1 Cardiovascular Research Unit, Center for Molecular Medicine L8:03, Karolinska Institute, S-17176 Stockholm, Sweden; and 2 Institut für Prophylaxe der Kreislaufkrankheiten, D-80336 Munich, Germany
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ABSTRACT |
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Pyrrolidine dithiocarbamate (PDTC) has been found to
induce or inhibit apoptosis in different cell types. Here we show that PDTC dose-dependently reduced the viability of rat smooth muscle cells
(rSMC), human fibroblasts, and endothelial cells at low but not at high
cell density. Endothelial cells were least sensitive, fibroblasts
showed a medium sensitivity, and rSMC showed a high sensitivity to
PDTC-mediated cell death. An early reduction in the mitochondrial
membrane potential indicated a rapid onset of apoptosis in rSMC.
Apoptosis was further confirmed by annexin V staining and DNA
fragmentation analysis. Gel shift analysis demonstrated increased
nuclear factor (NF)-B activity in high-density rSMC compared with
low-density cells. NF-
B has recently been shown to regulate the
induction of anti-apoptotic proteins. Although PDTC is widely used as
an inhibitor for NF-
B and a radical scavenger, our data show that
PDTC rather enhanced NF-
B activity and, alone or in combination with
menadione, induced oxygen radical generation. Notably, PDTC failed to
reduce rSMC viability in medium without Cu2+ or
Zn2+, and addition of Cu2+ or Zn2+
resulted in a dose-dependent increase in PDTC-induced cell death. Addition of both Cu2+ and Zn2+ showed
synergistic effects. Our results indicate that the induction of
apoptosis by PDTC requires Cu2+ and Zn2+ and is
dependent on cell type and density. Such differential effects may have
implications for studies of PDTC as an anti-atherosclerotic or
immunomodulatory drug.
cell density; smooth muscle cells; copper ion; zinc ion
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INTRODUCTION |
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APOPTOSIS (14) is important in normal development, and
a dysregulation in programmed cell death may cause or contribute to
various diseases such as atherosclerosis, autoimmunity, or cancer (11,
30, 32). Radicals play an important role in apoptosis, and both
induction of apoptosis by reactive oxygen species (ROS) and ROS
production by apoptotic cells have been described (18, 26, 37, 39). The
antioxidant and radical scavenger pyrrolidine dithiocarbamate (PDTC)
has been shown to potently inhibit agonist-induced apoptosis in
lymphocytes and tumor cell lines (4, 26). In contrast, other studies
demonstrated the induction of apoptosis by PDTC in different cell
types, such as hepatocytes, thymocytes, smooth muscle cells (SMC), or
murine B cells (2, 22, 35, 38). Although the protective action is
mainly attributed to antioxidative properties of PDTC, hypothetical causes for apoptosis induction range from antioxidative (4, 7) or
oxidative (22) to inhibition of nuclear factor (NF)-B (2, 38) or
cell cycle proteins (7, 19). Another report claimed that metabolites of
dithiocarbamates, but not PDTC or diethyldithiocarbamate themselves,
induce apoptosis through coordinate modulation of NF-
B,
c-fos/c-jun, and p53 proteins (19). PDTC is a metal
chelator, and an increase of redox reactive Cu2+ has also
been held responsible for PDTC-induced apoptosis (5, 22). Although
endothelial cells have previously been shown to be resistant to
treatment with PDTC (35), a more recent report has demonstrated
induction of endothelial cell death by PDTC (15).
These contradictory findings in various cell types prompted us to
further investigate potential mechanisms of PDTC-induced cell death in
different adhesive, primary cells, namely rat vascular SMCs (rSMC),
human umbilical vein endothelial cells (HUVEC), and human fibroblasts
(hFB) with respect to their sensitivity to PDTC. A reduction of the
mitochondrial membrane potential () has been described as a
reliable marker to detect an early point of no return in the apoptotic
cascade (6), followed by an increase in production of intracellular ROS
(39). Because early events in PDTC-induced apoptosis and the role of
ROS are not well characterized, we used measurement of
and ROS
generation, as well as annexin V binding and DNA fragmentation, to
further investigate PDTC-mediated cell death.
