INTRODUCTION

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Adult periodontitis is a chronic destructive disease characterized by the interaction between gram-negative bacteria and the host inflammatory response. A recent study indicated that severe destructive adult periodontitis is a multibacterial infection and that certain combinations of periodontopathogens, namely, Porphyromonas Prevotella, and Fusobacterium spp., seem to be important in the pathogenesis of the disease (38). These bacteria produce an elaborate variety of virulence factors such as proteases, lipopolysaccharides (LPS), and fimbriae (37).

The respective metabolism of these bacteria is also characterized by the production of an identifiable fingerprint of short-chain fatty acids, which are major by-products of anaerobic metabolism that are released into the microenvironment at the infection site (11) and can diffuse across biological membranes (35). Previous studies have demonstrated that these fatty acids inhibit gingival fibroblast proliferation (36), colon cancer cell growth (9), and phagocytosis (3). Moreover, short-chain fatty acids stimulate a gingival inflammatory response via inflammatory cytokine release (30). Our previous study (19) demonstrated that short-chain fatty acids, especially volatile fatty acids present in the culture filtrates of Porphyromonas gingivalis, Prevotella loescheii, and Fusobacterium nucleatum, markedly inhibited murine T- and B-cell proliferation and cytokine production by mitogen-stimulated splenic T cells. Furthermore, we found that a representative volatile fatty acids, butyric acid, induced cytotoxicity and apoptosis in murine thymocytes, splenic T cells, and human Jurkat T cells (20).

Apoptotic cell death generally occurs in a tightly regulated manner, which proceeds mainly through highly conserved genetic mechanisms (41). Several cys-proteases (caspases), especially interleukin-1 (IL-1)-converting enzyme (now called caspase-1) (1), homologous to the product of the Caenorhabditis elegans ced-3 cell death gene, and other IL-1-converting enzyme-like proteases have been implicated in different types of apoptosis (22). Of these caspases, caspase-3 (CPP32) has been well characterized and seems to act in the central pathway of the apoptotic process (22). Since caspase-3 shows the highest homology to the ced-3 product (29) and its tetrapeptide inhibitor (DEVD-CHO) often blocks apoptosis induced by a variety of inducers (2), it is now thought to be the human equivalent of the ced-3 product (29).

LPS, a common component of the cell walls of gram-negative bacteria, exhibits various types of bioactivity and is believed to play important roles in the infectious diseases caused by gram-negative bacteria. LPS is a potent stimulant of the host immune system (26). LPS stimulates the proliferation and differentiation of B cells and initiates the activation of macrophages, resulting in an enhanced immune response. However, little is known about the effects of LPS on T cells. Studying the interaction of representative virulence factors of periodontopathic bacteria, butyric acid, and LPS in the apoptosis-inducing mechanism, we found that LPS potentiates butyric acid-induced peripheral blood mononuclear cell (PBMC) apoptosis. In this study, we found that butyric acid induces apoptosis in human PBMC through a mechanism dependent on caspase-3. Furthermore, we found evidence that LPS, which does not cause PBMC apoptosis itself, potentiates CD3⁺-T-cell apoptosis induced by butyric acid with the increased apoptosis of CD4⁺ and CD8⁺ T lymphocytes.

	MATERIALS AND METHODS

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Reagents. Highly purified butyric acid and LPS from Escherichia coli K-235 were purchased from Sigma Chemical Co. (St. Louis, Mo.). Solutions of butyric acid ranging in concentration from 3.1 to 10 mM were diluted in RPMI 1640 (Gibco Laboratories, Grand Island, N.Y.) medium and adjusted to pH 7.2 with sodium hydroxide.

Isolation of lymphocytes. PBMC were separated from the heparinized venous blood of healthy adults by Ficoll-Hypaque (Pharmacia Biotech, Uppsala, Sweden) gradient centrifugation. After being washed twice in Hanks' balanced salt solution (Gibco), PBMC were cultured at 37°C in a moist atmosphere of 5% CO₂ in a complete medium consisting of RPMI 1640 supplemented with 10% heat-inactivated fetal calf serum, 2 mM L-glutamine, 100 U of penicillin per ml, 100 µg of streptomycin per ml, and 0.05 mM 2-mercaptoethanol. In certain experiments, T cells were separated from PBMC by immunomagnetic cell sorting. PBMC were incubated for 45 min at 4°C with a mixture of monoclonal antibodies (MAb) to CD14, CD16, CD19, and CD56 (Stemcell Technologies, Vancouver, Canada). The cells were then washed, and antibody-loaded cells were depleted by negative magnetic selection with using anti-mouse immunoglobulin G-coated magnetic beads (Dynal, Oslo, Norway). T cells isolated in this fashion were typically >98% CD3⁺ cells as analyzed by flow cytometry (Becton Dickinson, San Jose, Calif.).

