 |
INTRODUCTION |
Effects of nitric oxide (NO·) on cellular functions
are complex and even appear to be contradictory, janus-headed.
NO· may act cytotoxically but may also protect cells from toxic
insults (1), it may compromise the cellular redox state but may also act as an antioxidant (2, 3), and it may activate or inhibit signal
transduction pathways (4, 5) and gene transcription (6-9),
respectively. Many of these effects can, at least in part, be explained
by different NO· concentrations achieved in the respective
microenvironment. Nanomolar concentrations of NO·, as typically
synthesized in a tightly regulated fashion by constitutively expressed
nitric oxide synthase
(cNOS),1 serve as a signal
molecule activating the soluble guanylate cyclase to produce cGMP,
which acts as a second messenger. However, in addition to two cNOS
there is also an inducible NOS (iNOS) expressed in a variety of acute
or chronic disease states (for reviews see Refs. 10, 11). Originally
described as a cytotoxic activated macrophage effector molecule (12) it
is now evident that iNOS-derived NO· exerts a multitude of
biological functions. In an apparently unregulated fashion iNOS
synthesizes NO· for hours or even days resulting in micromolar
concentrations of NO·. Under these conditions NO· may
react with oxygen in a reaction mainly depending on the NO·
concentration to yield higher nitrogen oxides (NOx such as
N2O3, etc.), which display a much broader
chemical reaction spectrum than NO· itself (13).
A growing body of evidence suggests that NO· after reaction to
NOx helps to orchestrate gene expression, e.g.
via posttranslational modifications of transcription factors. A
prevalent DNA binding motif of transcription factors is the zinc finger
structure with Zn2+ tetrahedrally coordinated between a
-hairpin and a short 
-helix, creating a small, functional and
independently folded domain (14). In these zinc fingers cysteine thiols
and histidine imidazole nitrogens serve as direct ligands for the zinc ion.
We previously found that NO· S-nitrosates cysteines
in metallothionein, mediating the release of Zn2+ from this
zinc-storing protein (15), induces Zn2+ release within
cells (16) and is able to inhibit zinc finger-dependent transcription (17-19). However, zinc fingers can easily be disrupted by cysteine oxidation, e.g. by reactive oxygen species, or
by electrophilic attack, e.g. by alkylating compounds (for
reviews see Refs. 20, 21). The question therefore arises, whether the
reaction of NO· with zinc fingers may have a special role,
different from the reaction with other reactive species generated
during inflammatory reactions. Such species are superoxide, its
dismutation product hydrogen peroxide, the product of its reaction with
NO·, peroxynitrite, or singlet oxygen, as well as peroxyl radicals.
To investigate the impact of these species on the zinc finger
integrity, we used the transcription factors VDR and RXR as a model
system both containing two Cys4-type zinc fingers, which bind to specific promoter sequences as the heterodimeric complex VDR/RXR. VDR/RXR-DNA complex formation was quantified after treatment of VDR/RXR with reactive oxygen and nitrogen oxide species in vitro. In addition, VDR/RXR-dependent reporter gene
activity was investigated in living cells after treatment with
NO· or reactive oxygen species. Our results demonstrate that,
among the reactive species investigated, NO· shows unique
characteristics with a potential role in gene regulation.
| |
MATERIALS AND METHODS |
Materials--
The NO-donors MAMA/NO
((Z)-1-{N-methyl-N-[6-(N-methylammoniohexyl)amino]}diazen-1-ium-1,2-diolate),
and DETA/NO
((Z)-1-[N-(2-aminoethyl)-N-(2-ammonioethyl)amino]diazen-1-ium-1,2-diolate) were synthesized as described (22). Peroxynitrite (ONOO
)
was synthesized by reaction of H2O2 with
isoamyl nitrite (Sigma) on ice at pH 12.5 (23). Contaminating
isoamyl alcohol was removed by intense washing with chloroform (23),
and unreacted H2O2 was removed by reaction with
MnO2 (24). The resulting solution was adjusted to pH 14, and the concentration of peroxynitrite was determined spectroscopically
at 302 nm (
= 1670 M1
cm1) (25). Stock solutions were stored at 
80 °C for
several months without loss of absorbance. Degraded ONOO
(ONOO
) was obtained after
neutralization of a 6-mM ONOO stock solution
with HCl and incubation for 2 h at 37 °C. The singlet oxygen
generator disodium 3,3'-(1,4-naphthylidene) dipropionate 1,4-endoperoxide (NDPO2) was synthesized as described
(26-28). Decomposed endoperoxide (NDPO2deg), verified
spectroscopically as NDP (28), was obtained by incubating a
50-mM stock solution of NDPO2 for 2 days at
37 °C. Hydrogen peroxide and tert-butyl hydroperoxide
(t-BHP) were purchased from Merck, 2,3-dimethoxy-1,4-naphthoquinone (DMNQ) was purchased from Alexis (Grünberg, Germany),
2,2-azobis-(2-amidinopropane) dihydrochloride (AAPH) was purchased from
Polysciences (Warrington, PA), and 1,25-dihydroxyvitamin
D3 (VD) was kindly provided by Dr. L. Binderup (Leo
Pharmaceutical Products, Ballerup, Denmark).
