A Fenton reaction at the endoplasmic reticulum is involved in the redox control of hypoxia-inducible gene expression.
ABSTRACT It has been proposed that hydroxyl radicals (.OH) generated in a perinuclear iron-dependent Fenton reaction are involved in O(2)-dependent gene expression. Thus, it was the aim of this study to localize the cellular compartment in which the Fenton reaction takes place and to determine whether scavenging of.OH can modulate hypoxia-inducible factor 1 (HIF-1)-dependent gene expression. The Fenton reaction was localized by using the nonfluorescent dihydrorhodamine (DHR) 123 that is irreversibly oxidized to fluorescent rhodamine 123 while scavenging.OH together with gene constructs allowing fluorescent labeling of mitochondria, endoplasmic reticulum (ER), Golgi apparatus, peroxisomes, or lysosomes. A 3D two-photon confocal laser scanning microscopy showed.OH generation in distinct hot spots of perinuclear ER pockets. This ER-based Fenton reaction was strictly pO(2)-dependent. Further colocalization experiments showed that the O(2)-sensitive transcription factor HIF-1alpha was present at the ER under normoxia, whereas HIF-1alpha was present only in the nucleus under hypoxia. Inhibition of the Fenton reaction by the.OH scavenger DHR attenuated HIF-prolyl hydroxylase activity and interaction with von Hippel-Lindau protein, leading to enhanced HIF-1alpha levels, HIF-1alpha transactivation, and activated expression of the HIF-1 target genes plasminogen activator inhibitor 1 and heme oxygenase 1. Further,.OH scavenging appeared to enhance redox factor 1 (Ref-1) binding and, thus, recruitment of p300 to the transactivation domain C because mutation of the Ref-1 binding site cysteine 800 abolished DHR-induced transactivation. Thus, the localized Fenton reaction appears to impact the expression of hypoxia-regulated genes by means of HIF-1alpha stabilization and coactivator recruitment.
Article: The role of mitochondria in the regulation of hypoxia-inducible factor 1 expression during hypoxia.[show abstract] [hide abstract]
ABSTRACT: Hypoxia-inducible factor 1 (HIF-1) is a heterodimeric transcription factor that regulates transcriptional activation of several genes responsive to the lack of oxygen, including erythropoietin, vascular endothelial growth factor, glycolytic enzymes, and glucose transporters. Because the involvement of mitochondria in the regulation of HIF-1 has been postulated, we tested the effects of mitochondrial electron transport chain deficiency on HIF-1 protein expression and DNA binding in hypoxic cells. The neurotoxin 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) inhibits electron transport chain at the level of complex I. MPTP is first converted to a pharmacologically active metabolite 1-methyl-4-phenylpyridinum (MPP+). MPP+ effectively inhibited both complex I activity and hypoxic accumulation of HIF-1alpha protein in dopaminergic cell lines PC12 and CATH.a. In C57BL/6 mice, a single dose of MPTP (15 mg/kg, intraperitoneal) inhibited complex I activity and HIF-1alpha protein accumulation in the striatum in response to a subsequent hypoxic challenge (8% O(2), 4 h). In a genetic model system, 40% complex I-inhibited human-ape xenomitochondrial cybrids, hypoxic induction of HIF-1alpha was severely reduced, and HIF-1 DNA binding was diminished. However, succinate, the mitochondrial complex II substrate, restored the hypoxic response in cybrid cells, suggesting that electron transport chain activity is required for activation of HIF-1. A partial complex I deficiency and a mild reduction in intact cell oxygen consumption effectively prevented hypoxic induction of HIF-1alpha protein.Journal of Biological Chemistry 12/2000; 275(46):35863-7. · 4.77 Impact Factor
Article: Three-dimensional organization of microtubules in tumor cells studied by confocal laser scanning microscopy and computer-assisted deconvolution and image reconstruction.[show abstract] [hide abstract]
ABSTRACT: Confocal microscopy, image deconvolution and computer-assisted methods have been used to reconstruct the three-dimensional (3-D) distribution of tubulin in cells. The techniques were applied to tumor cells growing under regular culture conditions (planar cultivation) and those penetrating into reconstituted collagen matrices (spatial cultivation). As expected the application of deconvolution algorithms enhanced the resolution of results. The deconvolution using the maximum likelihood estimation included the measurement of the point spread function of the optical setup. The data visualization of the resulting data sets uses volume as well as surface rendering approaches. The 3-D reconstruction gives a clear image of the spatial arrangement of tubulin fibers in relation to cell shape and position of other cellular organelles, particularly the nucleus. The tubulin forms an intricate network of fibers of variable thickness. The highest tubulin concentrations appear in the cell periphery and particularly in pseudopodia/invadopodia. This is indicative of an enhanced transport of intracellular material facilitating cell movement and lysis of the extracellular matrix. The investigation is assumed to stimulate further experiments using a variety of techniques leading to the complete understanding of the spatial architecture of living cells.Cells Tissues Organs 02/2000; 167(1):1-8. · 2.20 Impact Factor