Cell proliferation assay. PBMC (4.0 × 10⁵ per well) were stimulated by precoating the wells with 10 µg of anti-CD3 MAb (145-2C11; PharMingen, San Diego, Calif.) per ml or with 5 µg of concanavalin A (ConA) (Sigma) per ml in 0.1 ml of complete medium in flat-bottom 96-well plates. Butyric acid in RPMI 1640 was added to give final concentrations of 0.31 to 10 mM, and each concentration of butyric acid was tested in quadruplicate. After incubation for 42 h, 20 µl of MTT [3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl tetrazolium bromide, 5 mg/ml in phosphate-buffered saline (pH 7.2); Sigma] was added to each well. After 6 h of incubation, the supernatants were decanted and the formazan precipitates were solubilized by the addition of 150 µl of 100% dimethyl sulfoxide (Sigma) and placed on a plate shaker for 10 min. The absorbance at 550 nm was determined on an MT32 spectrophotometric microplate reader (Corona Electric Co., Ibaraki, Japan). The absorbance of the untreated cultures was set at 100%. The mean relative absorbance and the standard error of the mean (SE) were calculated for every concentration of butyric acid tested.

Cell culture for apoptosis. PBMC (4 × 10⁶ per well) were cultured in 1 ml of complete medium in 24-well tissue culture plates (Falcon; Becton Dickinson Labware, Lincoln Park, N.J.) with various concentrations of butyric acid, in the presence or absence of LPS. At the times indicated in the figures, the cells were harvested, centrifuged at 400 × g for 5 min, and washed twice with ice-cold phosphate-buffered saline. The cells were resuspended in 400 µl of hypotonic lysis buffer (0.2% Triton X-100, 10 mM Tris, 1 mM EDTA [pH 8.0]) and centrifuged for 15 min at 13,800 × g (28). Half of the supernatant, containing small DNA fragments, was subjected to gel electrophoresis, while the other half, as well as the pellet containing large pieces of DNA and cell debris, was used for the diphenylamine (DPA) assay (see below).

Gel electrophoresis. One-half of the supernatant was treated with an equal volume of absolute isopropyl alcohol and 0.5 M NaCl to precipitate the DNA and stored at 20°C overnight. After centrifugation at 13,800 × g for 15 min, the pellet was washed with 200 µl of 70% ethanol and allowed to dry at room temperature. The DNA was resuspended in 12 µl of TE solution (10 mM Tris-HCl, 1 mM EDTA [pH 7.4]) plus 3 µl of loading buffer (50% glycerol, 1× Tris-acetate-EDTA, 10% saturated bromophenol blue, 1% xylene cyanol) at 37°C for 20 min and then electrophoresed for 1 h in a 1.7% agarose gel containing 0.71 µg of ethidium bromide per ml. The gels were photographed under UV transillumination.

DNA fragmentation assay. The DPA reaction was performed by the method of Paradones et al. (33). Perchloric acid (0.5 M) was added to the pellets containing uncut DNA (resuspended in 200 µl of hypotonic lysis buffer) and to the other half of the supernatant containing DNA fragments. Then 2 volumes of a solution containing 0.088 M DPA, 98% (vol/vol) glacial acetic acid, 1.5% (vol/vol) sulfuric acid, and 0.5% (vol/vol) of a 1.6% acetaldehyde solution was added. The samples were stored at 4°C for 48 h. The colorimetric reaction was quantified spectrophotometrically at 575 nm with a model UV-160A UV spectrophotometer (Shimazu Co. Ltd., Tokyo, Japan). The percentage of fragmentation was calculated as the ratio of DNA in the supernatant to the total DNA.

