mucosal pathogen P. gingivalis or its LPS activates DCs to undergo maturation and up-regulate costimulatory molecule expression in vitro
Recent in situ studies of periodontitis in rats (7)
and humans (37) suggest that costimulatory molecule
expression on gingival APCs might play a role in alveolar bone loss and
the local T cell response, respectively. Here, human immature MDDCs
(Fig. 3A) were pulsed with the
oral pathogen P. gingivalis for from 4 to 14 h.
Previous studies from our laboratory (6) as well as
unpublished observations established the optimum MOI (25:1) for
internalization of P. gingivalis by DCs, as confirmed by
trifluorochrome phagocytosis microassay (5) and electron
microscopy (data not shown). As our previous studies indicate that the
LPS of P. gingivalis is an immunodominant Ag capable of
inducing a strong IgG response in human subjects with CP
(31), MDDCs were also pulsed with LPS (100 ng/ml) from
P. gingivalis. The gut commensal E. coli ATCC
type strain 25922 and its purified LPS served as the controls. Our
results show that P. gingivalis (Fig. 3, D and
E), its LPS (Fig. 3F), or LPS of E.
coli (Fig. 3G) induced DCs to undergo maturation in a
time-dependent manner, as evidenced by up-regulation of CD83. Moreover,
expression of costimulatory molecules required for efficient Ag
presentation, including CD80, CD86, CD40, and HLA-Dr also were
up-regulated. The morphologic changes as immature DCs become matured by
LPS also are shown (Fig. 3, B and C).
P. gingivalis-matured DCs release relevant cytokines and stimulate a limited T cell response
Based on in situ, in vivo, and in vitro correlates detailed above,
we had reason to expect that DCs matured by the oral pathogen P.
gingivalis would be potent at stimulating T cell responses, as
determined by IFN- production and proliferation. However, we had
reservations in this regard because of the GCF evidence that IFN-
did not increase significantly in active CP. This was possibly
attributable to the presence of counterregulatory Th2-biasing
cytokines, such as IL-10 (27) and
PGE2 (38). Therefore, we analyzed
the supernatants from the DCs pulsed as above with P. gingivalis,
E. coli and their LPS moieties for proinflammatory cytokines
(IL-1, PGE2), the Th2-biasing cytokine IL-10,
and the Th1-biasing cytokine IL-12. DCs pulsed with either P.
gingivalis or E. coli release IL-1,
PGE2, IL-10, and IL-12, although P.
gingivalis-pulsed DCs released significantly less
(p < 0.05) of all four cytokines (Table III).
The ratio of IL-10:IL-12 elicited by P. gingivalis was
3-fold higher than that by E. coli (7:1 vs 2:1,
respectively). This elevation in IL-10:IL-12 ratio induced by P.
gingivalis may be reflected in the significantly
(p < 0.05) lower IFN- levels from
CD4+ T cells cocultured with DC-P.
gingivalis, relative to DC-E. coli (35 vs 1031 pg/ml,
respectively) and in the limited proliferation of autologous T cells
elicited by P. gingivalis-pulsed DCs as compared with
E. coli-pulsed DCs (Table III).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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164%);
immature DCs in LP (
198%); and CD8+ cells in
epithelium (169%). Although these percentages were not tested
statistically, we have interpreted the increase in mature DCs and
decrease in immature DCs in the same tissues as evidence for local
redistribution of LCs to LP, and subsequent maturation in situ (Fig. 4); however, the actual source of mature
DCs is unknown presently. Our results further show that although
certain cytokines released in this microenvironment (i.e., IL-1 and
PGE2) could promote in situ DCs
activation/maturation and T cell expansion, the counterregulatory
cytokine IL-10 also is released. Finally, the oral pathogen P.
gingivalis is able to induce cultured MDDCs to undergo maturation
and to release relevant proinflammatory (i.e., IL-1 and
PGE2) and Th cytokines (IL-10 and IL-12), but
elicits a limited T cell response compared with E.
coli-pulsed DCs.
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) within the LP
around the dentition. We acknowledge that the term "follicle" might
imply a proliferating B cell lymphoid organ and hence may be
misleading, but nonetheless propose this terminology. We propose that
the development of OLF is initiated in gingivitis by LCs homing to the
gingival epithelium in response to bacteria/Ags in the oral biofilm
(39), as well as a source of the earliest signals for
lymphoid/myeloid trafficking, i.e., epithelium/keratinocytes
(40). That the CD1a+ cells
enumerated in this study were in fact LCs was confirmed by scanning
laser confocal microscopy and double immunofluorescence with CD1a and
the LC-specific marker of Birbeck granules, LAG
(41), as well as single immunoenzyme staining with
langerin (Ref. 43 ; data not shown). Earlier studies have
shown the dynamics of LCs trafficking in the respiratory mucosa in
response to bacterial Ags (15), as well as LCs in oral
mucosa/gingiva in response to plaque/allergen accumulation (10, 39, 42, 44, 45). One study identified increased numbers of LCs
in the gingival epithelium of CP patients and also identified P.
