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1
Departments of
* Oncology, Transplants and Advanced Technologies in Medicine,
Physiology and Biochemistry,
Psychiatry and Neurobiology, University of Pisa, School of Medicine; and
Institute for Neurophysiology, CNR, Pisa, Italy
1Correspondence: Department of Oncology, Transplants, and Advanced Technologies in Medicine, University of Pisa, Via Paradisa 2, I-56124 Pisa, Italy. E-mail: biancam{at}dfb.unipi.it
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: confocal microscopy • reactive oxygen species • antioxidants • dihydrorhodamine 123 • melatonin • transplantation
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. However, severe side effects
have been found to be associated with life-long treatment of
transplanted patients with CSA, including renal and vascular toxicity
(2
3
4)
.
The observation that CSA causes oxidative stress in rat hepatocytes
(5)
and that antioxidants such as vitamin E, melatonin,
and other indolic compounds inhibit lipid peroxidation in animal models
of cyclosporin A-induced nephrotoxicity (6
7
8
9)
suggests a
role for oxidative stress in CSA-induced damage. As the endothelium is
known to influence vascular tone (10
, 11)
and is
considered to be the main site of impaired vascular regulation by CSA
(12
13
14
15)
, endothelial cell (EC) damage by oxidative stress
may play a key role in the vascular problems of transplanted patients
treated with CSA. Accordingly, we have recently shown that CSA induces
a concentration-dependent increase in oxidants and lipid peroxidation
in cultured human endothelial cells (16)
. Considering the
well-known relationship between oxidative stress and apoptosis
(17
, 18)
, CSA-induced oxidative stress may damage EC via
apoptotic cell death.
However, oxidative stress-induced apoptosis occurs for acute exposure
to CSA concentrations outside its therapeutic range (19
, 20)
, and it is presently unclear whether it may play a causal
role in the vascular problems of patients chronically treated with
therapeutic doses of CSA.
The main aim of the present work was to provide a better understanding of the pathogenesis of CSA-induced vascular toxicity by comparing its acute (high levels for short time) vs. chronic (low levels for long time) effects on oxidative stress and apoptosis in human EC.
Our study suggests that acute treatment with CSA is associated with apoptosis and with a decrease of Bcl-2 protein expression in EC. On the other hand, chronic treatment with therapeutic levels of CSA affects EC via mechanisms that involve adaptation to CSA-induced oxidative stress through Bcl-2 up-regulation, which may have a key role in promoting the survival of EC with an impaired function, as suggested by their altered morphology.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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with minor modifications. After collection, umbilical
cords were placed in cold saline solution containing penicillin 25 U/ml
and streptomycin 50 µg/ml. The umbilical veins were cannulated and
perfused with saline solution to wash out the residual blood. Both ends
of the cord were clamped and then infused with saline containing 0.25%
type IA collagenase. After 10 min incubation at 37°C, the collagenase
solution containing detached HUVECs was flushed out of the cord with 10
ml of Medium 199. The cells were then centrifuged at 100 g
and resuspended in complete Medium 199 containing 20 mM HEPES buffer,
20% heat-inactivated fetal calf serum, 0.2% growth factor for
endothelial cells, 2 mM glutamine, 5 U/ml heparin, 25 U/ml penicillin,
and 50 µg/ml streptomycin. HUVECs were cultured overnight in 25
cm2 gelatin-coated tissue culture flasks
(Corning, Corning, N.Y.) at 37°C in a NAPCO incubator (5%
CO2, 95% air). The next day contaminating cells
were rinsed out of the flask and 9 ml of complete medium was added. At
confluence, HUVECs were trypsinized and plated into three flasks
for an additional passage. HUVECs were used for experiments in the
third subpassage. HUVECs were identified by their typical cobblestone
appearance and positive indirect immunofluorescence for factor VIII.
All reagents used for cell dissociation and culture were obtained from
Sigma (Milan, Italy).