In the present study, the susceptibility to induction of cell death by
PDTC was compared in different cell types at defined cell densities,
parameters that have not previously been studied. We show that cell
type and density crucially affect the sensitivity to induction of cell
death. The activity of NF-B and oxygen radical generation in rSMC
was rather enhanced by PDTC, and the presence of Cu2+ and
Zn2+ was required for PDTC activity, thereby excluding
antioxidative effects while confirming alternative mechanisms of PDTC
in triggering apoptosis.
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MATERIALS AND METHODS |
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Cell culture. Rat aortic SMC were harvested from male Sprague-Dawley rats by enzymatic digestion, as described (8), and were grown in DMEM/Ham's F-12 (F12; 1:1; all media from Life Technologies) with 10% FCS and 50 µg/ml gentamicin (Life Technologies). For some experiments, rSMC were incubated in Eagle's MEM. HUVEC were isolated as described (9) and were cultured in medium 199 with 10% FCS, 5% human serum, 25 µg/ml endothelial cell growth factor (Boehringer Mannheim), and antibiotics. hFB were purchased from Clonetics and were cultured in HUVEC medium. Cells were passed every 5-10 days and were seeded at a density of 1 × 104 cells/cm2 for propagation in T-75 flasks or at the specified densities 48 h before the respective treatment in 24-well plates and T-75 or T-25 flasks. Cell numbers were adjusted when comparing low- and high-density cells. Experiments were performed on cells 4-10 passages from primary culture. For comparing SMC, HUVEC, and hFB under identical conditions, DMEM/F12 culture medium (10% FCS) was used for treatment with PDTC. To incubate SMC with defined concentrations of Cu2+ and Zn2+, MEM that is essentially free of Cu2+ or Zn2+ was used. PDTC, CuSO4, and ZnSO4 were dissolved in water, sterile filtered, and stored at 4°C. All other reagents were from Sigma Chemical
Cell viability assays. Cell viability was determined by a modified 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) reduction assay (8). For the MTT test, cells were grown in 24-well plates and washed with PBS, and 300 µl MTT solution (0.5 mg/ml) in culture medium (0.5% FCS) was added to each well. After 3 h of incubation at 37°C, MTT was discarded, and the purple formazan product in the cells was solubilized by addition of acidic isopropanol (0.04 M HCl). After lysing for 10 min, 2 × 100 µl from each well were transferred to a 96-well plate, and absorbency was analyzed in an ELISA reader at 570 nm. Absorbency at 690 nm was used to correct for background. MTT activity was expressed as the percentage of untreated controls. To validate the MTT assay as a measure of cell number and viability, MTT and trypan blue assays were performed side by side for all cell types and PDTC concentrations used.
Measurement of and oxygen radical
production.
Detection of apoptotic and nonviable cells was achieved by a
multiparameter assay measuring a decrease in
, positive staining for propidium iodide (PI+), and oxygen radical generation
(8, 39). To measure
, cells were incubated with
3,3'-dihexyloxacarbocyanine iodide (DiOC; Molecular Probes) at
37°C for 15 min (16 nM; 525 nm), followed by immediate analysis of
fluorochrome incorporation in a Becton-Dickinson FACSCalibur flow
cytometer. Each sample was incubated with propidium iodide (PI) 10 s
before analysis (1 µg/ml; 600 nm) to detect nonviable cells.
Hydroethidine (HE; 2 µM, 15 min at 37°C; Molecular Probes) was
used to measure superoxide radical (ROS) generation. In control experiments, cells were treated with the ROS-generating agent menadione
(100 nM) or in the presence of carbonyl cyanide
m-chlorophenylhydrazone (50 µM, data not shown) to induce
breakdown. An acquisition gate was set to exclude cell debris
and aggregates, and 10,000 cells within this gate were analyzed.