Measurement of caspase-1 and caspase-3 protease activity. After incubation of cells (16 × 10⁶) in 24-well tissue culture plates for the indicated times with or without 5 mM butyric acid, all the cells were collected, washed as described above, and suspended in 50 mM Tris-HCl (pH 7.4)-1 mM EDTA-10 mM EGTA. After the addition of 10 mM digitonin, the cells were incubated at 37°C for 10 min. The lysates were clarified by centrifugation at 18,360 × g for 3 min, and cleared lysates containing 50 mg of protein were incubated with 50 µM each Ac-YVAD-MCA (caspase-1 substrate) and Ac-DEVD-MCA (caspase-3 substrate) at 37°C for 1 h. Levels of released 7-amino-4-methylcoumarin were measured with MT32 spectrofluorometer (Corona Electric Co.) with excitation at 380 nm and emission at 460 nm. One unit was defined as the amount of enzyme required to release 0.22 nmol of 7-amino-4-methylcoumarin per min at 37°C.

Determination of hypodiploid DNA. After treatment with the reagents, the cells (10⁶) were washed twice with phosphate-buffered saline and fixed with 80% ethanol in phosphate-buffered saline. After fixation, the medium was removed by centrifugation, 500 µl of phosphate-buffered saline was added to each sample, and the cells were treated with DNase-free RNase A (20 µg per ml) for 30 min at 37°C. Then the supernatants were removed and the cells were stained with 500 µl of propidium iodide (PI; 50 µg per ml) for 15 min. Flow-cytometric analysis was performed with a FACScan flow cytometer (Becton Dickinson) with Cell Fit software. For each sample, 10,000 cells were analyzed and cell debris and clumps were excluded from the analysis by using forward- and side-scatter parameters.

Flow-cytometric analysis. To measurement the annexin-V binding and PI staining of PBMC, cells (10⁶) were harvested and stained with fluorescein isothiocyanate (FITC)-labeled annexin V and propidium iodide (Genzyme, Cambridge, Mass.) as specified by the supplier. Briefly, PBMC (2 × 10⁶) in 1 ml of medium were cultured as indicated for 21 h, washed, and then stained with PI and annexin V-FITC in annexin binding buffer at room temperature for 15 min. Samples were then diluted with the binding buffer and analyzed with Lysis II software by FACScan within 1 h. Annexin-V binding was also performed on CD3⁺, CD4⁺, and CD8⁺ lymphocytes by flow cytometry. Briefly, cells (10⁶) were stained with 4 µl of phycoerythrin (PE)-conjugated anti-CD3, anti-CD4, or anti-CD8 MAb (Becton Dickinson, Mountain View, Calif.) for 30 min at 4°C, washed, and then stained with annexin V-FITC in binding buffer at room temperature for 15 min. The samples were then diluted with the binding buffer and analyzed with a FACScan apparatus within 1 h. Data from 10⁶ cells were analyzed for each sample.

Statistics. Multiple-group comparisons were made by using a one-way analysis of variance followed by post hoc intergroup comparison by the Bonferroni-Dunn test. Where appropriate, Student's t test was used to compare two groups.

	RESULTS

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Butyric acid inhibits proliferative response and induces apoptosis in PBMC. To determine the effect of butyric acid on the proliferative response of PBMC, the cells were stimulated with anti-CD3 MAb or ConA in the presence of different concentrations of butyric acid. As shown in Fig. 1A, butyric acid inhibited both the proliferative responses in a dose-dependent fashion. An inhibitory effect was observed at butyric acid concentrations as low as 1.25 mM with ConA stimulation. With 10 mM butyric acid, the proliferative responses of PBMC were significantly suppressed (by 45.3% with anti-CD3 stimulation and by 40.0% with ConA stimulation). Moreover, the viability of PBMC was more effectively suppressed by butyric acid in the absence than in the presence of the mitogen (data not shown).

View larger version (26K): [in this window] [in a new window]   FIG. 1.   (A) Dose-dependent effects of butyric acids on cell proliferation. PBMC were cultured with the indicated concentration of butyric acid in the presence of solid-phase anti-CD3 MAb () or ConA () for 48 h. Cellular proliferation was determined by an MTT assay and is expressed as the percentage of the absorbance obtained without butyric acid. The results are expressed as the mean ± SE of three different experiments with triplicate cultures. Values significantly different from the corresponding negative controls without butyric acid at P < 0.05 are indicated by asterisks. (B) Agarose gel electrophoresis of DNA extracted from PBMC treated with butyric acid for 16 h. Lanes: M, molecular weight markers (HaeIII-digested X174 DNA); 1, untreated control cells; 2 to 6, cells treated with 5, 2.5, 1.25, 0.62, and 0.31 mM butyric acid, respectively. (C) Activities of caspase-1 and caspase-3 in butyric acid-treated PBMC. PBMC were cultured with or without 5 mM butyric acid for the indicated times. Cell extracts were prepared and caspase activities were measured as described in Materials and Methods. The results are expressed as the mean ± SE of three different experiments with triplicate cultures. Values significantly different from the corresponding negative controls without butyric acid at P < 0.05 are indicated by asterisks.