Gel Shift Assays--
Human VDR and human RXR cDNA were
subcloned into the pSG5 expression vector (Stratagene,
Heidelberg, Germany) (29). Linearized cDNA from VDR and
RXR, respectively, were transcribed with T7 RNA polymerase
and translated using wheat germ lysate as recommended by the supplier
(Promega, Mannheim, Germany). Similar amounts of in vitro
translated VDR and RXR proteins were mixed and incubated in the
presence or absence of 1 µM VD for 15 min at room
temperature in a total volume of 20 µl of binding buffer (10 mM Hepes, pH 7.9, 100 mM KCl, 0.2 µg/µl
poly[d(I-C)] and 5% glycerol). Indicated amounts of MAMA/NO,
H2O2, NDPO2, AAPH or
ONOO were added, and the mixtures were incubated for
various times. The DR3-type VD response element (VDRE) of the rat ANF
gene promoter (30) (core sequence
AGAGGTCATGAAGGACA) was labeled by a fill-in
reaction using [-32P]dCTP and the Klenow fragment of
DNA polymerase I (Promega). Approximately 1 ng of labeled probe (50,000 cpm) was added to the receptor-ligand mixture, and incubation was
continued for 30 min at room temperature. Protein-DNA complexes were
resolved on 8% nondenaturing polyacrylamide gels at room temperature
in 0.5 × TBE (45 mM Tris, 45 mM boric
acid, 1 mM EDTA, pH 8.3). The gels were dried and exposed
to a Fuji MP2040S imager screen overnight. The ratio of free probe to
protein-probe complexes was quantified with a Fuji FLA2000 reader using
Image Gauge software (Fuji, Tokyo, Japan).
Transfection and Reporter Gene Assay--
MCF-7 (human breast
adenocarcinoma) cells obtained from ATCC (Manassas, VA) were seeded
into 6-well plates (3 × 105 cells/well) and grown
overnight in RPMI 1640 supplemented with 10% charcoal-treated fetal
calf serum. Liposomes were formed by incubating 0.25 µg of the
expression vectors for VDR and for RXR each plus 1 µg of the reporter
gene constructs (four copies of the rat ANF DR3-type VDRE fused to the
tk promotor driving the luciferase reporter gene) with 15 µg of N-[1-(2,3-dioleoyloxy)propyl]-N, N,N-trimethylammonium methylsulfate (Roth,
Karlsruhe, Germany) for 15 min in a total volume of 100 µl of
H2O. After dilution with 0.9 ml of RPMI 1640, the liposomes
were added to the cells that had been prewashed with phosphate-buffered
saline. The transfection medium was removed after 5 h, and 1 ml of
RPMI 1640/10% fetal calf serum was added. After a further 8 h 0.5 ml of RPMI 1640 containing 10% fetal calf serum plus 100 nM VD or the solvent control (0.1% ethanol) together with
the indicated concentrations of DETA/NO, H2O2,
t-BHP, or DMNQ were added. The cells were lysed 8 h after the
onset of VD stimulation using a reporter gene assay lysis buffer (Roche
Molecular Biochemicals, Mannheim, Germany), and a constant-light-signal
luciferase reporter gene assay was performed using
LucLiteTMPlus (Canberra-Packard, Dreieich, Germany). The
luciferase activities were normalized to the protein content determined
by the method of Bradford (31), and induction was calculated as the
ratio of luciferase activity of VD-stimulated cells to that of solvent controls.
Semiquantitative Reverse Transcription (RT)-PCR--
Total
cellular RNA was isolated using the RNeasy Mini Kit (Qiagen,
Hilden, Germany) and the RNase-free DNase Set (Qiagen) exactly as described by the manufacturer and was used (1 µg of RNA
each) for cDNA synthesis (32). RT was carried out at 37 °C for
1 h using the Omniscript RT Kit and oligo(dT)16
primers (Qiagen). PCR was performed with this cDNA using the
Taq PCR Core Kit (Qiagen) and primers for luciferase
(Photinus pyralis) cDNA (GenBankTM
accession number X84848); sense: 5'-ATTGCTTTTACAGATGCACA-3' (bases
232-251), antisense: 5'-TAGGATCTCTGGCATGCGAG-3' (bases 771-790).