Detection of apoptosis by annexin V staining and DNA fragmentation analysis. After treatment with PDTC, SMC were washed with binding buffer and incubated with FITC-labeled annexin V in binding buffer (annexin V Kit; Immunotech) for 20 min at 20°C in the dark. Cells were washed three times, and adherent cells were harvested by scraping, pooled with nonadherent cells, resuspended in PBS with 1.5 mM Ca2+ and 1% FCS, and fixed with 1 ml 2% paraformaldehyde to prevent cell aggregation. SMC were analyzed by flow cytometry using a single cell gate, and the percentage of annexin V-positive cells was determined after setting appropriate markers for negative and positive populations (16). To quantify DNA breakdown by endonuclease activity, we stained isolated nuclei with PI and measured the percentage of apoptotic nuclei with DNA fragmentation by cell cycle analysis. Apoptotic cells with a DNA content <2 N appear in the sub-G1 region (8, 29). Cells were grown for 48 h and were treated with PDTC for 24 h. Cultures were scraped, and adherent and nonadherent cells were pooled, washed, resuspended in 0.5 ml PBS, and fixed with 5 ml of fluorescence-activated cell sorter (FACS) lysing solution (Becton-Dickinson). Cells were then processed using the Cycle TEST PLUS DNA Reagent Kit (Becton-Dickinson). After being washed with 10 ml PBS and 2 × 1 ml citrate buffer, cells were lysed in 250 µl buffer A for 10 min, and 200 µl buffer B were added for 10 min. Nuclei were stained with PI solution for 5 min and were analyzed in a FACSCalibur with instrument settings from the DNA Quality Control Kit (Becton-Dickinson) in activated doublet discrimination mode and in a single nuclei gate. Histograms were analyzed by manual gating, setting a marker (M1) on the sub-G1 peak.
Preparation of nuclear extracts and electrophoretic mobility shift
assay.
Electrophoretic mobility shift assay (EMSA) was performed as described
(8, 9). Briefly, cells in T-25 flasks were rinsed with ice-cold PBS and
harvested in 1 ml of ice-cold buffer (20 mM HEPES, 22% glycerol, 20 mM
KCl, 1.5 mM MgCl2, 0.2 mM EDTA, 1 mM dithiothreitol, and 1 mM phenylmethylsulfonyl fluoride). Isolated nuclei were resuspended in
41 µl of 20 mM KCl buffer, and 39 µl of 600 mM KCl buffer were
added. Nuclear proteins were extracted by incubation on ice for 30 min.
After centrifugation for 15 min at 8,000 g, supernatants were
transferred to precooled tubes, and the protein concentration was
determined spectrophotometrically. Nuclear protein (10 µg) was mixed
with a double-stranded oligonucleotide corresponding to a NF-B
binding motif (5'-AGTTGAGGG GACTTTCCCAGGC-3'; Santa Cruz)
and was labeled with [32P]dATP using T4
polynucleotide kinase. After binding for 15 min, samples were separated
on nondenaturating 4% polyacrylamide gels and exposed to X-ray films.
An excess of unlabeled oligonucleotide and antibodies to the p50 and
p65 subunits were used to identify specific bands (data not shown). The
absorbency of specific NF-
B bands on autoradiograms was analyzed by
laser densitometry and was given as optical density units times millimeters.
Statistical analysis. Values are given as means ± SD. A paired Student's t-test was used to analyze for statistical differences. P values <0.05 were considered significant. To test for synergistic effects, a factor was calculated using the following equation: %decrease MTT (Cu2+ + Zn2+)/%decrease MTT (Cu2+) × %decrease MTT (Zn2+). Values greater than one are indicative of a synergistic effect.
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RESULTS |
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Cell density affects induction of cell death by PDTC in rSMC, hFB,
or HUVEC.
Treatment of rSMC with PDTC for 24 h dose dependently reduced cell
numbers and viability, as detected by MTT assays (Fig. 1A) and trypan blue exclusion (Fig.
2A). This was consistent with findings that exponentially proliferating SMC but not confluent endothelial cells are sensitive to induction of apoptosis with PDTC
(35). Because endothelial cells grow to a confluent monolayer, whereas
SMC grow in overlying layers exhibiting a typical hill-and-valley pattern, we compared rSMC, HUVEC, and hFB seeded at three defined densities for their sensitivity to induction of cell death by PDTC.