Activation of caspase-3 in butyric acid-induced apoptosis. The requirement for caspase-1 and caspase-3 in butyric acid-induced apoptosis was determined by their capacity to cleave the caspase-1 substrate Ac-YVAD-MCA and the caspase-3 substrate Ac-DEVD-MCA. Analysis of protease activation during the cell death induced by treatment of PBMC with butyric acid resulted in increased caspase-3 but not caspase-1 protease activity (Fig. 1C). The increase in caspase-3 protease activity began about 8 h after the addition of butyric acid and peaked after 18 to 21 h, reaching levels more than seven times those of control populations. Similar results were obtained with the butyric acid-treated Jurkat cells (data not shown). Enhancement of caspase-3 proteolytic activity induced by treatment of PBMC with butyric acid was inhibited in a dose-dependent manner by treatment with the caspase-3 inhibitor DEVD-CHO, indicating that DEVD-CHO inhibits the activation of caspase-3 protease induced by butyric acid (data not shown).

LPS stimulates butyric acid-induced apoptosis. To examine the effect of LPS on butyric acid-induced apoptosis, PBMC were treated with LPS in the presence of butyric acid. When PBMC were cultured in the presence of 0.625 to 5.0 mM butyric acid for 21 h and quantified by the DNA fragmentation assay, a dose-dependent increase in DNA fragmentation was seen (Fig. 2A). With 5 mM butyric acid, a maximal increase (28.2% ± 2.0%) in DNA fragmentation was induced in PBMC. The addition of 100 µg of LPS per ml potentiated butyric acid-induced DNA fragmentation in PBMC, and increased DNA fragmentation was observed in all the cultures treated with various concentrations of butyric acid (P < 0.05). The addition of LPS alone had no effect on the DNA fragmentation in PBMC. In similar experiments, cells were cultured with 5 mM butyric acid in the presence or absence of different concentrations of LPS (1 to 100 µg/ml) and examined for DNA fragmentation (Fig. 2B). LPS also potentiated the induction of apoptosis by butyric acid in a dose-dependent fashion, and even 1 µg of LPS per ml accelerated butyric acid-induced apoptosis (P < 0.05). On the other hand, PBMC cultures pretreated with anti-CD3 MAb slightly resisted the induction of apoptosis by butyric acid and LPS (data not shown). These apoptosis data, combined with the results of proliferative-response studies, indicate that intact PBMC are sensitive to apoptosis whereas when activated by mitogen, the cells become slightly unsusceptible to butyric acid-induced apoptosis unless they are treated with high concentrations of butyric acid.

View larger version (37K): [in this window] [in a new window]   FIG. 2.   Effect of LPS on butyric acid-induced apoptosis in PBMC. (A) PBMC were treated with the indicated concentration of butyric acid in the presence () or absence () of LPS (100 µg/ml) for 21 h. (B) PBMC were treated with the indicated concentration of LPS (micrograms per milliliter) in the presence of 5 mM butyric acid (BA) for 21 h. Harvested cells were assayed by the DPA assay. The results are expressed as the mean ± SE of three different experiments with triplicate cultures. Values significantly different from corresponding LPS-free butyric acid values at P < 0.05 are indicated by asterisks.