Phase-contrast microscopy
For phase-contrast microscopy, cells were cultured on 75
cm2 culture flasks (Greiner) and exposed to
experimental conditions. Phase-contrast microscopic examination was
performed using an inverted Zeiss Televal 3 (light source 25 W halogen
lamp) equipped with a Nikon camera.
Cell density was evaluated by the trypan blue exclusion procedure. The cell suspension was diluted 1:2 with the dye (trypan blue 0.4%, Sigma) prior to counting of viable cells with a hemocytometer.
Monitoring of reactive oxygen species production
For microscopic observation, HUVECs were plated on
gelatin-coated glass coverslips until they reached subconfluence. The
glass coverslip was adapted to a Plexiglas chamber and HUVECs were
loaded with 10 µM dihydrorhodamine 123 (DHR123) (Molecular Probes,
Eugene, Oreg.) for 30 min at 37°C in culture medium. The chamber was
positioned for imaging on the stage of an inverted microscope (Nikon
Eclipse TE300) equipped with a laser confocal scanning system (Radiance
Plus, Bio-Rad, Hercules, Calif.). The stock solution (10 mM) of
dihydrorhodamine 123 in DMSO contained less than 1% of contaminating
rhodamine 123 (RHD123), as assessed by spectrofluorometry using a
Perkin-Elmer LB50 Fluorescence Spectrometer. Contamination of rhodamine
123 present in traces in the dihydrorhodamine stock was responsible for
the low background fluorescence observed at the beginning of the
recording session, which did not change in untreated cells over a
period of 2 h. Cyclosporin A was added after 15 min of observation
(baseline); 2 µM SNARF (Molecular Probes) was also included in some
experiments to assess cell morphology and viability. Acridine orange
(Molecular Probes) was used at 10 µg/ml as a membrane permeant and an
intercalating agent to label nucleic acids.
Fluorescence images were collected every 3 min using the 488 nm
excitation wavelength from an argon laser and the 515 nm emission
filter. To minimize photo-oxidation of the probe, the laser beam was
attenuated to 50% of maximal illumination and exposure of cells to
light was limited to the image acquisition intervals (
2 s every 3
min) via the acquisition software. Analysis of the fluorescent signal
was performed with Adobe Photoshop 5.0 (Adobe, San Jose, Calif.) for
each cell and for each period of time over a similar region of
interest, which included most of the cells. To compensate for the
pronounced variability of fluorescence from cell to cell (see Results,
Fig. 1A
, and Fig. 6A
) that was already present in
controls, for each cell the effect of CSA on fluorescence was expressed
as the relative increase over the basal value at the beginning of
recording. In control experiments, without dihydrorhodamine 123, CSA
did not affect the low autofluorescence of HUVECs.
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Lactic dehydrogenase (LDH) assay
LDH release was measured as the enzymatic activity present in
the cell medium. Media were collected from cell cultures at appropriate
times and centrifuged to remove contaminating cells and debris. LDH
activity was assessed by standard enzymatic methods (22)
.
Lipid peroxidation assay
Lipid peroxidation was evaluated by using a commercial
colorimetric kit by Calbiochem (San Diego, Calif.) for malonaldehyde
and 4-hydroxyalkenals (MDA+4-HNA). MDA + 4-HNA production was
determined at different time points in control and CSA-treated cell
cultures in accordance with the manufacturer’s instructions.
