Homocysteine Induces Protein Kinase C Activation and Stimulates c-Fos and Lipoprotein Lipase Expression in Macrophages

FCS was purchased from Hyclone Laboratories (Logan, UT); DL-Hcys, l-cysteine, and heparin were obtained from Sigma (St. Louis, MO). Dulbecco’s modified Eagle’s medium and Trizol reagent were purchased from Gibco BRL (Burlington, Ontario, Canada). Calphostin C was obtained from Calbiochem (La Jolla, CA).

The J774 murine Mo cell line was obtained from the American Type Culture Collection (Rockville, MD). J774 cells were cultured in Dulbecco’s modified Eagle’s medium containing 10% FCS and 100 μg/ml penicillin-streptomycin (Gibco BRL).

Six million J774 cells were plated in plastic Petri dishes (100 × 20 mm; Falcon, Lincoln Park, NJ). After treatment with Hcys, cells were lysed with Trizol reagent. Total RNA was isolated and separated in a 1.2% agarose gel containing 2.2 mol/l formaldehyde, as previously described (33). The blots were prehybridized for 8 h. The mRNA expression was analyzed by hybridization with [³²P]dCTP-labeled LPL, c-fos, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) cDNA probes. Hybridization was detected by autoradiography with Kodak X-Omat-AR films (Rochester, NY). mRNA expression was quantified by high-resolution optical densitometry (Alpha Imager 2000; Packard Instruments, Meriden, CT).

The isolation of the nuclei was performed as follows. Briefly, 5 × 10⁷ J774 cells were collected, washed with cold phosphate-buffered salt solution, and lysed in 1 ml of ice-cold buffer A (15 mmol/l KCL, 2 mmol/l MgCl₂, 10 mmol/l HEPES, 0.1% phenylmethylsulfonyl fluoride, and 0.5% Nonidet-P-40). After a 10-min incubation on ice, lysed cells were centrifuged, and the nuclei were washed with buffer A without Nonidet P-40. The nuclei were then lysed in a buffer containing 2 mol/l KCL, 25 mmol/l HEPES, 0.1 mmol/l EDTA, and 1 mmol/l dithiothreitol (DTT). After a 15-min incubation period, a dialysis buffer (25 mmol/l HEPES, 1 mmol/l DTT, 0.1% phenylmethylsulfonyl fluoride, 2 μg/ml aprotinin, 0.1 mmol/l EDTA, and 11% glycerol) was added to the nuclei preparation. Nuclei were collected by centrifugation for 20 min at 13,000 rpm. Aliquots (50 μl) of the supernatants were frozen at − 70°C, and protein concentration was determined. DNA retardation (mobility shift) electrophoresis assays were performed as previously described by Fried and Crothers (34). Briefly, 5 μg of nuclear extracts was incubated for 15 min in the presence of 5× binding buffer (125 mmol/l HEPES [pH 7.5], 50% glycerol, 250 mmol/l NaCl, 0.25% Nonidet P-40, and 5 mmol/l DTT). End-labeled double-stranded consensus sequences of the LPL promoter AP-1-enhancing element (20,000 cpm per sample) were then added to the samples for 30 min. Samples were analyzed on a 4% nondenaturing polyacrylamide gel containing 0.01% Nonidet P-40. The specificity of the nuclear protein binding was assessed by incubating the nuclear proteins isolated from murine Mo with labeled DNA probe in the presence of a 1,000-fold molar excess of unlabeled DNA probe.

The cDNA probe for detection of murine LPL was prepared by the polymerase chain reaction technique. cDNA was obtained from total RNA using a reverse transcription reaction. Two synthetic primers spanning bases 255–287 and 1,117–1,149 of the LPL cDNA were used to enzymatically amplify a 894-bp region of the LPL probe. The cDNA probes for murine c-fos and GAPDH were purchased from American Type Culture Collection. A 20-mer double-stranded oligonucleotide (5′ -GGGCACCTGACTAAGGCCAG-3′; 5′-TGTGCTGGCCTTAGTCAGGT-3′) containing the consensus sequence for the AP-1-responsive element of the murine LPL gene promoter (35,36) was synthesized with the aid of an automated DNA synthesizer. After annealing, the double-stranded oligonucleotide was labeled with [γ -³²P]ATP by using the Boehringer Mannheim 5′ end-labeling kit (Indianapolis, IN).