Treatment with PDTC for 24 h dose dependently reduced MTT activity in
all three cell types, but with marked differences in sensitivity
depending on cell type and seeding density. At 1 or 10 µM, rSMC but
not HUVEC or hFB showed up to 75% reduction in MTT activity (Fig.
1A) and trypan blue cell count (Fig. 2A) at a density
of 0.5 × 104 cells/cm2, indicating that
rSMC were most sensitive to induction of cell death. Moreover, HUVEC
and hFB were sensitive to PDTC treatment at low cell densities before
reaching confluence (Figs. 1 and 2). Under certain conditions such as a
high cell density, PDTC may exert growth stimulatory effects (Fig. 1).
Treatment with N-acetylcysteine (NAC; 25 mM, 24 h) did not
induce cell death in rSMC, despite resulting in a 25 ± 3% reduction
in MTT activity. Microscopic inspection revealed no evidence for
induction of cell death by NAC up to 48 h of incubation (data not
shown). To validate the MTT assay as a measure for cell number and
viability, we performed trypan blue exclusion assays in side-by-side
experiments. In all cell types, PDTC induced a dose-dependent reduction
in cell number and an increase in trypan blue positive cells (Fig. 2).
Although SMC showed a marked increase of trypan blue positive cells
(70-95%) at relatively low concentrations of PDTC at the lowest
density tested (Fig. 2A), treatment of HUVEC or hFB
resulted in a reduction of cell count with a only fraction of trypan
blue positive cells (10-40%), even at the highest concentrations
of PDTC (Fig. 2, B and C). This suggests that in HUVEC
or hFB PDTC treatment results primarily in reduced proliferation and
partial induction of cell death.
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PDTC effects in SMC are dependent on serum and show an early onset.
Serum removal is known to induce apoptosis (21), whereas reducing serum
concentrations to 0.1% does not induce apoptosis in SMC (24). Hence,
we tested whether the effects of PDTC on SMC viability are modulated by
different serum concentrations. Incubation of rSMC in medium with 0.1%
FCS enhanced the PDTC-mediated reduction in viability (Fig.
3), suggesting that serum components provide survival signals. To determine the minimal incubation time
required for induction of SMC death, we removed PDTC at defined time
points by threefold washing and incubated the cells for up to 24 h in
medium containing 0.1 or 10% FCS. Again, low serum conditions rendered
the cells more sensitive to PDTC with ~70% reduction in viability
after 1 h of exposure, whereas in 10% FCS incubation for 2 h or longer
was required to induce substantial cell death (Fig. 3). In both cases,
the effect was almost maximal at 6 h of exposure to PDTC.
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Effects of PDTC on and membrane
permeability in low-density SMC.
To verify whether PDTC induces early apoptotic changes in rSMC, we used
a flow cytometric multiparameter analysis to detect sequential
alterations in
and plasma membrane integrity. As determined by
DiOC staining, we found a dose- and time-dependent reduction in
after treatment of rSMC with PDTC at 10, 100, or 1,000 µM (data not
shown) or with 100 µM PDTC for 2, 4, or 6 h (Table
1). Moreover, we observed an increase in PI
staining, indicating plasma membrane leakage and subsequent cell death. The
reduction was accompanied by an increase in PI+
cells (Table 1), in accordance with previous studies in lymphocytes (6,
39). As observed for MTT activity (Fig. 1A), high-density rSMC
did not exhibit signs of cell death or apoptosis, such as a reduced
or increased staining for PI after PDTC treatment, even in
reduced serum conditions that potentiated PDTC effects in low-density
SMC (Fig. 4A).
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Induction of apoptosis in low-density SMC by PDTC.