Effect of LPS on flow cytometry analysis. We next examined the effect of LPS on the degree of butyric acid-induced apoptosis by assessing the emergence of hypodiploid DNA (percentage of sub-G₁ cells) in PI-stained samples of 16-h-cultured cells by flow cytometry. As shown in Fig. 3, treatment with 5 mM butyric acid increased the percentage of PBMC with hypodiploid DNA. Furthermore, the addition of LPS stimulated the butyric acid-induced PBMC apoptosis in a dose-dependent manner, as demonstrated by an increase in the percentage of sub-G₁ cells (P < 0.05). A significant increase in effect was noted at 100 µg of LPS per ml. The addition of LPS alone had no effect on the emergence of sub-G₁ cells (data not shown). These results were confirmed by another method of apoptosis detection, annexin V-FITC staining. Figure 4 shows the results of bivariate FITC-annexin V-PI flow cytometry of PBMC after incubation with butyric acid in the presence or absence of LPS (1 to 100 µg/ml) for 21 h. The lower left quadrant of the histogram shows the viable cells, which exclude PI and are negative for FITC-annexin V binding. The lower right quadrant represents the early apoptotic cells, which are PI negative and annexin V positive, indicating the translocation of phosphatidylserine to the external cell surface but the integrity of the cytoplasmic membrane (44). The upper right quadrant represents the nonviable necrotic and late-stage apoptotic cells, which are positive for annexin V binding and PI uptake. After 21 h of incubation in the presence of butyric acid only, there were 7.5% early-stage apoptotic cells and 32.1% late-stage apoptotic and necrotic cells. The addition of LPS increased the number of early- and late-stage apoptotic and necrotic cells induced by butyric acid. The addition of 100 µg of LPS per ml markedly increased the number of late-stage apoptotic and necrotic cells, by up to 46.6%.

View larger version (51K): [in this window] [in a new window]   FIG. 3.   Effect of LPS on sub-G1 accumulation in butyric acid-treated PBMC. PBMC were treated with the indicated concentrations of LPS (micrograms per milliliter) in the presence of 5 mM butyric acid (BA) for 16 h. The DNA content was analyzed by PI staining. The results are expressed as the mean ± SE of three different experiments with triplicate cultures. Values significantly different from corresponding LPS-free butyric acid values at P < 0.05 are indicated by asterisks.

View larger version (47K): [in this window] [in a new window]   FIG. 4.   LPS increases annexin V-FITC staining of apoptotic cells in butyric acid-treated PBMC. PBMC were double-stained with annexin V-FITC and PI after treatment with 5 mM butyric acid (BA) in the presence or absence of LPS (concentrations given in micrograms per milliliter) for 21 h and analyzed by FACScan analysis. Annexin V+, PI cells are the early apoptotic cells, and annexin V+, PI+ cells are the late apoptotic cells. The figure is representative of five experiments with similar results.

LPS increases apoptosis in CD3⁺, CD4⁺, and CD8⁺ T cells. To analyze which populations are associated with apoptosis, cytofluorometric studies with annexin V labeled with FITC were performed on different lymphocyte populations. The expression of annexin V on CD3⁺, CD4⁺, and CD8⁺ T cells was determined by using dual-color flow cytometry. Table 1 shows results obtained with the CD3⁺-T-cell population. After incubation with 5 mM butyric acid, annexin V preferentially bound to CD3⁺ T cells (31.7%) and LPS dose-dependently accelerated CD3⁺-annexin V cell accumulation induced by the butyric acid. The maximum increase (46.6%) in CD3⁺-annexin V cells occurred with the addition of 100 µg of LPS per ml. To further analyze which subpopulations are associated with apoptosis, the expression of annexin V on CD4⁺ and CD8⁺ T cells was also determined. Annexin V preferentially bound to CD4⁺ T cells, and LPS dose-dependently accelerated CD4⁺-annexin V cell accumulation induced by butyric acid (Fig. 5). Increased annexin V binding was not restricted to CD3⁺ and CD4⁺ T cells, since similar increases with butyric acid were seen on CD8⁺ T lymphocytes (Fig. 6). LPS also dose-dependently increased butyric acid-induced CD8⁺-annexin V cell accumulation. The addition of LPS alone had no effect on the increase of annexin V-positive and CD3⁺, CD4⁺, or CD8⁺ cells in PBMC (data not shown).

View larger version (52K): [in this window] [in a new window]   FIG. 5.   Cytofluorimetric analysis of annexin V on CD4+-cell populations. PBMC were double stained with FITC-labeled annexin V and PE-labeled anti-CD4 MAb after treatment with 5 mM butyric acid (BA) in the presence or absence of LPS (concentrations given in micrograms per milliliter) for 21 h and analyzed by FACScan analysis. Data show the expression of CD4 on apoptotic cells. The figure is representative of five experiments with similar results.