Expression of the housekeeping gene GAPDH was analyzed using primers
for GAPDH cDNA (GenBankTM accession number M17851);
sense: 5'-ATGCCCGATGGCACCATCAGA-3' (bases 153-175), antisense:
5'-TCTCCAGGCCCATCCTCCTGC-3' (bases 548-568). PCR was carried out
following standard procedures (33). To ensure that amplification
conditions were within the linear phase, PCR was performed using
various numbers of cycles with RNA isolated from transfected and
VD-stimulated cells. The following cycle profiles were found to be
suitable: 20 cycles at 94 °C/30 s, 56 °C/60 s, and 72 °C/60 s
for luciferase mRNA amplification, and 18 cycles at 94 °C/30 s,
58 °C/30 s, and 72 °C/30 s for GAPDH mRNA amplification,
respectively. As a control, PCR was performed with all additives but
without cDNA or with all additives but only with RNA, respectively,
to exclude unspecific amplifications. Equal amounts of DNA were
electrophoresed on a 1.8% agarose gel. The luciferase/GAPDH ratio was
obtained by densitometric analysis of ethidium bromide-visualized
amplification product bands using the Kodak 1D software (Kodak,
Stuttgart, Germany).
| |
RESULTS |
In VitroStudies--
To investigate effects of nitrosative and/or
oxidative stress toward zinc finger transcription factors, in
vitro translated VDR and RXR proteins were used as a model system.
VDR-RXR heterodimers (VDR/RXR) were incubated with various
concentrations of NO· or reactive oxygen species before
analyzing VDR/RXR-DNA complex formation to the DR3-type VDRE of the rat
ANF gene promoter by gel shift experiments.
Nitric Oxide--
NO· was generated by decomposition of the
NO-donor MAMA/NO (half-life at 37 °C: 2 min). Fig.
1A shows that NO·
generated by MAMA/NO inhibits VDR/RXR-DNA complex formation in a
concentration-dependent manner with an IC50
value of about 0.5 mM MAMA/NO. Using preformed VDR/RXR-VDRE
complexes as targets, MAMA/NO with a nearly identical IC50
value (about 0.4 mM) induced the release of VDR/RXR from
its response element (Fig. 1B). The DNA complex formation of
MAMA/NO-treated VDR/RXR was nearly completely restored by subsequent
incubation with 1 or 10 mM dithiothreitol (DTT) for 30 min
at 30 °C (Fig. 1C). To investigate whether NO·
affects the response element as well, VDRE was preincubated with up to
5 mM MAMA/NO for 30 min at 30 °C before VDR/RXR was
added. A significant inhibition was observed only with the highest
NO-donor concentration (Fig. 1D).
View larger version (29K):
[in this window]
[in a new window]
|
Fig. 1.
Effects of NO· on VDR/RXR-DNA complex
formation in vitro. Gel shift experiments were
performed using in vitro translated VDR and RXR and the
DR3-type VDRE of the rat ANF gene. Treatment of (A) free or
(B) already VDRE-bound VDR/RXR with MAMA/NO for 30 min at
30 °C resulted in a concentration-dependent inhibition
of VDR/RXR-DNA complex formation. C, subsequent addition of
1-10 mM DTT for 30 min at 30 °C nearly completely
reversed the inhibitory effect of MAMA/NO. D, treatment of
VDRE with MAMA/NO before addition of VDR/RXR resulted in a partial
inhibition of VDR/RXR-DNA complex formation at high MAMA/NO
concentrations. Representative gels are shown. Relative complex
formation is presented in reference to the control in the absence of
MAMA/NO. Columns represent means ± standard deviation of at least
three independent experiments.
|
|
Hydrogen Peroxide--
Incubating VDR/RXR with increasing
concentrations of H2O2 for 90 min at 30 °C
provided a significant inhibition of the VDR/RXR-DNA complex formation
only at relatively high H2O2 concentrations (>20 mM) with an IC50 value of 40-50
mM H2O2 (Fig.
2A). Similar high
H2O2 concentrations were needed to release
DNA-bound VDR/RXR from its response element (Fig. 2B).
Subsequent incubation with 10-20 mM DTT showed partial
restoration of the VDR/RXR-DNA complex formation (Fig. 2C).
DTT has been reported to complex zinc ions (34), but even high DTT
concentrations (20 mM) apparently did not lead to ejection
of Zn2+ from the zinc fingers as the DNA binding activity
of VDR/RXR was not affected by treatment with DTT alone (Fig.
2C). Pretreatment of the VDRE with
H2O2 prior to addition of VDR/RXR showed
similar IC50 values with regard to the formation of
VDR/RXR-DNA complexes compared with VDR/RXR (Fig. 2D).
View larger version (34K):
[in this window]
[in a new window]
|
Fig. 2.