To confirm that the reduction of viability and was due to
apoptosis, we used FITC-conjugated annexin V, which binds to phosphatidylserine that is translocated from the inner to the outer
leaflet of the cell membrane and therefore is exposed when a cell
enters apoptosis (16). Consistent with reduced viability and
,
flow cytometric analyses demonstrated that treatment of low-density SMC
with 100 µM PDTC time dependently increased the percentage of annexin
V-positive cells from 3 ± 1% in control cells to 15 ± 4% after 6 h and to 80 ± 9% after 24 h (Fig. 4B). High-density SMC
showed slightly increased annexin V binding in the untreated condition
compared with low-density cells, which were not increased by treatment
with PDTC (Fig. 4B). Treatment of low-density cells with 10 mM
NAC for 24 h did not increase binding of annexin V, excluding
proapoptotic effects of antioxidative compounds (Fig. 4B). To
verify that PDTC induces typical apoptotic changes of nuclear DNA and
to evaluate DNA fragmentation in a quantitative setting, we determined
the percentage of cells with a DNA content <2 N appearing in a sub-G1
region by flow cytometric cell cycle analysis (8, 29). Treatment of SMC
with PDTC (100 µM) for 24 h resulted in a marked sub-G1 peak (42 ± 12 vs. 2 ± 1%, mean ± SD, n = 4) indicative of an
apoptotic subpopulation (Fig. 4C), whereas PDTC treatment of
high-density cells, which showed slightly increased basic apoptosis,
did not result in a substantial increase of the sub-G1 peak (10 ± 5 vs. 5 ± 2%, n = 3; Fig. 4C). Treatment of hFB or
HUVEC with 100 µM PDTC for 24 h did not result in an
increased sub-G1 peak but showed a marked shift from the G2/M to the
G0/G1 phase or the G0/G1 to the G2/M phase, respectively. This suggests
that PDTC reduces proliferation of hFB or HUVEC (Fig. 4C),
which would largely account for the reduction in MTT activity, and was
also seen from the trypan blue data (Figs. 1 and 2). These data clearly
indicate that PDTC induces characteristic signs of apoptosis in low-
but not in high-density SMC. In hFB or HUVEC, treatment with PDTC
primarily resulted in reduced proliferation and was accompanied by the
occurrence of cell death.
NF-B activity in low- and high-density SMC.
To elucidate possible mechanisms of PDTC-induced cell death, we focused
on rSMC because these cells were most sensitive to PDTC treatment.
First we looked at differences in cell density. Seeding rSMC at 10 or
15 × 104 cells/cm2 resulted in 10 ± 0.5 × 104 cells/cm2 after 48 h of culture,
whereas seeding at 104 cells/cm2 equally
resulted in 10 ± 0.7 × 104 cells/cm2
after 2 wk of culture, indicating that this level constitutes a density
maximum for rSMC under in vitro culture conditions. Inhibition of
NF-
B has been reported to induce apoptosis in different cell types
(2, 31, 38) that share a constitutive NF-
B activity with SMC (17).
Hence, we analyzed NF-
B activity in rSMC at low and high density
(Fig. 5A). Interestingly, EMSA
demonstrated a higher NF-
B activity in rSMC in high-density as
opposed to low-density cells (Fig. 5B, P < 0.025).
Because we have recently shown that inhibition of NF-
B induces
apoptosis in SMC (8) and PDTC is a potent inhibitor of NF-
B
mobilization in a variety of cell types (27, 36, 40), we investigated
whether the induction of apoptosis in rSMC by PDTC may be due to
inhibition of NF-
B. Surprisingly, NF-
B in low-density cells was
increased by 4 h of PDTC treatment (Fig. 5B), whereas the basic
NF-
B activity was slightly reduced by treatment with PDTC in
high-density cells (Fig. 5B).
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Oxygen radical generation after PDTC treatment.
Free radicals play an important role in apoptosis. Both induction of
apoptosis by ROS in SMC and ROS production by cells undergoing apoptosis have been described (18, 39). To measure intracellular ROS,
we used HE, a substance showing increased fluorescence upon reaction
with intracellular oxygen radicals that can be detected by FACS.
Because PDTC is a potent antioxidant and radical scavenger, we tested
its effect on radical production in rSMC. Surprisingly, PDTC resulted
in an induction of ROS at 4 or 6 h of treatment (Table 1) that was more
pronounced in reduced serum conditions (data not shown). At 2 h, the
reduction of was not accompanied by an increased ROS generation
(Tables 1 and 2). Incubation of SMC with
menadione, a substance known to induce oxygen radicals, decreased
after 1 h of incubation, preceding an increase in ROS (Table 2).
