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Introduction
The vascular endothelium serves as the key barrier
between the intravascular compartment and extravascular
tissues, and plays a critical role in a large number of
physiological and pathological
processes[1]. The progressive dysfunction of the endothelium is regarded as a common
deno-minator of impaired organ function after a variety of
pathological tissue alterations such as ischemia, shock, or
sepsis[2].
Lipopolysaccharide (LPS) is a critical glycolipid
component of the outer wall of Gram-negative bacteria.
LPS can induce apoptosis in bovine and ovine endothelial cells
in vitro. When the expression of new genes is inhibited using
cycloheximide (CHX), human endothelial cells become
responsive to the apoptotic signals induced by
LPS[6]. The induction of reactive oxygen species (ROS) by LPS
correlates with the onset of apoptosis, and agents that inhibit the
formation of ROS are known to protect against LPS-induced
endothelial cell apoptosis[1,3].
The phenylethanoid 3,5-dicaffeoylquinic acid
(3,5-diCQA, a white power, Mw 516.44, purified by CC and
HPLC) is present in a variety of plants, including
green coffee beans (Lactuca
indica)[4,5], and is an important
component of the Chinese herb Erigeron
breviscapus. Apart from functioning as a free radical scavenger, 3,5-diCQA
inhibits tyrosinase activity and has a neuroprotective
capacity[5,14]. Anti-oxidant or anti-apoptotic activity has not
previously been described for 3,5-diCQA in endothelial cells.
In the present study, we investigated the effect of 3,5-diCQA
on lipid peroxides in microsomes, LPS-stimulated ROS
generation, and LPS-induced injury in human dermal
microvascular endothelial cells (HMEC-1).
Materials and methods
Reagents 3,5-diCQA was a gift from the Kunming
Institute of Botany (Chinese Academy of Sciences, Kunming
Shanghai, China). The purity of the compounds was greater
than 95%, as verified by HPLC. LPS (Escherichia
coli 0111:B4), CHX, BSA, 3-(4,5-dimethylthiazol-2-yl)-2,5-
diphenyl-tetrazolium bromide (MTT) and MCDB-131 medium were
purchased from Sigma (St Louis, MO, USA).
2,7-dichloro-dihydrofluorescein diacetate (DCFH-DA) was obtained from
Beyotime Institute of Biotechnology (Haimen, China). The
lactate dehydrogenase (LDH) and malondialdehyde (MDA)
assay kits were purchased from the Nanjing Jiancheng
Bioengineering Institute (Nanjing, China). The CaspACE
assay system colorimetric kit was from Promega (USA). All
other reagents or drugs were of analytical grade.
Preparation of microsomes and assay of lipid peroxides
The microsomes were prepared from rat livers. The liver was
cut into small pieces and homogenized in cold buffer A
[(0.l mol/L sucrose, 0.05 mol/L KCl, 0.04 mol/L
KH2PO4, and
0.03 mol/L EDTA (pH 7.2)]. After centrifugation
(1000×g, 20 min), the supernatant was transferred to another tube and
recentrifuged at 105 000×g for 60 min. The sediment was
washed and resuspended in buffer A. All operations were
carried out at 4 °C[9]. Lipid peroxidation of microsomes was
measured as described previously[17] with the following
modifications: the membranes, at a concentration of 1.5 mg
protein, were incubated at 37 °C with 0.05 mol/L phosphate
(pH 7.4) and 0.4 mmol/L ascorbic acid. The final volume was
1.5 mL. Phosphate buffer was contaminated with sufficient
iron to provide the necessary ferric iron for lipid peroxidation.
Control membrane preparations were performed in parallel in
the absence of ascorbate. Lipid peroxidation was initiated
by incubating the membrane preparations in a water bath at
37 °C for 30 min. Lipid peroxidation was assessed by
detection of thiobarbituric acid-reactive MDA, an end product of
the peroxidation of polyunsaturated fatty acids and related
esters. The level of MDA was determined using the MDA
assay kit. The results were normalized to the level of MDA
equivalents/mg of protein.