DNA extraction and gel electrophoresis
Endothelial cells (107 cells/sample) were
washed with NaCl 0.9%, trypsinized, and harvested by centrifugation
(100 g for 10 min). The pellet was washed with NaCl 0.9%
and resuspended in 500 µl of digestion buffer (100 mM NaCl, 10 mM
Tris-HCl pH 8, 25 mM Na2EDTA pH 8, 0.5% SDS, 0.1
mg/ml proteinase K, and 1% 2-mercaptoethanol). After overnight
incubation at 50°C, the samples were extracted twice with
phenol/chloroform/isoamyl alcohol (25/24/1) and centrifuged at 10,000
g for 10 min. DNA in the aqueous phase was precipitated by
adding to the sample 0.5 volume of 7.5 M ammonium acetate and 2 volumes
of ethanol, and recovered by centrifuging at 10,000 g for 2
min. The pellet was rinsed with 70% ethanol and air dried, and DNA was
resuspended in TE buffer (10 mM Tris-HCl pH 8, 1 mM
Na2EDTA pH 8). Residual RNAs were removed with 1
µg/ml DNase-free RNase-A by incubating for 1 h at 37°C. DNA
was resolved in 1% agarose gel in the presence of 2 µg/ml ethidium
bromide and visualized with UV transillumination. Photographs were
directly taken with a Polaroid M4 camera.
Bcl-2 and nucleosome ELISA
Human Bcl-2 protein and nucleosomes were quantified in HUVEC
extracts by using the Bcl-2 and the nucleosome ELISA kits from Oncogene
Research Products (Cambridge, Mass.) in accordance with the
manufacturer’s instructions. Readings of colored products were taken
by using a spectrophotometer plate reader (Titertek Multiskan MCC) and
compared with the standard curves. Values were expressed as units of
Bcl-2 or nucleosomes/mg of protein.
Protein measurement
Protein concentration was determined in accordance with the
method of Lowry et al. (23)
.
CSA application
CSA was obtained either as a powder from Alexis Corporation
(Läufelfingen, Switzerland) or as a solution in castor oil
(Novartis, Pharma, Basel, Switzerland). CSA powder was dissolved in
ethanol at 10 mM and brought to final concentrations in Medium 199. The
CSA formulation in castor oil was dissolved in Medium 199. Similar
effects were obtained with both formulations of CSA; control
experiments with ethanol and castor oil ruled out a possible effect of
the vehicle. As CSA binds to plasma proteins in vivo, the
free CSA concentration ([CSA]) in serum-containing Medium 199 is
expected to be lower than the analytical concentration
(Ctot). Determination of [CSA] in Medium 199
was performed with an FPIA kit (Fluorescence Polarisation Immunoassay)
from Abbott Laboratories (Abbott Park, Ill.). The measurement of
[CSA] over a range of Ctot values indicates
that [CSA]/Ctot 
0.5; in other words, the
effective CSA concentration is approximately half
Ctot. Concentrations of CSA are reported here as
free concentrations ([CSA]).
Statistical analysis
Measurements are reported as mean ± standard error of the
mean (SE). Analysis of HUVEC fluorescence shows the
presence of cells with large differences in intensity, even in basal
conditions. Accordingly, plots of average cellular fluorescence result
in a bimodal distribution (data not shown), indicating that HUVECs may
not be considered as a homogeneous population normally distributed.
Consequently, the statistical significance of the differences in
fluorescence observed before and after CSA application was assessed by
either nonparametric tests (Mann-Whitney U test) or by 2-way
analysis of variance (2-way ANOVA), using tabulated values of
probabilities from the F distribution (24)
.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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13 min after application of CSA, when
a steady state has been reached, and may be compared to control
conditions (Fig. 1A
).
2-way ANOVA for the effect of the duration of CSA application vs.
control was carried out in 75 cells from three independent experiments
and indicated that the increase in fluorescence induced by CSA was
statistically significant after both 10 and 15 min of exposure to the
drug (P<0.005 and P<0.001, respectively). Note
that despite clear differences in fluorescence intensity between cells
already present in control conditions, the average relative increase in
fluorescence over basal values (see methods) induced by CSA was similar
for all cells, averaging 105.4 ± 11.9%
(mean±SE) after 13 min of exposure to the drug.