PDTC treatment did not decrease
or significantly increase ROS
generation after 1 h (Table 2). Interestingly, a combination of
menadione and PDTC further increased the percentage of cells showing
ROS formation and low
(Table 2), excluding antioxidant or
radical scavenging activities of PDTC in our experimental setting. This
indicates that ROS generation may be a consequence rather than a cause
of apoptosis induction by PDTC.
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Effects of PDTC are mediated by
Cu2+ and
Zn2+.
PDTC is known as a metal chelator. Chelation of extracellular
Cu2+ by PDTC and subsequent transport into the cells as
PDTC-metal complexes induced apoptosis in thymocytes (22). The
Cu2+ content in the DMEM/F12 culture medium used was 5.3 nM. To test whether Cu2+ was crucially
involved in PDTC-induced rSMC death, we used serum-free MEM without
Cu2+. In contrast to the varying degrees of PDTC-mediated
cell death in DMEM/F12 with 0.1% FCS or 10% FCS (Fig. 3), incubation
of rSMC with 10 or 100 µM PDTC in serum-free MEM for 24 h did not
reduce viability, as assessed by MTT activity (Fig.
6A) and phase-contrast microscopy
(Fig. 7). The addition of Cu2+
alone to MEM had no effect but resulted in a dose-dependent increase in
SMC death in the presence of 10 µM PDTC (Fig. 6A). Addition of Cu2+ at 0.1 µM resulted in a less marked reduction of
viability by PDTC in MEM (39 ± 6%, mean ± SD, n = 3) than in serum-free DMEM/F12 culture medium (91 ± 5%, mean ± SD, n = 4). This inferred that components other than
Cu2+ contributed to PDTC effects. Hence, we focused on
differences in the concentration of Zn2+ between DMEM/F12
(1.5 µM) and MEM (without Zn2+). As found for
Cu2+, addition of Zn2+ to MEM resulted in a
dose-dependent increase in PDTC-induced SMC death (Fig. 6B).
The IC50 for Cu2+ and for Zn2+ was
~0.3 and 0.45 µM, respectively. The combination of Cu2+
(0.03 µM) and Zn2+ (0.3 µM) revealed a synergistic
effect (Fig. 6C) that was reflected by a synergy factor of 1.87 ± 0.43 compared with 1.0 ± 0.14 for untreated control cells
(P < 0.01). Interestingly, addition of 10% FCS containing
Cu2+ and Zn2+ cations to MEM restored the
susceptibility for induction of cell death by 100 µM PDTC (Fig. 7,
E and F). Hence, the combination of PDTC with metal
cations in serum may shift the apoptotic balance toward cell death,
despite anti-apoptotic effects of survival factors in serum that were
evident from the serum reduction experiments (Fig. 3). These data
suggest that Cu2+ and Zn2+ are the components
crucially required for mediating PDTC-induced cell death.
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DISCUSSION |
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The induction of apoptosis by PDTC has been described in several cell
types (2, 7, 22, 35, 38); however, the data were inconclusive with
respect to responsible mechanisms. We now demonstrate that PDTC dose
dependently reduced the cell number and viability of rSMC, HUVEC, and
hFB at low but not high cell density. SMC were most sensitive, HUVEC
were least sensitive, and hFB showed intermediate sensitivity. A rapid
onset of PDTC effects was supported by an early reduction in .
NF-
B activity and radical generation were stimulated by PDTC in
low-density rSMC. Notably, PDTC effects were dependent on the presence
of Cu2+ and Zn2+.
Treatment with PDTC for 24 h dose dependently reduced viability in rSMC, HUVEC, or hFB. This is consistent with previous reports showing that PDTC induces apoptosis in colorectal tumor, PC-12, SMC, or B cells (2, 7, 35, 38). On the other hand, studies show inhibition of agonist-induced apoptosis by PDTC in HL-60 or T cells (4, 26). A possible explanation for these discrepancies could be the use of different cell types or culture conditions. PDTC was shown to inhibit or induce apoptosis in PC-12 cells (31, 35), indicating a dual role of PDTC, depending on assay conditions. We found that endothelial cells are sensitive to induction of cell death by PDTC at lower cell density, whereas confluent cells were not affected, providing a possible explanation for contrasting observations on PDTC-induced cell death in this cell type. Our data implicate that the susceptibility to induction of apoptosis may depend on factors such as cell-cell contact, proliferation, or the presence of survival factors. Consistent with the latter idea, low serum conditions did not induce cell death but rendered SMC more sensitive to PDTC. This complies with findings that only complete serum deprivation but not reduction to 0.1% results in apoptosis of SMC (24).