Cell culture and drug treatment HMEC-1 were
maintained in a 5% CO2 atmosphere at 37 °C in MCDB-131
medium supplemented with 10% FBS and 10 µg/L human
recombinant epidermal growth factor. The cells treated with
CHX (50 mg/L) were used as controls unless otherwise
specified. Cell injury was induced by incubation with LPS
(100 µg/L) and CHX (50 mg/L). 3,5-diCQA was added to the
cells 1 h prior the addition of LPS and CHX. All experiments
were carried out on confluent cultures.
Assay of intracellular ROS The measurement of
intracellular ROS was based on the ROS-mediated conversion of
non-fluorescent DCFH-DA into DCFH. DCFH is membrane
impermeable and rapidly oxidized to the highly fluorescent
2,7-dichlorofluorescein in the presence of intracellular
ROS[23]. The intensity of fluorescence reflects the level of oxidative stress.
Intracellular ROS was measured after incubation in LPS
and/or CHX for 12 h by replacing the medium with 50 mmol/L
phosphate buffer (pH 7.4) containing 10 µmol/L DCFH-DA,
and incubating in the dark for 1 h at 37 °C. The level of DCFH
fluorescence was measured at an emission wavelength of
530 nm and an excitation wavelength of 485 nm by flow
cytometry[19]. The results were expressed as the percentage
of control fluorescence intensity.
Assay of cell survival and cell damage To quantitate the
proportion of viable cells, HMEC-1 were seeded onto
96-well plates at a density of 40 000 cells/well. The cells were
incubated in LPS and/or CHX for 16 h. The viable cell
number was estimated using the MTT assay. Briefly, the
medium was removed and replaced with Krebs (120 mmol/L
NaCl, 5.6 mmol/L KCl, 1.2 mmol/L MgSO4, 1.2 mmol/L
NaH2PO4, 25 mmol/L
NaHCO3, 10 mmol/L glucose, and 2.5 mmol/L
CaCl2) containing 1% BSA and 1 g/L MTT, and then
incubated for 5 h. The medium was aspirated and the
formazan product was solubilized with DMSO. Absorbance
at 630 nm (back-ground absorbance) was subtracted from
absorbance at 570 nm for each well. Viability was expressed
as a proportion of the viable cell number incubated without
LPS[6,15]. Both released and total LDH concentrations were
determined. The release of LDH from cells is a pathological
manifestation of increased plasma membrane permeability.
The total LDH was determined after incubation in the
presence and absence of LPS for 16 h by releasing the compound
from the cells with 1% Triton-X 100 and incubating for 30
min at 37 °C. The level of LDH was determined in samples of
culture medium (30 µL) with or without Triton-X 100 using
the LDH detection kit. The activity of LDH was corrected for
the volume, and released LDH activity was expressed as a
percentage of total cellular LDH[8].
DNA fragmentation assay After incubation in the
presence or absence of LPS for 16 h, the cells
(5×105) were harvested by centrifugation and washed twice with
phosphate buffered solution (PBS). The cells were then lysed
by incubating in lysis buffer [20 mmol/L EDTA, 100
mmol/L Tris (pH 8.0), and 0.8% (w/v) SDS] with RNaseA/T1 (RNase
A 500 U/mL, RNase T1 20 kU/mL (Amesco) for 2 h at 37
°C. The lysed cells were then incubated in the presence of
proteinase K (20 mg/mL) for 4 h at 55 °C. DNA samples were
loaded onto 2% horizontal agarose gels containing ethidium
bromide. The gels were run at 35 V for 6 h, and the DNA
fragments were visualized using UV
illumination[16].
PI apoptosis assay For the flow cytometric analysis of
fragmented DNA, 5×105 cells/well on the 6-well plates were
harvested after incubation in the presence or absence of
LPS for 12 h, incubated with 1 mL of 75% cold ethanol for 2 h
at -20 °C, and then washed with PBS. The cell pellets were
incubated with 50 mg/L PI and 100 mg/L RNase A at 37 °C for
30 min. Sub G1 cell counts were determined on a FACSort
flow cytometer using the Cell Quest analysis
program[7].