The increase in cellular oxidants monitored by the oxidation-sensitive
probe is associated with an increased release of malonaldehyde and
4-hydroxyalkenals, by-products of lipid peroxidation (25)
,
in the culture medium from 9.6 ± 0.7 pmol
ml-1 in control conditions to 109.4 ± 21.2
pmol ml-1 after 21 h in CSA 200 µM
(P<0.05, n=3, paired t test).
To correlate CSA-induced oxidative stress with apoptosis, HUVECs were
double stained with DHR 123 and the cytoplasmic indicator SNARF. As
illustrated in Fig. 2
, when 200 µM CSA treatment for 20 min had already caused an increase
in rhodamine 123 fluorescence (green), there was the appearance of
multiple cytoplasmic (white arrows) membrane blebs, an early sign of
apoptosis (Fig. 2A
).
|
Additional indications for the occurrence of apoptosis in response to
200 µM CSA were assessed by staining the cells with acridine orange,
a membrane-permeant and nucleic acid-intercalating probe. After 50 min
from the application of 200 µM CSA, clear changes in nuclear
morphology, including chromatin condensation and lobule formation, were
present (Fig. 2B
). Note that these early signs of apoptosis
were not present simultaneously in every cell. CSA did not cause an
acute release of lactic dehydrogenase, suggesting that pharmacological
concentrations of CSA trigger an apoptotic rather than a necrotic
process in HUVECs.
Molecular basis of CSA-induced damage
To assess the ability of HUVECs to complete the apoptotic process
triggered by 200 µM CSA, we performed DNA fragmentation analysis. As
illustrated in Fig. 3A
, the DNA from control cells appears to be intact (lane 2)
whereas treatment with CSA 200 µM (lane 3) for 21 h induces DNA
degradation, with the ladder-like appearance of the classic
nucleosomal-sized DNA fragmentation characteristic of apoptosis
(26
27
28)
. Note that simultaneous treatment with 300 µM
melatonin, a hormone with antioxidant properties (29
, 30)
,
fails to protect HUVECs from DNA degradation (lane 4).
|
To verify the role of oxidants in the proapoptotic effects of CSA, we
performed quantitative estimates of DNA fragmentation by measuring the
levels of nucleosomes with ELISA. Column bars of Fig. 3B
indicate that in HUVECs acutely treated with CSA (200 µM for 21 h), nucleosome levels increased from a control value of 0.6 ± 0.1
U/mg protein to 29.3 ± 2.3 U/mg protein. However, nucleosome
levels in HUVECs treated simultaneously with 200 µM CSA and either
300 µM melatonin or 100 µM Trolox (a water-soluble analog of
vitamin E) were 27.8 ± 1.0 or 27.3 ± 2.0 U/mg protein,
significantly higher than control levels (P<0.001 for both
antioxidants) and not significantly different from CSA alone (1-way
ANOVA). The increase in oligonucleosome formation was not significant
2 h after CSA application (data not shown), suggesting that an
induction time is required for the activation of enzymes involved in
DNA fragmentation.
To further characterize the molecular basis of the apoptotic process in
consideration of the importance of antiapoptotic proteins, like Bcl-2,
we measured the effects of acute CSA treatment on expression of the
antiapoptotic protein Bcl-2 in HUVECs. Figure 3C
shows that
treatment with 200 µm CSA for 21 h, which induces ladder-like
DNA fragmentation, also induces a decrease in Bcl-2 protein expression
from 4.6 ± 0.1 U/mg protein (control) to 1.4 ± 0.1 U/mg
protein (treated cells); melatonin did not prevent Bcl-2 from falling
to 1.8 ± 0.3 U/mg of protein.
These results suggest that pharmacological levels of CSA affect HUVEC survival by altering the balance between pro- and antiapoptotic factors although by mechanisms independent of the drug’s ability to induce oxidative stress.