One hour before the end of the incubation period, 50 units/ml heparin was added to the medium. The amount of LPL immunoreactive mass was measured by enzyme-linked immunosorbent assay using the Markit-F LPL kit (Dainippon, Pharmaceutical, Osaka, Japan) (37). Extracellular LPL activity was determined using the Confluolip kit (Progen, Heidelberg, Germany) (38). LPL mass and activity values were normalized to the levels of total cell proteins.

Adherent murine J774 Mo were recovered and homogenized (Dounce: 15 strokes) in 500 μl of ice-cold buffer A (20 mmol/l Tris [pH 7.5], 0.5 mmol/l EDTA, 0.5 mmol/l EGTA, 25 μg/ml aprotinin, and 25 μg/ml leupeptin). The membrane and cytosolic fractions were separated by ultracentrifugation (100,000g for 30 min at 4°C). After recovery of high-speed supernatants containing cytosolic PKC, the corresponding membrane pellets were homogenized in 500 μl of buffer A containing 0.5% Triton X-100. The enzyme from both fractions was partially purified through DE52 chromatography columns. After removal of unbound proteins by washing of the columns with buffer B (20 mmol/l Tris [pH 7.5], 0.5 mmol/l EDTA, 0.5 mmol/l EGTA, and 10 mmol/l β-mercaptoethanol), fractions containing PKC were eluted with buffer C (Tris [pH 7.5], 0.5 mmol/l EDTA, 0.5 mmol/l EGTA, 10 mmol/l β-mercaptoethanol, and 0.2 mol/l NaCl). Eluates were analyzed for PKC activity, following the optimum conditions of the assay, by measuring the incorporation of ³²P into the synthetic peptide Ac-myelin basic protein(4–14). The specificity of the assay was determined by subtracting the radioactivity obtained in the presence of the pseudosubstrate inhibitor PKC(19–36) from total radioactivity. PKC activity was normalized to the levels of total cell proteins. Data were expressed as percentages, considering the control as 100% activity.

Total protein content was measured according to the Bradford method (39) using a colorimetric assay (Bio-Rad, Mississauga, Ontario, Canada).

All values were expressed as the mean ± SE. Data were analyzed by Student’s t test for comparison of two groups or by one-way ANOVA followed by the Student-Newman-Keuls test or the Dunn’s method for multiple comparisons. A value of P < 0.05 was considered significant.

Incubation of J774 cells for 1–24 h with Hcys increased Mo LPL gene expression in a time-dependent manner. Induction of Mo LPL mRNA levels in response to Hcys (1–5 mmol/l) was biphasic, peaking at 1 and 6 h (Fig. 1A). Maximal effect was observed after a 6-h incubation period in the presence of 5 mmol/l Hcys. Under these experimental conditions, no modulation of the mRNA expression of GAPDH, used as internal control, was observed. LPL mRNA levels normalized to the levels of GAPDH mRNA are illustrated in Fig. 1A. No stimulatory effect of cysteine (1 mmol/l) was observed on Mo LPL mRNA levels, thereby indicating a specific effect of Hcys on Mo LPL gene expression (LPL mRNA levels [percent increase of control values]: cysteine 1 mmol/l: 1 h, 94.2 ± 8.2; 6 h, 100.0 ± 9.1).

To characterize further the kinetics of the early induction of LPL gene expression in response to Hcys, J774 cells were incubated in the presence of 1 mmol/l Hcys for 15, 30, 45, and 60 min. As shown in Fig. 1B, a significant increase in Mo LPL mRNA levels was detected after a 30-min incubation period with Hcys. Maximal effect was observed at the 45- and 60-min incubation periods. LPL mRNA levels normalized to the levels of GAPDH are presented in Fig. 1B.

To assess the optimal concentration of Hcys required to stimulate Mo LPL gene expression, we incubated J774 cells for 60 min in the presence of increasing concentrations (0.01, 0.05, 0.1, 0.5, 1, or 5 mmol/l) of Hcys. Whereas Mo LPL mRNA expression remained unchanged in cells exposed to 0.01 and 0.05 mmol/l Hcys, a significant increase in this parameter was found in the presence of 0.1 mmol/l Hcys (Fig. 2A). Maximal levels of LPL mRNA were observed after culture of the cells with 0.5–1 mmol/l Hcys. A similar dose-dependent effect of Hcys on LPL mRNA expression was observed in Mo exposed for 6 h to Hcys (data not shown). Recovery of enhanced amounts of LPL immunoreactive mass from the culture media reflected the increase in LPL mRNA expression of Hcys-treated J774 cells. To determine the dose-dependent effect of Hcys on Mo LPL secretion, we incubated J774 cells for 24 h with increasing concentrations (0.1, 0.2, 0.5, 1, or 5 mmol/l) of Hcys. Whereas a small induction of LPL secretion was already observed in Mo exposed to 0.2–0.5 mmol/l, the effect of Hcys on Mo LPL secretion reached statistical significance only at concentrations of 1–5 mmol/l (Fig. 2B).