Caspase-3 activity assay Caspase-3 activity was
evaluated using the CaspACE assay system according to the
protocol provided by the manufacturer. The cells plated in the
flasks were treated with LPS and/or CHX for 8 h, were then
harvested, washed, and lysed by 4 cycles of freezing and
thawing, followed by incubation on ice for 15 min. The
samples were then centrifuged at 15
000×g for 20 min at 4 °C. The cell lysate supernatant was aliquoted into the wells of a
96-well plate, and 32 µL buffer, 2 µL DMSO, 10 µL DTT (100
mmol/L), and 2 µL of DEVD-pNA (10 mmol/L) were added to
a final volume of 100 µL. The samples were incubated for 4 h
at 37 °C. The absorbance at 405 nm was measured and
enzyme activity was calculated as pmol pNA liberated per
hour/µg protein according to the formula provided by the
manufacturer.
Statistical analysis All experiments were repeated at least
3 times. All data were expressed as mean±SD. Statistical
analysis was performed by the Student's t-test using the
software SPSS 11.0 for Windows (SPSS, Chicago, USA).
P<0.05 was considered statistically significant.
Results
Effect of 3,5-diCQA on microsome lipid peroxidation
The anti-oxidative activity of 3,5-diCQA was assessed by
measuring its effect on microsome lipid
peroxidation. The exposure of microsomes to
ascorbate-Fe2+ led to an increase in the level of MDA from the control level of 0.49 µmol/g
protein to 0.93 µmol/g protein (P<0.01) indicating
lipoper-oxidation. A concentration-dependent decrease in
ascorbate-Fe2+-induced MDA formation to 0.72, 0.64, and 0.52
µmol/g protein was observed for 3,5-diCQA at 5, 10, and 50
µmol/L, respectively (Figure 2). Treatment with 20 µmol/L
α-tocopherol, a well known anti-oxidant, gave an MDA value
of 0.73 µmol/g protein (P<0.01).
Inhibitory effect of 3,5-diCQA on ROS formation
According to the result in the model of the microsome above,
we further determined intracellular ROS levels to assess the
anti-oxidative activity of 3,5-diCQA in LPS-stimulated
HMEC-1. Cells treated with CHX and LPS for 12 h gave an
increase in intensity of fluorescence indicative of ROS
formation, 1.8-fold the intensity observed for the control cells
treated with CHX alone. Pretreatment with 3,5-diCQA (5, 10,
or 50 µmol/L) or α-tocopherol (20 µmol/L) significantly
decreased the intensity of fluorescence (Figure 3).
Effect of 3,5-diCQA on cell viability and damage
figure Consistent with previously reported data, and supported by
the predictive assay presented in the present report, LPS did
not induce injury in HMEC-1 alone. Injury was induced when
HMEC-1 were treated with LPS together with CHX. The
effect of 3,5-diCQA on LPS-induced apoptosis in HMEC-1 in
the presence of CHX was examined. After 16 h incubation
with both LPS and CHX, cell viability was reduced to 60% of
the level of the control group treated with CHX alone.
Pretreatment with 3,5-diCQA (5, 10, or 50 µmol/L) decreased the
cell death rate to 19%, 14%, and 12%, respec-tively (Table 1).
The release of LDH after LPS treatment for 16 h was
significantly higher than the cells treated with CHX alone.
Treatment with 3,5-diCQA at 5, 10, or 50 µmol/L decreased LDH
release in a concentration-dependent manner. α-Tocopherol
(20 µmol/L) also showed significant suppression of
LPS-induced cell death and LPS-induced LDH release (Table 1).
Effect of 3,5-diCQA on cell apoptosis Apoptosis induced
by LPS in the presence of CHX was confirmed by the DNA
fragmentation assay. CHX alone did not induce
fragmentation of chromosomal DNA. Treatment with 3,5-diCQA
or α-tocopherol suppressed LPS-induced DNA
laddering (Figure 4). Flow cytometric analysis was
performed after PI staining. DNA fragmentation was observed in
27% of cells with hypodiploid DNA in LPS-induced apoptosis. The percentage of
cells with DNA fragmentation decreased significantly on
treatment with 10 or 50 µmol/L 3,5-diCQA, while 5 µmol/L
3,5-diCQA and 20 µmol/L α-tocopherol had no effect (Figure 5).