Effects of H2O2 on
VDR/RXR-DNA complex formation in vitro. Gel shift
experiments were performed as described in the legend of Fig. 1.
Treatment of free (A) or already VDRE-bound (B) VDR/RXR with
H2O2 for 90 min at 30 °C resulted in a
concentration-dependent inhibition of the VDR/RXR-DNA
complex formation. C, subsequent addition of 10-20
mM DTT for 30 min at 30 °C partially reversed the
inhibitory effect of H2O2 on VDR/RXR-DNA
complex formation. D, treatment of VDRE with
H2O2 before addition of VDR/RXR plus 50 mM DTT to protect VDR/RXR from residual
H2O2 resulted in a
concentration-dependent inhibition of VDR/RXR-DNA complex
formation. Representative gels are shown. Relative complex formation is
presented in reference to the control in the absence of
H2O2. Columns represent means ± standard
deviation of at least three independent experiments.
|
|
Singlet Oxygen--
Singlet oxygen (1O2)
was generated by decomposition of NDPO2 (half-life at
37 °C: 23 min) leading to the formation of NDP and molecular oxygen
with a 1O2 yield of 50% (28, 35). Incubating
VDR/RXR with increasing concentrations of NDPO2 for 90 min
at 30 °C resulted in a concentration-dependent inhibition of the VDR/RXR-DNA complex formation with an
IC50 value of about 0.75 mM NDPO2,
while the decomposition product NDPO2deg showed no effects
(Fig. 3A). Already DNA-bound
VDR/RXR were found to be slightly more sensitive to NDPO2
(IC50 about 0.4 mM) compared with unbound
VDR/RXR (Fig. 3B). Subsequent incubation with up to 20 mM DTT for 30 min at 30 °C did not restore the
VDR/RXR-DNA complex formation (Fig. 3C). Experiments using
10 mM of the 1O2 quencher sodium
azide revealed partial protection from the effects of 5 mM
NDPO2 indicating that indeed 1O2
was the effective compound in our system (Fig. 3D).
Pretreatment of the VDRE with NDPO2 induced inhibition of
subsequent VDR/RXR-DNA complex formation with an IC50 value
of about 4 mM NDPO2 (Fig. 3E). This
is in line with the capability of 1O2 to
oxidatively modify DNA (36).
View larger version (35K):
[in this window]
[in a new window]
|
Fig. 3.
Effects of 1O2 on
VDR/RXR-DNA complex formation in vitro. Gel shift
experiments were performed as described in the legend of Fig. 1.
Treatment of free (A) or already VDRE-bound (B)
VDR/RXR with the singlet oxygen generator NDPO2 for 90 min
at 30 °C resulted in a concentration-dependent
inhibition of the VDR/RXR-DNA complex formation, while degraded
NDPO2 (NDPO2deg) had no effect. C,
subsequent addition of up to 20 mM DTT for 30 min at
30 °C did not restore VDR/RXR-DNA complex formation. D,
treatment of VDR/RXR with NDPO2 in the presence of 10 mM of the 1O2 quencher sodium azide
resulted in a partial protection of VDR/RXR showing that indeed
1O2 was the active compound. E,
treatment of VDRE with NDPO2 before addition of VDR/RXR
plus 50 mM DTT to protect VDR/RXR from residual-generated
1O2 resulted in a
concentration-dependent inhibition of VDR/RXR-DNA complex
formation. Representative gels are shown. Relative complex formation is
presented in reference to the control in the absence of
NDPO2. Columns represent means ± standard deviation
of at least three independent experiments.
|
|
Peroxyl Radicals--
Peroxyl radicals (ROO·) were
generated by decomposition of AAPH (half-life at 37 °C about
175 h) forming carbon-centered radicals that react swiftly with
O2 to yield ROO· (37). Incubating VDR/RXR with
increasing concentrations of AAPH for 2 h at 30 °C resulted in
a concentration-dependent inhibition of the VDR/RXR-DNA
complex formation with an IC50 value of about 10 mM AAPH (Fig.
4A). DNA-bound VDR/RXR was
found to be slightly more sensitive toward treatment with AAPH
(IC50 value of about 7 mM) than unbound VDR/RXR
(Fig. 4B). Subsequent treatment with up to 20 mM
DTT for 30 min at 30 °C did not restore VDR/RXR-DNA complex
formation (Fig. 4C). Pretreatment of the VDRE with AAPH prior to addition of VDR/RXR resulted in an IC50 value for
VDR/RXR-DNA complex formation of ~7 mM AAPH (Fig.
4D).
View larger version (31K):
[in this window]
[in a new window]
|
Fig. 4.