Mo exposed for 24–72 h to Hcys (1–5 mmol/l) showed a sustained twofold increase in LPL mass secretion (Fig. 3A). Determination of Mo LPL activity in the supernatants of J774 cells exposed to 5 mmol/l Hcys showed a moderate increase in this parameter (Fig. 3B). In contrast, the levels of extracellular LPL activity in cells treated with 1 mmol/l Hcys remained unchanged (Fig. 3B).

To determine whether PKC may represent the signaling pathway involved in the upregulation of Mo LPL gene expression by Hcys, we pretreated J774 cells for 30 min in the presence or absence of the specific PKC inhibitor Calphostin C (0.5 μg/ml), and then we treated cells for 1 h with 1 mmol/l Hcys. As shown in Fig. 4A, calphostin C totally abolished the stimulatory effect of Hcys on Mo LPL gene expression. A similar effect was observed when J774 cells were pretreated with the PKC inhibitors GF109203X (20 nmol/l) and chelerythrine (0.5 μmol/l) (LPL mRNA levels [percent of control values]: controls: 100.0 ± 1.6; Hcys: 145.2 ± 5.5; GF109203X: 79.0 ± 11.8, GF109203X + Hcys: 73.0 ± 11.1, P < 0.05 versus Hcys; chelerythrine: 95.3 ± 7.2, chelerythrine + Hcys: 94.3 ± 7.7, P < 0.05 versus Hcys).

To establish further the involvement of PKC in Hcys-stimulated Mo LPL mRNA expression, we measured PKC activity in cytosolic and particulate fractions of Hcys-treated J774 cells (Fig. 4B). A maximal increase in PKC activity in the membrane fraction of Mo was observed after a 10-min exposure of the cells to 1 mmol/l Hcys.

To assess the effect of Hcys on Mo c-fos gene expression, we incubated J774 cells for 15 to 60 min with 1 mmol/l Hcys (Fig. 5A). Exposure of Mo for 15 min to Hcys led to a significant induction of basal c-fos gene expression. Maximal effect was observed after a 30- to 45-min incubation period. As illustrated in Fig. 5B, induction of Mo c-fos gene expression was dose-dependent, with a maximal effect occurring at a Hcys concentration of 1 mmol/l.

We next determined whether incubation of J774 cells in the presence of Hcys might induce changes at the level of LPL gene promoter binding proteins. We found that a 15- to 30-min exposure of J774 cells to Hcys led to a major increase in the binding of nuclear proteins to the AP-1 consensus sequence of the LPL promoter. Maximal effect was observed after a 15-min incubation period (Fig. 6). This binding complex was specifically competed in the presence of a 1,000-fold molar excess of the unlabeled AP-1 oligonucleotide.

Hyperhomocysteinemia is an independent risk factor for cardiovascular disease (1–5). Although Hcys has been implicated in many biological events associated with atherogenesis, the mechanism(s) by which this amino acid exerts its proatherogenic effects is still not well known.