View larger version (61K): [in this window] [in a new window]   FIG. 6.   Cytofluorometric analysis of annexin V on CD8+-cell populations. PBMC were double stained with FITC-labeled annexin V and PE-labeled anti-CD8 MAb after treatment with 5 mM butyric acid (BA) in the presence or absence of LPS (concentrations given in micrograms per milliliter) for 21 h and analyzed by FACScan analysis. Data show the expression of CD8 on apoptotic cells. The figure is representative of five experiments with similar results.

	DISCUSSION

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Human PBMC are the first cells to migrate into periodontal tissue and gingival crevices in response to invading pathogens. Therefore, it is interesting to know how PBMC interact with butyric acid, an extracellular metabolite of periodontopathic bacteria, and also with LPS, a common component of the cell walls of gram-negative bacteria. Our previous study (20) demonstrated that butyric acid induced cytotoxicity and apoptosis in murine thymocytes, splenic T cells, and human Jurkat T cells. To support the hypothesis that butyric acid can modulate the immunoregulatory cell population in periodontal tissue by inducing T-cell death through apoptosis, we further examined the capacity of butyric acid to regulate the proliferation and apoptosis of PBMC and the effect of LPS on butyric acid-induced PBMC apoptosis.

In this study, we demonstrated the dose-dependent suppression by butyric acid of the proliferation of mitogen-activated human PBMC (Fig. 1A). This inhibition of PBMC by butyric acid depended in vitro on apoptosis, a specific form of programmed cell death characterized by internucleosomal DNA digestion. This was revealed by gel electrophoresis (Fig. 1B) followed by a colorimetric DNA fragmentation assay (Fig. 2A).

Butyric acid-induced DNA fragmentation of PBMC was accompanied by an increase in caspase-3 activity but not in caspase-1 activity. This suggests that caspase-3 plays a required role in butyric acid-induced PBMC death. Caspase(-like) proteases have been implicated in many forms of apoptosis (18). In particular, Fas-induced apoptosis is inhibited in caspase-1 knockout mice (17) and is also partially inhibited by tetrapeptide inhibitors of caspase-1 (6) and caspase-3 (8). On the other hand, the apoptosis induced by dexamethasone or ionizing radiation is not suppressed in caspase-1 knockout mice (17), although it induces the activation of caspase 3-like enzymes (14, 27). Furthermore, tumor necrosis factor-induced apoptosis in U937 cells is not blocked by a specific peptide inhibitor (DEVD-CHO) of the caspase-3 (8). These findings would suggest that there might be multiple pathways for initiating apoptosis. Therefore, butyric acid-induced PBMC apoptosis may presumably occur in a similar manner to glucocorticoid- and radiation-induced apoptosis. A recent study (24) also showed that butyrate induces a novel pathway of caspase-3 activation and apoptosis in Jurkat cells. Even in our preliminary study, butyric acid-induced apoptosis in Jurkat cells was partially inhibited by a specific peptide inhibitor of caspase-3 (data not shown). Unfortunately, butyric acid-induced apoptosis in PBMC was not suppressed by DEVD-CHO, a specific inhibitor of caspase-3 (data not shown). Although it is not clear why DEVD-CHO does not function in butyric acid-induced PBMC apoptosis in spite of increased caspase-3 activity, it is possible that DEVD-CHO is not readily internalized by intact cells, since it has poor membrane permeability (29). Our results also suggest that normal systemic cells and leukemic cells have different susceptibilities to the inhibitor. In fact, some evidence suggests that phorbol ester potentiates NO-related apoptosis in transformed cell lines but has no effect on normal PBMC (15). This indicates that a more efficient defense mechanism may exist in the differentiated cells than in the undifferentiated or transformed cells.

We have shown that LPS dose-dependently potentiates butyric acid-induced DNA fragmentation in PBMC (Fig. 2B). Flow-cytometric analysis revealed that LPS increased the proportion of sub-G₁ cells (Fig. 3) and the number of late-stage apoptotic cells (Fig. 4) induced by butyric acid. Numerous efforts have been made to define a role for LPS in the regulation of apoptotic events. For example, an inhibitory influence is supported by the observation that LPS prevents spontaneous and TNF--induced apoptosis in human neutrophils (12), spontaneous apoptosis in murine neonatal splenic B cells (45), and Staphylococcus enterotoxin A-induced apoptosis in murine peripheral V3⁺ T cells (43). Conversely, a stimulatory role is suggested by the ability of LPS to induce apoptosis in swine lymphocytes (31), rat pancreatic acinar cells (21), and murine endothelial cells (13). The latter findings appear to be the most consistent with our observations. However, the exact mechanism by which LPS augments butyric acid-induced apoptosis in PBMC remains to be established. In addition, the observation that the addition of LPS alone had no effect on PBMC apoptosis suggests that LPS may indirectly affect PBMC apoptosis via an interaction with butyric acid.