Effects of peroxyl radicals on VDR/RXR-DNA
complex formation in vitro. Gel shift experiments
were performed as described in the legend of Fig. 1. Treatment of free
(A) or already VDRE-bound (B) VDR/RXR with the
peroxyl radical generator AAPH for 2 h at 30 °C resulted in a
concentration-dependent inhibition of the VDR/RXR-DNA
complex formation. C, subsequent addition of up to 20 mM DTT for 30 min at 30 °C did not restore VDR/RXR-DNA
complex formation. D, treatment of VDRE with AAPH before
addition of VDR/RXR plus 50 mM DTT to protect VDR/RXR from
residual-generated peroxyl radicals resulted in a
concentration-dependent inhibition of VDR/RXR-DNA complex
formation. Representative gels are shown. Relative complex formation is
presented in reference to the control in the absence of AAPH. Columns
represent means ± standard deviation of at least three
independent experiments.
|
|
Peroxynitrite--
NO· and the superoxide anion radical
react to form the strong oxidant peroxynitrite anion
(ONOO), which has a half-life of about 1 s under
physiological conditions (24). We used chemically synthesized authentic
ONOO as a source. Prior to use, the 260 mM
ONOO stock solution was diluted with ice-cold deionized
H2O to a ONOO concentration of 6 mM. To avoid pH changes resulting from the use of up to 2 mM ONOO diluted from the 6 mM
ONOO stock solution, 200 mM phosphate buffer
was employed. This buffer had no effect on the VDR/RXR-DNA complex
formation (not shown). Incubating VDR/RXR with increasing
concentrations of ONOO for 10 min at room temperature
resulted in a concentration-dependent inhibition of the
VDR/RXR-DNA complex formation with an IC50 value in the
range of about 0.3-0.4 mM ONOO (Fig.
5A). Adding VDR/RXR various
times after the addition of ONOO showed that inhibition
of the VDR/RXR-DNA complex formation occurred within 10 s (data
not shown). Addition of multiple low ONOO doses (5 × 0.2 mM every minute) did not result in a shift of the
IC50 value for ONOO (not shown). DNA-bound
VDR/RXR were slightly more sensitive toward treatment with
ONOO (IC50 value of ~0.3 mM)
than unbound VDR/RXR (Fig. 5B). Subsequent treatment with up
to 20 mM DTT for 30 min at 30 °C did not restore the
VDR/RXR-DNA complex formation (Fig. 5C). Addition of 20 mM methionine nearly completely inhibited the inhibitory
effect of 1 mM ONOO (Fig. 5D).
Pretreatment of the VDRE with up to 5 mM ONOO
had no effect on VDR/RXR-DNA complex formation (Fig. 5E).
Higher concentrations of ONOO could not be used due to
problems with the pH and the salt concentration essential for optimal
VDR/RXR-DNA complex formation.
View larger version (39K):
[in this window]
[in a new window]
|
Fig. 5.
Effects of ONOO on VDR/RXR-DNA
complex formation in vitro. Gel shift experiments
were performed as described in the legend in Fig. 1. Treatment of free
(A) or already VDRE-bound (B) VDR/RXR with
authentic ONOO for 10 min at room temperature resulted in
a concentration-dependent inhibition of the VDR/RXR-DNA
complex formation, while degraded ONOO
(ONOO) had no effect. C,
subsequent addition of up to 20 mM DTT for 30 min at
30 °C did not restore VDR/RXR-DNA complex formation. D,
treatment of VDR/RXR with ONOO in the presence of 20 mM methionine resulted in a significant protection of
VDR/RXR-DNA complex formation. E, treatment of VDRE with up
to 5 mM ONOO before addition of VDR/RXR
showed no inhibitory effect on the VDR/RXR-DNA complex formation.
Representative gels are shown. Relative complex formation is presented
in reference to the control in the absence of ONOO.
Columns represent means ± standard deviation of at least three
independent experiments.
|
|
Studies with Living Cells--
To investigate whether the results
found in vitro are also found with cells, MCF-7 cells were
transiently transfected with expression vectors for VDR and RXR,
respectively, and a luciferase reporter gene plasmid under the control
of four copies of the rat ANF DR3-type VDRE (30). To avoid effects of
NO or reactive oxygen species on the protein expression values, the
cells were allowed to recover after transfection for 8 h and to
overexpress inactive VDR and RXR, respectively. Cells were then
stimulated with 100 nM VD or solvent (0.1% ethanol) for
various time intervals. Subsequent determination of the luciferase
activity revealed that VD activated the reporter gene time dependently
in a linear fashion: 2 h, 6.1 ± 0.6-fold; 4 h,
25.8 ± 2.7-fold; 6 h, 49.1 ± 4.9-fold; 8 h,
79.6 ± 4.8-fold; 10 h, 98.5 ± 6.6-fold; 24 h,
288.8 ± 22.6-fold (n = 3). To investigate effects
of NO· or reactive oxygen species on
VDR/RXR-dependent transcription, the 8 h time period
was chosen for further experiments. Only those concentrations of the
various compounds were used that proved to be subtoxic (
95% cell
viability investigated by morphological analysis and trypan blue
exclusion after 12 h compared with untreated controls).