Effect of 3,5-diCQA on caspase-3
activity Caspase-3 activity in HMEC-1 was 8.92 pmol pNA liberated/hour in
LPS-induced apoptosis. In the cells treated with CHX alone, the
caspase-3 activity was similar to the control group (data not
shown). Caspase-3 activity decreased in a dose-dependent
manner on treatment with 3,5-diCQA. Caspase-3 activity
induced by LPS was suppressed by 20 µmol/L α-tocopherol
(Figure 6).
Discussion
In the present study, 3,5-diCQA effectively suppressed
the generation of oxide free radicals in the microsome
model, reduced the production of intracellular ROS induced by
LPS in HMEC-1, protected HMEC-1 from LPS-induced cell injury,
and inhibited the activity of caspase-3.
LPS is a bacterial endotoxin that functions as a pro-
inflammatory mediator inducing significant levels of
endogenous TNF-α and mediating endothelial cell
damage[11]. LPS induces apoptosis in bovine and ovine endothelial cells
in vitro and elicits apoptosis in human endothelial cells in the
absence of new gene expression. The induction of apoptotic
cell death by LPS is confirmed by several criteria, including
morphological changes, DNA laddering, TdT-mediated dUTP
nick end labeling, the activation of caspase, and poly
(ADP-ribose) polymerase cleavage[1].
In the present study, LPS-induced cytotoxicity of HMEC-1
was suppressed by treatment of the cells with 3,5-diCQA or
α-tocopherol. Cell injury, determined by the release of LDH,
was also suppressed by treatment with 3,5-diCQA. The DNA
fragmentation associated with LPS-induced cell death was
partially inhibited by treatment with 3,5-diCQA.
Oxidants or pro-oxidants are significant regulators of
apoptosis and can induce the apoptotic
process[24,25]. The significance of oxidative stress in apoptosis is strongly
supported by the capacity of various cellular anti-oxidants to
block apoptosis induced by a diverse number of
agents[25,26]. Oxidative stress is a common factor in apoptosis and is
induced by stimuli including LPS and TNF-α
exposure[3,19,27_29]. Several phenylethanoids also possess free radical
scavenging properties and protect cells against oxidative stress-
induced injury[30_32]. In the present study, we investigated
the anti-oxidant capacity of the phenylethanoid 3,5-diCQA
and found that both lipid peroxidation of microsomes and
intracellular ROS accumulation induced by LPS were
significantly suppressed. These results suggest that 3,5-diCQA
may protect endothelial cells against apoptosis by directly
scavenging intracellular ROS.
The signaling pathways induced by apoptosis converge
to the caspase pathway to execute the final phase of the
apoptotic process[12]. The caspase family of proteases
consists of at least 14 members in mammals that are
constitutively expressed in almost all cell types as inactive pro-
enzymes (zymogens) that become processed and activated
in response to a variety of pro-apoptotic
stimuli[13]. Caspase-3 is a downstream member of the caspase cascade and acts
as a central effector in the execution phase, the proteolytic
events of which can lead to apoptosis and contribute to
DNA fragmentation, nuclear morphological changes, and
eventual cell death. The effect of LPS-induced caspase
activation on endothelial cells is dramatic, resulting in the
cleavage of nuclear proteins and structural proteins that mediate
cell-cell and cell-substrate
adhesion[18,21]. In vivo studies
support a role for caspases in mediating LPS-induced EC
apoptosis[20, 22]. Thus, substances that inhibit the activity of
caspase-3 may protect cells from apoptosis. The
anti-apoptotic capacity of 3,5-diCQA was identified in the present
study by the marked suppression of caspase-3 activity in
LPS-treated cells.
In summary, 3,5-diCQA displays anti-oxidative and
anti-apoptotic activity in HMEC-1 due to the scavenging of
intracellular ROS and the reduction of caspase-3
activity. The anti-oxidative and anti-apoptotic capacity of
3,5-diCQA provides clinical potential for the treatment of diseases
involving the progressive dysfunction of the endothelium,
including ischemia, shock, or sepsis.
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