gingivalis (i.e., Bacteroides gingivalis) and
Actinobacillus actinomycetemcomitans within the same tissues
by immunohistochemistry (47). Our previous studies have
shown by scanning laser confocal microscopy that LCs are infected with
P. gingivalis in situ (6). In this same study,
the pathophysiology of chronic gingivitis/mild CP was equated with a
bacterially induced contact hypersensitivity response
(46), wherein Ag (i.e., P.
gingivalis)-sensitized gingival epithelial LCs engage
CD8+ T cells homing to the gingival epithelium
(6). This supposition is consistent with the cellular
infiltrates within the epithelium but is too simplistic to account for
all of our findings, particularly those observed within the LP. The
foci of myeloid and lymphoid cells within the LP of the diseased
periodontium have been characterized by other investigators
(8, 9, 10, 11, 12, 13, 14). Some have interpreted the predominance of
CD4+ activated/memory T cells and B cells within
the LP as evidence that the CP "lesion" is in fact a tertiary
lymphoid organ (48). It is necessary to differentiate,
within OLF, expression of primary immune response markers from those
consistent with recall responses expected within a tertiary lymphoid
organ.
We propose that OLF are not fully developed unless and until sufficient
antigenic challenge is provided by specific pathogens/commensals in the
subgingival flora such as P. gingivalis and
Fusobacterium nucleatum (1, 2, 40). In like
manner, lymphoid follicles in Peyer’s patches do not develop
normally in germ-free animals, but are restored when animals are
monoinfected (49, 50). Both P. gingivalis and
F. nucleatum have been shown to stimulate human gingival
epithelial cells in vitro to produce -defensins, proinflammatory
cytokines, and chemokines in vitro that are required for leukocyte
recruitment (40). We further propose that although T cell
expansion and IFN- production (Table I) occurs in OLF, the
development of a protective cell-mediated response is limited by the
presence of elevated IL-10 and PGE2 levels in
vivo (Table III).
Our in vitro results indicate that DCs matured by P.
gingivalis are able to stimulate a very limited autologous
CD4+T cell response, as compared with DCs matured
by E. coli. Our efforts are now being directed to
understanding the mechanism(s) of this disparity. One possibility being
investigated in the human and murine (51)
systems is that P. gingivalis or its LPS induce a Th2-biased
response in DCs. This is supported by the data in Table III showing an
elevation in IL-10:IL-12 ratio from P. gingivalis-pulsed
DCs, relative to E. coli-pulsed DCs (7:1 vs 2:1), but
confirmation of a Th2-biased response requires more rigorous
investigation.
In conclusion, our results support the role of DCs in the pathophysiology of CP through their activation and in situ maturation in the T cell-rich LP under the influence of oral pathogens and a cytokine milieu that may be counterregulatory to a protective T cell-mediated response.
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Christopher W. Cutler, Department of Periodontics, School of Dental Medicine, State University of New York, Stony Brook, NY 11794-8703. E-mail address: ccutler{at}notes.cc.sunysb.edu
3 Abbreviations used in this paper: CP, chronic periodontitis; DC, dendritic cells; GCF, gingival crevicular fluid; LP, lamina propria; LC, Langerhans cell; MDDC, monocyte-derived DC; MOI, multiplicity of infection; OLF, oral lymphoid follicles.
Received for publication May 29, 2001. Accepted for publication August 7, 2001.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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R. Jotwani and C.W. Cutler Multiple Dendritic Cell (DC) Subpopulations in Human Gingiva and Association of Mature DCs with CD4+ T-cells in situ Journal of Dental Research, September 1, 2003; 82(9): 736 - 741. [Abstract] [Full Text] [PDF]
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 Y.-T. A. Teng THE ROLE OF ACQUIRED IMMUNITY AND PERIODONTAL DISEASE PROGRESSION Critical Reviews in Oral Biology & Medicine, July 1, 2003; 14(4): 237 - 252. [Abstract] [Full Text] [PDF]
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 V. J. Cavanaugh, Y. Deng, M. P. Birkenbach, J. S. Slater, and A. E. Campbell Vigorous Innate and Virus-Specific Cytotoxic T-Lymphocyte Responses to Murine Cytomegalovirus in the Submaxillary Salivary Gland J. Virol., February 1, 2003; 77(3): 1703 - 1717. [Abstract] [Full Text] [PDF]
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