Nitric Oxide--
To investigate effects of NO· on
VDR/RXR-dependent transcription we used DETA/NO, an
NO-donor similar to MAMA/NO but with a considerably longer half-life
(about 8 h at 37 °C). Increasing DETA/NO concentrations
inhibited VD-stimulated luciferase activity with an IC50
value of about 100 µM DETA/NO (Fig.
6A). In contrast, 1 mM of the control compound DETA had no effect (97.2 ± 4.1% of control) showing that indeed NO· was the effective
compound.
View larger version (25K):
[in this window]
[in a new window]
|
Fig. 6.
Effects of NO·, peroxides, or
intracellular O generation on
VDR/RXR-dependent reporter gene activity. MCF-7 cells
were transiently transfected with expression vectors for VDR and RXR,
respectively, together with a luciferase reporter gene plasmid under
the control of four copies of the rat ANF DR3-type VDRE. After 8 h
recovery and synthesis of inactive VDR/RXR, cells were stimulated with
100 nM VD or solvent (0.1% ethanol) in the presence of
various concentrations of DETA/NO (A),
H2O2 (B), t-BHP (C), or
the intracellular O generator DMNQ (D). Luciferase
activity normalized to the cellular protein content was quantified
after 8 h, and stimulation was calculated as the ratio of
luciferase activity of VD-stimulated cells to the luciferase activity
of solvent controls. All compounds investigated inhibited
VDR/RXR-dependent reporter gene activity in a
concentration-dependent manner. Data represent means ± standard deviation of at least three independent experiments.
|
|
Hydroperoxides--
The transfected cells were incubated with
increasing concentrations of hydroperoxides.
H2O2 induced a
concentration-dependent inhibition of VD-stimulated
luciferase activity with an IC50 value of about 300 µM H2O2 (Fig. 6B).
t-BHP also induced a concentration-dependent inhibition of
the reporter gene activity with an IC50 value of about 45 µM t-BHP (Fig. 6C).
Intracellular Superoxide--
As source for intracellular
O generation we used the redox cycler DMNQ. Fig.
6D shows that DMNQ inhibited VDR/RXR-dependent
luciferase activity in a concentration-dependent manner
with an IC50 value of about 5 µM DMNQ.
Reversibility--
To ask whether the observed effects on zinc
finger-dependent transcription in cells are reversible, the
transiently transfected cells were treated with the IC50
value concentrations of DETA/NO, H2O2, t-BHP,
or DMNQ for 8 h. After a recovery period of 5 h in fresh
medium, cells were stimulated with 100 nM VD or solvent vehicle for another 8 h. After treatment with 100 µM
DETA/NO, 50 µM t-BHP, or 5 µM DMNQ, cells
could again be stimulated by VD for reporter gene activity (about 80%
restored activity compared with non-pretreated controls). After
treatment with 300 µM H2O2, however, cells were refractory to stimulation with VD (Fig.
7).
View larger version (30K):
[in this window]
[in a new window]
|
Fig. 7.
Reversibility of the inhibitory effects of
NO·, peroxides, or intracellular O
generation on VDR/RXR-dependent reporter gene
activity. MCF-7 cells were transiently transfected as described in
the legend of Fig. 6. After 8 h recovery and synthesis of inactive
VDR/RXR, cells were incubated with (empty bars) or without
(black bars) 100 nM VD or solvent (0.1%
ethanol) and treated for an additional 8 h with the
IC50 value concentrations of DETA/NO,
H2O2, t-BHP or the intracellular O
generator DMNQ. Cells cultured without VD (black
bars) were then incubated in fresh medium for 5 h to recover
from nitrosative or oxidative stress and were subsequently stimulated
with 100 nM VD or solvent (0.1% ethanol) for additional
8 h. Luciferase activity normalized to the cellular protein
content was quantified, and stimulation was calculated as the ratio of
luciferase activity of VD-stimulated cells to the luciferase activity
of solvent controls. While the inhibitory effects of NO·, t-BHP,
and DMNQ, respectively, were reversible, the effect of
H2O2 proved to be irreversible. Data represent
means ± standard deviation of three independent
experiments.
|
|
With this experimental design it was not possible to discriminate
whether the disrupted zinc finger transcription factors were repaired
by the cells or whether they were synthesized de novo.