Effects of acute CSA treatment on endothelial cell density and
morphology
This idea was further tested by observing the effects of CSA on
HUVEC vitality. Treatment of HUVECs with 200 µM CSA for 21 h
causes a decrease in cell density (Fig. 4B
) compared with control cultures (Fig. 4A
).
|
Moreover, the acute treatment also affects cellular morphology, with
the appearance of round cells (arrows in Fig. 4B
) quite
different from the typical polygonal shape of HUVECs (Fig. 4A
). Simultaneous treatment of cells with CSA and
antioxidants such as Trolox (Fig. 4C
) or melatonin (Fig. 4D
) failed to prevent the effect of acute CSA treatment on
HUVEC density. A similar lack of protection has also been noted with
other antioxidants, such as ascorbic acid or N-acetyl-serotonin (data
not shown). The failure of antioxidants to afford protection against
the effects of acute treatment indicates that CSA triggers apoptotic
cell death of HUVECs via an oxidative stress-independent mechanism.
Effects of chronic CSA treatment on endothelial cell density and
morphology
As illustrated in Fig. 5
, chronic treatment of HUVECs with CSA (2 µM for 8 days) did not
appreciably affect cell density (Fig. 5B
) compared with
control cultures (Fig. 5A
).
|
However, HUVECs treated with CSA have an altered morphology (Fig. 5B
), with a transition to an elongated shape, when compared
with controls (Fig. 5A
). These morphological changes induced
by chronic treatment with low CSA concentrations were suppressed by
simultaneous treatment with Trolox (Fig. 5C
) or vitamin C
(Fig. 5D
). A cobblestone morphology was also preserved with
melatonin (data not shown). This indicates that chronic treatment with
CSA may cause oxidative stress, which in turn affects the expression of
specific proteins involved in maintaining the endothelial phenotype.
Oxidative stress on exposure to therapeutic levels of CSA
In agreement with this notion, confocal microscopy of EC exposed
to CSA concentrations of clinical importance revealed an increased
production of oxidants, albeit to a lower extent than in response to
acute treatment. Figure 6
shows the increase in RHD 123 fluorescence in cells treated with 2.5
µM CSA for 15 min (Fig. 6B
) compared with control
conditions (Fig. 6A
) before CSA; 2-way ANOVA indicates that
the increase in fluorescence is statistically different from controls
10 and 15 min after CSA application.
The average relative increase was 37.0 ± 4.1%, calculated over
11 cells. Similar relative increase were measured in 70 cells from
three independent experiments. In addition to the increased generation
of oxidants monitored by DHR 123, chronic treatment of cells with 2.5
µM CSA significantly increased the release of malonaldehyde and
4-hydroxyalkenals, as shown in Table 1
.
|
Despite the generation of oxidants in response to therapeutic levels of
CSA (2.5 µM), no morphological signs of apoptosis were apparent (data
not shown). Moreover, EC chronically treated (8 days) with 0.5–2.5
µM CSA did not undergo nucleosomal-size DNA fragmentation, as shown
in Fig. 7A
. On the other hand, HUVECs surviving the chronic challenge
with clinical levels of CSA show an increased expression of the
antiapoptotic protein Bcl-2. Figure 7B
shows that Bcl-2
protein expression significantly increased in cells exposed to chronic
CSA treatment (0.5–2.5 µM for 8 days).
|
Simultaneous treatment of EC with the antioxidant melatonin (300 µM),
which was ineffective in preventing the DNA degradation induced by 200
µM CSA (see Fig. 4
), suppresses the increase of Bcl-2 protein
expression in response to 0.5–2.5 µM CSA, suggesting that low levels
of oxidants may act as signaling molecules that trigger the adaptation
process (Fig. 7B
).