A rapid onset of PDTC-induced apoptosis at low density was supported by
the dose- and time-dependent reduction of in SMC in accordance
with studies in lymphocytes (6, 39). Apoptosis of SMC was confirmed by
increased binding of annexin V to phosphatidylserine exposed on the
cell membrane after induction of apoptosis (16). In addition, DNA
fragmentation was quantified by flow cytometric cell cycle analysis.
After 24 h, almost 50% of nuclei exhibited a DNA content <2 N,
clearly indicating massive induction of apoptosis by PDTC in low- but
not in high-density rSMC. In contrast, hFB or HUVEC showed a shift in
the cell cycle, indicating marked reduction of proliferation with a
fraction of dead cells, as detected by trypan blue. Our data add to
recent findings that proposed resistance of endothelial cells to PDTC
treatment (35) and demonstrate a crucial role of cell density in the
susceptibility to apoptosis.
To find an explanation for the different reaction to PDTC, we further
focused on differences associated with cell density. Although HUVEC or
hFB stop to proliferate at confluence, SMC grow on postconfluence,
forming multiple cell layers. EMSA demonstrated a higher NF-B
activity in rSMC at high density as opposed to low density. This adds
to the finding that SMC show NF-
B activity under normal culture
conditions, which is essential for proliferation (3, 17). The
constitutive activity of NF-
B has been ascribed to serum components,
and serum is a potent inducer of NF-
B in SMC (23). The higher
NF-
B activity in high-density cells in our study was not due to
serum components because identical medium composition was used for low-
and high-density cultures. Serum reduction to 0.1% for 24 h resulted
in a significant decrease of NF-
B in low-density rSMC, whereas
NF-
B in high-density cells was hardly affected by this treatment
(data not shown), indicating a serum-independent increase of NF-
B in
high-density cells. Interestingly, fibronectin, which is produced in
large amounts by SMC (12, 33), has been shown to stimulate NF-
B in
SMC (25). In addition, cell-cell contact-dependent effects, such as
cell activation by integrin signaling, may contribute to NF-
B
activation in high-density cells growing in overlying layers. Because
we have recently demonstrated that inhibition of NF-
B induces
apoptosis in SMC and PDTC is a potent inhibitor of NF-
B induction in
a variety of cell types (27, 36, 40), we investigated whether the
effects of PDTC were due to inhibition of NF-
B. Surprisingly,
NF-
B in low-density cells was not inhibited but rather was
stimulated by PDTC, consistent with previous reports showing NF-
B
mobilization after induction of apoptosis (10, 28). Constitutive
NF-
B activity in thymocytes is increased by stimuli that promote
apoptosis and is not inhibited by PDTC (28). We have shown that the
expression of anti-apoptotic proteins is controlled by NF-
B and is
consequently increased in high-density human SMC (8), providing a
possible explanation for protection of high-density SMC from induction
of apoptosis.
Free radicals are involved in the apoptotic process. Surprisingly,
PDTC, which is a strong antioxidant, caused increased radical production in rSMC. This indicates that ROS generation may be a
consequence rather than a cause of apoptosis induction by PDTC. We have
found that the decrease in precedes an increase in ROS
production. Because HE only detects oxygen radicals, the effects of
PDTC may be mediated by other radical species. In fact, a recent report
shows that PDTC induces apoptosis in thymocytes without release of ROS
(5). In addition, menadione, a substance known to induce intracellular
oxygen radical generation by redox cycling, decreased
after 1 h
of incubation, preceding an increase in ROS. The combination of
menadione and PDTC further increased the percentage of cells with ROS
formation. Interestingly, we did not find induction of cell death by
NAC, a structurally unrelated antioxidant. Although annexin V binding
was not increased after treatment with NAC for 24 h, we could detect a
25% reduction in MTT activity and trypan blue cell count that is
comparable to values reported previously (35). An absence of trypan
blue positive cells, as has been similarly demonstrated with NAC in
bovine aortic SMC (3), indicated that this reduction in MTT activity
was most probably due to decreased proliferation and was not due to induction of cell death. Taken together, this infers that properties of
PDTC other than antioxidative radical scavenging are crucial for the
effects observed in our study.