Therefore, transiently transfected cells were treated for 8 h with
the IC50 value concentrations of DETA/NO,
H2O2, t-BHP, or DMNQ, and were then stimulated
with 100 nM VD or solvent vehicle in the presence of the
translational inhibitor cycloheximide. Treatment with 10 µg/ml
cycloheximide inhibited VD-stimulated luciferase activity by 96.3 ± 2.6% (n = 3). RNA was isolated after 15 h, and
RT-PCR was performed to quantitate VDR/RXR-dependent luciferase mRNA expression. Fig. 8
shows that VD stimulated luciferase mRNA expression about 7-fold
compared with unstimulated cells. Interestingly, even in the presence
of cycloheximide a pretreatment of the cells with 100 µM
DETA/NO for 8 h still allowed full stimulation of
VDR/RXR-dependent luciferase mRNA expression by VD
(Fig. 8, lane 4). In contrast, pretreatment with 300 µM H2O2, 50 µM
t-BHP, or 5 µM DMNQ allowed for subsequent significant
VD-stimulated luciferase mRNA expression (Fig. 8, lanes
5-7). These results demonstrate that cells are able to repair
VDR/RXR after nitrosative stress but not after oxidative stress.
View larger version (38K):
[in this window]
[in a new window]
|
Fig. 8.
Cells repair VDR/RXR after nitrosative but
not after oxidative stress. MCF-7 cells were transiently
transfected as described in the legend of Fig. 6. After 8 h of
recovery and synthesis of inactive VDR/RXR, cells were incubated in the
absence (lanes 1-3) or presence of 100 µM
DETA/NO (lane 4), 300 µM
H2O2 (lane 5), 50 µM
t-BHP; (lane 6), or 5 µM of the
intracellular O generator DMNQ (lane 7)
for 8 h. Cells were then treated without (lane 2) or
with 100 nM VD (lanes 1 and 3-7) or
solvent (0.1% ethanol, lane 2) in the presence of 10 µg/ml cycloheximide (CHX) to inhibit de novo
synthesis of VDR and RXR. After 15 h, total cellular RNA was
isolated and luciferase- and GAPDH-specific RT-PCR was performed. Under
these conditions, VDR/RXR-dependent luciferase mRNA
expression was not affected after nitrosative stress, while after
oxidative stress luciferase mRNA expression was significantly
inhibited. Values are means ± standard deviation of three
independent experiments.
|
|
| |
DISCUSSION |
In Vitro--
Nitric oxide is able to nitrosate cysteine thiols in
proteins yielding S-nitrosothiols under aerobic conditions,
probably via formation of N2O3 (13, 38). In the
case of zinc fingers, which contain up to four cysteines, this reaction
leads to the ejection of the zinc ion (15, 39, 40). However, various sulfhydryl-oxidizing compounds such as H2O2
(41-46), O (47), ONOO (48), HOCl (45, 48), or
diamide (46, 49), as well as sulfhydryl-alkylating compounds, such as
iodoacetamide (43, 46) or N-ethylmaleimide (43, 50), have
also been shown to disrupt zinc-sulfur complexes in various proteins.
To investigate whether effects of nitrosative and oxidative stress,
respectively, differ with regard to zinc finger-dependent transcription factors, we treated VDR/RXR with compounds that are
generated under conditions of nitrosative and/or oxidative stress,
i.e. NO·, H2O2,
1O2, ROO·, or ONOO. All
these compounds inhibited the DNA complex formation and thus the DNA
binding activity of VDR/RXR in a concentration-dependent manner (Figs. 1A-5A). The millimolar range of the
IC50 value for H2O2 appears to be
rather high but fits with concentrations reported for the inhibition of
other zinc finger transcription factors, such as the glucocorticoid
receptor (41, 42, 51), Sp1 (43, 44), or the estrogen receptor (49).
Prebinding to its response element did not protect VDR/RXR from the
inhibitory effects (Figs. 1B-5B). In contrast, we found
equal or in some cases even slightly enhanced sensitivity of prebound
VDR/RXR to our treatment regimens as compared with pretreating VDR/RXR
before DNA binding. This suggests that the zinc fingers within the
protein-DNA complex are equally or even slightly more accessible for
the reactive species investigated. From a functional point of view this
means that reactive species are able to inhibit zinc
finger-dependent gene expression during all phases of their action.
NO-induced inhibition of VDR/RXR-DNA complex formation was nearly
completely restored by subsequent reduction with DTT (Fig. 1C). After treatment with H2O2, DTT
was able to partially (about 50%) restore the DNA complex formation of
VDR/RXR (Fig. 2C). In contrast, treatment with
1O2, ROO·, or ONOO led to
irreversible loss of the DNA binding activity of VDR/RXR since DTT did
not restore VDR/RXR-DNA complex formation (Figs. 3C-5C).