Chronic exposure to clinical concentrations of CSA induces
adaptation and protects HUVECs against acute challenge with
pharmacological CSA concentrations
To verify that clinical concentrations of CSA trigger an
adaptation process, HUVECs were challenged with 200 µM CSA after 6
day exposure to 2.5 µM CSA. Data in Fig. 8
show that HUVEC cultures acutely challenged for 21 h with 200 µM
CSA have a significantly higher survival rate when chronically treated
for 6 days with 2.5 µM CSA (Fig. 8C
) than cultures not
adapted to CSA prior to the acute challenge with the drug (Fig. 8B
).
|
Note that the cell density in Fig. 8C
was significantly
lower than in cultures treated for 6 days with control medium (Fig. 8A
) or 2 µM CSA (not shown). The cell density mirrored the
changes in the level of the Bcl-2 protein. This suggests that the Bcl-2
up-regulation induced by oxidants during the chronic treatment with
therapeutic doses of CSA allows the cells to survive the acute
challenge with pharmacological doses of CSA.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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We will next discuss the role that the properties of either CSA or HUVECs may play in eliciting the spectrum of responses to the drug. We will also discuss the possible clinical significance of adaptation to clinical concentrations of CSA through Bcl-2 up-regulation.
Pharmacological doses of CSA triggers oxidative stress-independent
apoptosis in HUVECs
Pharmacological concentrations of CSA have been observed to cause
oxidative stress in renal (31)
and hepatic
(5)
tissues and in glioma cells (19)
.
As oxidative stress is a well-known cause of apoptosis (17
, 18)
, we investigated the ability of pharmacological doses of CSA
to induce apoptosis in HUVECs. As shown in Figs. 2
and 3
, acute
exposure to nonclinical levels of CSA concentrations induces both the
morphological and biochemical hallmarks of apoptosis. Thus, CSA-induced
apoptosis probably plays a role in the decrease in HUVEC density
documented in Fig. 4B
. However, the observation that
simultaneous treatment with antioxidants does not suppress the
biochemical markers of apoptosis (Fig. 3A
) or the loss of
HUVECs (Fig. 4C
, D
), while blocking CSA-induced lipid
peroxidation (16)
, suggests that pharmacological
concentrations of CSA induce apoptosis by mechanisms that operate
independently of the simultaneously occurring oxidative stress.
Both CSA and its complexes with cyclophylins (32
, 33)
are
known to exert several biological effects in target cells (for a
review, see ref 34
), such as modulation of the protein
phosphatase calcineurin (35)
or of the permeability
transition pore at the mitochondrial level (36)
.
Considering that CSA has previously been reported to protect cells from
apoptosis because of its ability to block the mitochondrial
permeability transition pore (37)
, it seems unlikely that
this mitochondrial action is responsible for CSA-induced apoptosis in
HUVECs. Rather, the observation that apoptosis is independent of
oxidative stress, despite the occurrence of lipid peroxidation in
response to acute treatment with pharmacological concentrations of CSA
(16)
, suggests that perhaps CSA protects HUVECs from
oxidative stress-induced apoptosis.
On the other hand, inhibition of calcineurin (38)
, a
calcium-activated protein phosphatase that plays a key role in the
immunosuppressive action of CSA, may have a causal role in CSA-induced
apoptosis in HUVECs. Despite the lack of any direct evidence for a
specific role of calcineurin in inducing oxidative stress in HUVECs, it
seems reasonable to postulate that, in general, the interaction of the
CSA/cyclophilins complexes with specific proteins may trigger an
apoptotic program independently of oxidative stress. Further studies
are required to define the molecular basis underlying the ability of
CSA to induce both the increase in cellular oxidants and apoptosis and
to elucidate the possible relationships between these two pathways of
CSA action in HUVECs.
Clinical concentrations of CSA trigger oxidative stress-dependent
adaptation in HUVECs
The observation that CSA-induced oxidants do not play a role in
triggering apoptosis does not mean they are devoid of any effects in
HUVECs. Indeed, antioxidants suppress the effects of clinical
concentrations of CSA, including the up-regulation of Bcl-2 (Fig. 7B
) and the modification of cell morphology from a
cobblestone to an elongated shape (Fig. 5B
). This suggests
that clinical levels of CSA affect HUVECs through mechanisms regulated
by oxidants and may also indicate that cellular oxidants are the second
messengers mediating the effects of clinical levels of CSA in HUVECs.