Our results demonstrate, for the first time, that Hcys stimulates Mo LPL secretion in vitro. This observation is relevant in light of recent studies showing that Mo LPL production in the arterial wall promotes foam cell formation and atherosclerosis in vivo (26–29). Our finding that Hcys induces to a similar extent LPL mRNA levels and LPL mass suggests that the stimulatory effect of Hcys on Mo LPL secretion may, at least partly, involve transcriptional events. It is widely known that the regulation of LPL gene expression is controlled by several cis- and trans-acting factors surrounding the LPL transcriptional start site (40,41). Induction of LPL gene expression by Hcys may theoretically involve different signaling pathways. On the basis of the presence of an AP-1 site in the regulatory sequence of the LPL gene and of the previously documented effect of Hcys on PKC and c-fos activation (12), one may suggest that increased AP-1 activity resulting from PKC-induced c-fos expression could mediate the stimulatory effect of Hcys on Mo LPL gene expression. Our findings that Hcys led to PKC activation in Mo and that specific PKC inhibitors totally abolished the induction of Mo LPL gene expression in response to Hcys clearly demonstrated that PKC activation is required for the stimulatory effect of Hcys on Mo LPL gene expression. Furthermore, our observations that Hcys induces parallel changes in Mo LPL and c-fos mRNA levels and that Hcys induces the binding of nuclear proteins to the AP-1 sequence suggest a role for c-fos in the transcriptional regulation of Mo LPL gene expression by Hcys. From these observations, one may postulate that one signaling event mediating the increase of the transcriptional rate of the LPL gene in response to Hcys involves the activation of PKC and the interaction of its well-known target c-fos with the AP-1 sequence. Several lines of evidence indicate that PKC activates the mitogen-activated protein (MAP) kinase signal transduction pathway and that induction of MAP kinase activity results subsequently in c-fos expression. Members of the MAP kinase family include the extracellular signal-regulated kinases (ERKs) 1 and 2 and the stress-induced kinases Jun NH₂-terminal kinase and p38. It has been demonstrated that Hcys activates the Jun NH₂-terminal kinase signaling pathway in human endothelial cells (42) and stimulates ERK1 and -2 phosphorylation in smooth muscle cells and NIH/3T3 cells (43–45). On the basis of these findings and on the previously documented role of the MAP kinase pathway in the regulation of adipocyte LPL expression (46), it is tempting to speculate that PKC-dependent MAP kinase activation could mediate the upregulation of Mo LPL gene expression by Hcys. This question is especially relevant considering the close correlation between the reported biphasic activation of ERK, which rises 10 min and 4 h after stimulation (47), and the presently documented Hcys-induced early and delayed stimulation of Mo LPL gene expression that we observed at 1 and 6 h after Hcys exposure, respectively.

Besides c-fos, other transcription factors, such as peroxisome-proliferator activated receptors (PPARs), could mediate the upregulation of Mo LPL gene expression in response to Hcys. Indeed, evidence has been provided that PPARs regulate Mo LPL gene expression (32,48) by binding to a peroxisome-proliferator response element present in the LPL promoter (49). Because Hcys (50), PKC (51,52), and MAP kinases (53–57) regulate the expression and/or activation of PPARs, these transcription factors may play a key role in the upregulation of Mo LPL gene expression in response to Hcys. Our preliminary observations showing that Hcys induces the binding of nuclear proteins to the peroxisome-proliferator response element of the LPL gene support this possibility.

Our results finally demonstrated that Hcys increases Mo LPL secretion. Whereas the increment of LPL mass paralleled the induction in LPL mRNA, increase of LPL activity was not observed in cells exposed to 1 mmol/l Hcys and was well below the reported elevation in the enzyme mRNA levels in Mo treated with 5 mmol/l Hcys. Because the antibody used to measure LPL mass recognized both active and inactive LPL, these findings suggest that Hcys induces the release of mostly inactive LPL. The complex posttranslational processing of LPL allows for the expectation of different sites for this modulation of LPL activity by Hcys. Additional research will be necessary to determine exactly which steps of the process of LPL synthesis are target sites for the modulation of LPL activity by Hcys.

Recent data suggest that Hcys may be a risk factor for cardiovascular disease in the diabetic population. Plasma Hcys concentrations are elevated in patients with diabetes (58,59), and a relationship between hyperhomocysteinemia and increased risk of cardiovascular morbidity and mortality has been established in patients with type 2 diabetes (6–10). We previously demonstrated that Mo LPL, a proatherogenic protein, is increased in patients with type 2 diabetes (30) and that peripheral factors are, at least partly, responsible for this alteration (30–32). Our finding that Hcys increases Mo LPL secretion in vitro suggests that Hcys may contribute to the induction of Mo LPL in human diabetes.

One limitation of the present study is the supraphysiological Hcys concentrations needed to enhance Mo LPL expression in vitro. Indeed, we found that the DL-Hcys concentrations required to stimulate Mo LPL in the present study were >10 times those found in the plasma of patients with diabetes. Because the inactive d-isomer constitutes ∼50% of the DL-Hcys, use of the l-isomer of Hcys exclusively could have lowered the Hcys concentration needed to elicit a biological response. It is also possible that concentrations of Hcys in the limited environment of the atherosclerotic plaque may be sufficient to favor the induction of Mo LPL in vivo. Because LPL exerts its proatherogenic effects through both its structural and catalytic properties (60–65), generation of increased amounts of LPL, even in its inactive form, may contribute to the accelerated atherosclerosis associated with diabetes.

In conclusion, this study demonstrates for the first time that Hcys stimulates Mo LPL gene expression and secretion in vitro. Our data show that induction of Mo LPL mRNA expression by Hcys involves PKC activation and suggest a role of c-fos in the transcriptional regulation of Mo LPL expression. These results provide a new mechanism by which Hcys may promote atherosclerosis in human diabetes.