Furthermore, the LPS concentration that is sufficient to accelerate butyric acid-induced apoptosis in PBMC did not affect butyric acid-induced apoptosis in murine splenic T and B cells (data not shown). These results indicate that there may be differences among species in the capability of LPS to potentiate butyric acid-induced apoptosis. LPS can act as a powerful adjuvant for murine T-cell responses (23, 42) and can induce apoptosis in the murine thymus (48). For humans, however, clear information on the capacity of LPS to stimulate T cells or to induce apoptosis has not been reported.

Flow-cytometric analysis revealed that LPS also accelerated butyric acid-induced apoptosis in CD3⁺ T cells. Experiments with fractionated subpopulations of PBMC on flow-cytometric analysis revealed that LPS accelerated butyric acid-induced apoptosis of CD4⁺ and CD8⁺ T cells equally. Although we cannot explain the discrepancy with a previous murine study, which showed that butyric acid-induced murine apoptosis was observed predominantly in CD4⁺ T cells rather than in CD8⁺ T cells, it is possible that butyric acid and LPS modulate apoptosis of CD3⁺ T cells differently in different species. In fact, Estaquier et al. (7) recently showed that CD4⁺ and CD8⁺ cells from HIV-infected patients were equally susceptible to apoptosis induced by exposure to Fas ligand or by Fas triggering antibody. Further studies with a purified subpopulation will be required to determine the major target for the observed effect.

Although the mechanism by which sensitization with butyric acid and LPS primes CD4⁺ and CD8⁺ T cells to increase apoptosis is presently unknown, the effect of monocytes on T-cell apoptosis has become of interest. Monocyte-derived macrophages and phytohemagglutinin-treated monocytes prime peripheral T cells to undergo apoptosis by cell-cell contact (25, 47). Human immunodeficiency virus (HIV) gp120 induces anergy in human peripheral blood lymphocytes by inducing IL-10 production in monocytes (34). Recent reports also indicate that monocytes infected with HIV have enhanced expression of FasL, which can kill uninfected T cells (32), and that monocyte-dependent apoptosis of human T cells requires Fas, CD45, and CD11a/CD18 (46). Therefore, it is possible that cytokine production by monocytes and enhanced expression of FasL on monocytes induced by LPS triggering potentiate the T-cell apoptosis induced by butyric acid. In fact, LPS induces monocyte production of tumor necrosis factor alpha (4, 40), IL-10 (34, 40), and IL-1 (4). Tumor necrosis factor alpha is the crucial mediator of the apoptosis-enhancing effect triggered by LPS (5) and of CD4⁺-T-cell apoptosis in HIV-infected patients (16). IL-10 promotes activation-induced T-cell apoptosis via the Fas pathway (10). Although there is no report that LPS-stimulated monocytes enhance the expression of Fas ligand, the possibility should be considered that human monocytes produce IL-1 after stimulation by LPS and become Fas positive after IL-1 exposure and that normal human pancreatic beta cells that do not constitutively express Fas become strongly Fas positive after IL-1 exposure and are then susceptible to Fas-mediated apoptosis (39). To clarify whether monocytes affect the LPS stimulation mechanism in butyric acid-induced PBMC apoptosis, we performed additional experiments with purified T cells to directly examine the effect of butyric acid and LPS on T cells and demonstrated that LPS directly enhanced the butyric acid-induced apoptosis in purified T cells.

In conclusion, we reported that butyric acid induces apoptosis in human PBMC through a mechanism that is dependent on caspase-3. In addition, exposure of PBMC to LPS rendered them more susceptible to butyric acid-induced DNA damage and apoptosis. Since evidence is provided that LPS does not cause PBMC apoptosis itself, these findings imply that LPS, in combination with butyric acid, potentiates PBMC apoptosis and exerts deleterious effects on the immunoregulatory cell population in periodontal tissue, further supporting the hypothesized role of these bacterial products in the pathogenesis of periodontal disease.