It is important to note that the increase in oxidants takes place on a time scale of minutes, attaining a steady level as long as CSA is present in the medium, whereas the increase in Bcl-2 levels and the changes in cell morphology occur on a slow time scale, because they are present after 4 days but not after 24 h. This suggests that changes in Bcl-2 levels and cell shape represent a true adaptive response to a chronic increase in oxidants rather than a short-term response to a low level of oxidants.
The notion of an adaptive response to oxidants via an increase in Bcl-2
is further supported by the data in Fig. 8
, which show an improved cell
survival to acute challenge with 200 µM CSA after 6 days pretreatment
with 2.5 µM CSA that increases Bcl-2 levels.
Relationship between the level of CSA-induced oxidants,
apoptosis and adaptation
Previous work with mammalian fibroblasts (see ref 39
for a recent review) indicates that the switch-over between adaptation
and apoptosis may occur for increase of external
H2O2 between three- and
sixfold. Moreover, recent data obtained in bovine EC exposed to carbon
monoxide (CO) indicate that either adaptation or apoptosis develops
within a less than 10-fold increase in environmental CO levels
(40)
. These data suggest that EC share with other
mammalian cells the ability to trigger either an apoptotic or an
adaptive response, the switch operating over a narrow range of
concentrations of either oxidants or oxidant generators.
According to measurements based on laser scanning confocal microscopy,
CSA induces an increase in the fluorescent signal of 30% and 100%
over basal values in HUVECs exposed to CSA concentrations of 2.5 and
200 µM, respectively. Assuming a linear relationship between oxidant
levels and cellular fluorescence, a twofold increase in cellular
oxidants may be lower than the threshold for triggering an apoptotic
response. On the other hand, this increase is enough to cause lipid
peroxidation (16)
, which paves the way to apoptosis, but
CSA-induced inhibition of the mitochondrial permeability transition
pore may block the apoptotic response to CSA-induced oxidative stress.
Furthermore, the possibility that the failure of oxidative stress to
trigger apoptosis in HUVECs is related to differences between the
factors regulated in CO-treated EC (i.e., superoxide dismutase and
caspase-1) and in CSA-treated EC (Bcl-2) or to interspecies differences
(human vs. bovine) may not be ruled out at this time. It must also be
considered that CSA probably modulates the level of several pro- and
antiapoptotic factors in EC, perhaps including caspases in addition to
Bcl-2, and further studies will be required to address this issue. At
clinical concentrations of CSA, a 30% increase over basal values of
cellular oxidants may be enough to modulate Bcl-2 levels in a way
similar to that reported for other mammalian fibroblasts and
endothelial cells.
In conclusion, whereas the responses of HUVECs to acute treatment with pharmacological concentrations might mirror the presence of different branches in the CSA mechanisms of action, the ability to switch between adaptation and apoptosis for different levels of CSA may rely as much on the different effects of CSA on its target proteins as on the general properties of the response of HUVECs to cellular oxidants, which may operate independently to produce the full spectrum of drug effects in HUVECs.
Adaptation to oxidants generated in EC by clinical doses of
CSA: friend or foe?
The observation that clinical doses of CSA may affect HUVECs
through the generation of oxidants without causing apoptosis does not
support a role for CSA-triggered apoptosis in the pathogenesis of
vascular problems of transplanted patients. On the other hand, the
up-regulation of Bcl-2, which occurs at clinical concentrations and
indicates an adaptation to oxidants, may play a key pathogenetic role
in these patients. The morphological changes observed in response to
clinical levels of CSA with the loss of the cobblestone morphology
typical of endothelial cells and the development of an elongated
‘fibroblast-like’ phenotype may indicate a loss of specialization
that occurs in parallel to adaptation of HUVECs to CSA-induced
oxidants.