The antioxidant potential is a common feature of PDTC and NAC, but, in contrast to NAC, PDTC is also a metal chelator. We found that depletion of Cu2+ and Zn2+ abrogated PDTC effects in SMC and that the repletion of Cu2+ or Zn2+ by addition separately, in combination, or contained in serum dose dependently restored the potential of PDTC to induce cell death. Metal chelation by PDTC has been shown to be responsible for induction of apoptosis in thymocytes by raising intracellular levels of redox-reactive Cu2+ (22). In line with this observation, we found that PDTC induced cell death in the presence of comparable low amounts of Cu2+, although Cu2+ alone had no effect. In accordance with a study in cerebral endothelial cells (15), we now demonstrate that Zn2+ contributed to PDTC-induced SMC death. Decreasing intracellular Zn2+ by specific chelators has been reported to induce apoptosis in lymphocytes and to increase the apogenic potential of apoptosis inducers (13, 34), whereas raising intracellular Zn2+ levels has been postulated to exert anti-apoptotic effects (1). A key role for intracellular Zn2+ in controlling nuclear DNA cleavage has been suggested (1, 13, 34). In contrast, a recent study has reported that a PDTC-mediated rise in intracellular Zn2+ is responsible for the induction of endothelial cell death (15). In parallel, our data indicate that comparable low amounts of Zn2+ in a complex with PDTC may induce apoptosis in SMC. Because PDTC did not induce cell death in the absence of extracellular Zn2+, chelation of extracellular but not intracellular Zn2+ appears to be relevant for effects described herein. Why PDTC, in contrast to other Zn2+-chelating agents (13, 34), does not appear to bind intracellular Zn2+ remains to be investigated. Our findings suggest that induction of apoptosis was not due to a PDTC-mediated increase in intracellular Zn2+. Instead, dithiocarbamate-induced cell death may require its conversion to active metabolites, i.e., thiuram disulfides (19). Such a conversion has been demonstrated in vitro (5). It is unclear whether the simultaneous presence of Cu2+ and Zn2+ in living cells can facilitate or accelerate the metabolism of PDTC to thiuram disulfides, e.g., by enzymatic catalysis, which would explain the marked reaction of low-density rSMC to as low as 1 µM of PDTC. The ineffectivity of PDTC to induce apoptosis in Hep G2 cells (19) could indicate that these cells lack such a converting pathway, whereas tumor cell lines may be more sensitive to PDTC than primary cells (Ref. 7 and Erl, unpublished observations).
In conclusion, our data show that Cu2+ and Zn2+ are mediating PDTC-induced death at low cell density. Cells at confluence or high density are protected from PDTC effects. The results may have implications for further studies on PDTC as a potential anti-cancer, anti-atherosclerotic, or immunomodulatory drug (7, 20, 36). In addition to the role of PDTC, which can induce or inhibit apoptosis, our data suggest that Zn2+ may have similar properties, and future studies are needed to clarify the role of Zn2+ in the apoptotic process.
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ACKNOWLEDGEMENTS |
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We thank A. Olsson for performing EMSA.
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FOOTNOTES |
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This study was supported by grants from the Swedish Medical Research Council (project 6816), the Heart-Lung Foundation, the Cancer Society, the Johnson Foundation, Deutsche Forschungsgemeinschaft (Er270/1-1 to W. Erl and We1913/2-1 to C. Weber), and an exchange program of Deutscher Akademischer Austauschdienst and Svenska Institutet.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: W. Erl, Institut für Prophylaxe der Kreislaufkrankheiten, Pettenkoferstr. 9, D-80336 Munich, Germany (E-mail werl{at}klp.med.uni-muenchen.de).
Received 17 August 1999; accepted in final form 15 December 1999.
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