This suggests that H2O2,
1O2, ROO·, or ONOO at
least partially oxidize zinc finger cysteine thiols (Cys-SH) beyond the
level of disulfides, e.g. to sulfinic
(Cys-SO2H), sulfonic acid (Cys-SO3H) (21), or
even thiosulfinate (Cys-SO-S-Cys) (52). Among the amino acids present
in proteins only cysteines are modified by NO· (53), while the
other reactive species used in this study are able to modify additional
amino acids in proteins also, e.g. to oxidize methionine to
methionine sulfoxide (49, 54) or, in the case of ONOO, to
nitrate tyrosine (55, 56). VDR and RXR each contain 17 methionine
residues, which in part are located close to the zinc finger domains,
and RXR even contains four tyrosines within or nearby the DNA binding
domain. All these amino acid modifications may contribute to inhibition
of the VDR/RXR-DNA complex formation by H2O2,
1O2, ROO·, or ONOO.
The reactive species NO·, H2O2,
1O2, and ROO· were found to affect in a
dose-dependent manner the VDRE such that subsequently added
VDR/RXR were unable to bind anymore. Interestingly, the ratios of the
IC50 values found for VDRE and for VDR/RXR were calculated
as >10 for NO·, ~1 for H2O2, 10 for
1O2, ~1 for ROO·, and >10 for
ONOO. This provides NO· with an unique profile
among the studied reactive species, as the disruption of zinc fingers
by NO· is reversible and a more than 10-fold higher NO·
concentration is needed to half-maximally inactivate the VDRE compared
with VDR/RXR.
Cells--
Using living cells, we investigated whether effects of
nitrosative stress on VDR/RXR-dependent transcription
differ from effects of oxidative stress. Transiently transfected cells
were treated with subtoxic concentrations of the NO-donor DETA/NO, the
two hydroperoxides H2O2 and
tert-butyl hydroperoxide, and the redox cycler DMNQ, which
generates O intracellularly. The 1O2
generator NDPO2 could not be used as degraded
NDPO2 for unknown reasons showed a strong inhibitory effect
on VDR/RXR-dependent luciferase reporter gene activity
(data not shown). All compounds investigated inhibited the
VDR/RXR-dependent luciferase activity in a
concentration-dependent manner (Fig. 6). The
IC50 value found using t-BHP was about 7-fold lower than
the IC50 value found for H2O2. This
may be due to several cellular depletion pathways for H2O2 (e.g. glutathione peroxidase,
catalase) and/or the direction of the hydrophobic t-BHP to more
hydrophobic cellular compartments than H2O2.
The IC50 value found for H2O2
in vitro (40-50 mM) and that found in living cells
(300 µM) differed considerably. However, differences of
two orders of magnitude have also been reported using the zinc finger
transcription factor glucocorticoid receptor (51).
In living cells reversibility of the inhibition of VDR/RXR-DNA complex
formation was shown after exposure to NO·, t-BHP, or DMNQ at
concentrations corresponding to the IC50 values (Fig. 7).
In contrast, after treatment with H2O2 no
reversibility was found, although the H2O2
concentration used (300 µM) also proved to be nontoxic.
This suggests that in addition to VDR/RXR-dependent transcription additional cellular functions are affected by
H2O2.
To investigate whether cells are able to repair disrupted VDR/RXR, we
treated cells with the reactive species and then stimulated them with
VD in the presence of the translational inhibitor cycloheximide, which
inhibited de novo synthesis of VDR/RXR. Results showed that after exposure to NO· VDR/RXR-dependent luciferase
mRNA expression was unaffected (Fig. 8). In contrast, exposure to
the hydroperoxides or DMNQ resulted in significantly impaired
expression of luciferase mRNA. These results suggest that cells are
able to repair S-nitrosated zinc finger transcription
factors, whereas oxidative modifications appear to be irreversible.
Although in this study only transcription factors containing
Cys4-type zinc fingers were investigated, experiments using
the Cys2/His2-type zinc finger transcription
factor Sp1 have shown, that the latter is also very susceptible toward
H2O2 (43, 44) and NO· (17, 19),
respectively. As all known zinc finger structures contain at least two
cysteine residues (14) and as the cysteine thiols are the
redox-sensitive parts of zinc fingers it is highly likely that our
results can be generalized with regard to all zinc-finger containing
transcription factors. Whether nitrosative or oxidative stress also
affects the translocation of zinc finger transcription factors into the
nucleus remains to be determined.
In conclusion, we suggest that the more gentle reactions of NO·
permit cells to handle nitrosation of zinc finger transcription factors with minimal collateral damage as hypothesized recently (57)
compared with the irreversible modifications induced by oxidative
stress. Thus, NO· is not just another compound able to destroy
zinc finger domains of proteins but may potentially serve as a gene
regulatory molecule during inflammatory reactions.
§,
TOP
ABSTRACT
INTRODUCTION
Fellow of the National Foundation for Cancer Research,
Bethesda, MD.