By analogy with the increased resistance of cancer cells to
antineoplastic drugs because of their increased content of Bcl-2
(41)
, the up-regulation by CSA-induced oxidants of Bcl-2
may increase the survival of functionally impaired HUVECs, as suggested
by their altered morphology. Future studies are required to define the
functional properties of HUVECs chronically exposed to clinical levels
of CSA, with special reference to the regulation of
·NO production, which may eventually affect the
function of the vasculature.
Considering that endothelial cell lysis is an important step in the
cascade of events responsible for hyperacute rejection of
xenotransplanted organs (42)
, the adaptive properties of
endothelial cells may be useful to maintain the integrity of their
membrane and its function as a selective barrier during hyperacute
rejection. Recent results indicate that Bcl-2 transfection increases
the ability of HUVECs to colonize and form new microvessels when
xenotransplanted in immunodeficient mice (43)
. A potential
hazard of including HUVECs transfected by retroviral transduction into
synthetic tissues intended for human use is the malignant potential of
Bcl-2 overexpressing cells. However, we found no evidence for increased
cell proliferation in HUVECs with up-regulated Bcl-2 levels.
Furthermore, life-long treatment with CSA does not induce endothelial
cell proliferation in transplanted patients, suggesting that Bcl-2
overexpression in human endothelial cells may not pose major concerns
in clinical application. Our results may thus encourage further
development of Bcl-2 overexpressing cells in the engineering of tissue
equivalents.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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 S.-Y. Sun, N. Hail Jr, and R. Lotan Apoptosis as a Novel Target for Cancer Chemoprevention J Natl Cancer Inst, May 5, 2004; 96(9): 662 - 672. [Abstract] [Full Text] [PDF]
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 T. Chen, J. Guo, M. Yang, C. Han, M. Zhang, W. Chen, Q. Liu, J. Wang, and X. Cao Cyclosporin A impairs dendritic cell migration by regulating chemokine receptor expression and inhibiting cyclooxygenase-2 expression Blood, January 15, 2004; 103(2): 413 - 421. [Abstract] [Full Text] [PDF]
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 S. Carnevali, S. Petruzzelli, B. Longoni, R. Vanacore, R. Barale, M. Cipollini, F. Scatena, P. Paggiaro, A. Celi, and C. Giuntini Cigarette smoke extract induces oxidative stress and apoptosis in human lung fibroblasts Am J Physiol Lung Cell Mol Physiol, June 1, 2003; 284(6): L955 - L963. [Abstract] [Full Text] [PDF]
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 B. C. Santos, J. M. Pullman, A. Chevaile, W. J. Welch, and S. R. Gullans Chronic hyperosmolarity mediates constitutive expression of molecular chaperones and resistance to injury Am J Physiol Renal Physiol, March 1, 2003; 284(3): F564 - F574. [Abstract] [Full Text] [PDF]
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 G. Schneider, F. Oswald, C. Wahl, F. R. Greten, G. Adler, and R. M. Schmid Cyclosporine Inhibits Growth through the Activating Transcription Factor/cAMP-responsive Element-binding Protein Binding Site in the Cyclin D1 Promoter J. Biol. Chem., November 8, 2002; 277(46): 43599 - 43607. [Abstract] [Full Text] [PDF]
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 M. V. Alvarez-Arroyo, S. Yague, R. M. Wenger, D. S. Pereira, S. Jimenez, F. R. Gonzalez-Pacheco, M. A. Castilla, J. J. P. Deudero, and C. Caramelo Cyclophilin-Mediated Pathways in the Effect of Cyclosporin A on Endothelial Cells: Role of Vascular Endothelial Growth Factor Circ. Res., August 9, 2002; 91(3): 202 - 209. [Abstract] [Full Text] [PDF]
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