Article: Inhibition of CREB- and cAMP response element-mediated gene transcription by the immunosuppressive drugs cyclosporin A and FK506 in T cells.[show abstract] [hide abstract]
ABSTRACT: The clinically important immunosuppressant drugs cyclosporin A and FK506 (tacrolimus) inhibit in T-cells calcineurin phosphatase activity and nuclear translocation of the cytosolic component of the transcription factor nuclear factor of activated T-cells (NF-ATc) that is involved in the induction of early genes during T-cell activation. This effect has been proposed to explain at least part of the immunosuppressive effect of these drugs. Previous studies in pancreatic islet cell lines have shown that cyclosporin A and FK506 through inhibition of calcineurin interfere also with the function of the transcription factor cAMP response element binding protein (CREB) that is activated by cAMP and calcium signals and binds to cAMP/calcium response elements (CRE). By transient expression of CRE-reporter genes or GAL4-CREB fusion proteins, the present study shows that inhibition of CREB/CRE-directed transcription by cyclosporin A and FK506 occurs in a great variety of cell types including in cell lines derived from tissues in which adverse effects of the immunosuppressants develop. CREB activity and CRE-mediated transcription was blocked by these drugs also in Jurkat T-cells. When taken together with recent evidence for an essential role of CREB in T-cell activation and proliferation, the present results suggest that inhibition of CREB/CRE-directed transcription may be a molecular mechanism of the immunosuppressive effect of cyclosporin A and FK506.Archiv für Experimentelle Pathologie und Pharmakologie 11/1997; 356(4):433-40. · 2.65 Impact Factor
A Fenton reaction at the endoplasmic reticulum is
involved in the redox control of hypoxia-inducible
Qing Liu*†, Utta Berchner-Pfannschmidt†‡, Ulrike Mo ¨ller*, Martina Brecht*, Christoph Wotzlaw‡, Helmut Acker‡,
Kurt Jungermann*§, and Thomas Kietzmann*¶
*Institut fu ¨r Biochemie und Molekulare Zellbiologie, Humboldtallee 23, D-37073 Go ¨ttingen, Germany; and‡Max-Planck-Institut fu ¨r Molekulare Physiologie,
Otto-Hahn-Strasse 11, D-44227 Dortmund, Germany
Communicated by Robert E. Forster, University of Pennsylvania School of Medicine, Philadelphia, PA, January 13, 2004 (received for review
December 6, 2002)
It has been proposed that hydroxyl radicals (?OH) generated in a
perinuclear iron-dependent Fenton reaction are involved in O2-
dependent gene expression. Thus, it was the aim of this study to
localize the cellular compartment in which the Fenton reaction
takes place and to determine whether scavenging of ?OH can
modulate hypoxia-inducible factor 1 (HIF-1)-dependent gene ex-
pression. The Fenton reaction was localized by using the nonfluo-
fluorescent rhodamine 123 while scavenging ?OH together with
gene constructs allowing fluorescent labeling of mitochondria,
endoplasmic reticulum (ER), Golgi apparatus, peroxisomes, or ly-
sosomes. A 3D two-photon confocal laser scanning microscopy
showed ?OH generation in distinct hot spots of perinuclear ER
pockets. This ER-based Fenton reaction was strictly pO2-depen-
dent. Further colocalization experiments showed that the O2-
sensitive transcription factor HIF-1? was present at the ER under
normoxia, whereas HIF-1? was present only in the nucleus under
hypoxia. Inhibition of the Fenton reaction by the ?OH scavenger
DHR attenuated HIF-prolyl hydroxylase activity and interaction
genes plasminogen activator inhibitor 1 and heme oxygenase 1.
binding and, thus, recruitment of p300 to the transactivation
domain C because mutation of the Ref-1 binding site cysteine 800
abolished DHR-induced transactivation. Thus, the localized Fenton
reaction appears to impact the expression of hypoxia-regulated
genes by means of HIF-1? stabilization and coactivator recruitment.
contradictory evidence exists concerning the nature of the
sensor. First, it was proposed that heme proteins (1) such as
NADPH oxidases (2) or as exist within the mitochondrial
electron-transport chain (3, 4) may act as O2sensors by produc-
ing reactive oxygen species (ROS) as ‘‘second messengers.’’
Recently, it was proposed that a family of O2-using prolyl and
asparagyl hydroxylases involved in the modification of HIFs may
act as direct O2sensors (5).
The role of ROS as O2messengers has been supported by the
finding that, similar to a typical response to hypoxia, treatment
of healthy human volunteers with the antioxidant N-acetyl-
cysteine enhanced the hypoxic ventilatory response and eryth-
ropoietin (EPO) concentration in blood (6). Thus, N-acetyl-
cysteine or its biochemical derivatives, cysteine, and glutathione
mimic hypoxia. In contrast, the ROS H2O2mimicked normoxia
in the expression of O2-dependent genes in cell cultures (7).
We have demonstrated (7, 8) that an H2O2-degrading perinu-
clear Fenton reaction [H2O2? Fe2?3 Fe3?? OH?? hydroxyl
radical (?OH)] was involved in O2signaling. However, it remained
undetermined in which cellular compartment the Fenton reaction
he heterogeneous pO2 distribution in tissues requires an
O2-sensing system to adapt organ functions. Until now,
takes place and whether also transcription factors regulating the
O2-dependent gene expression are located in this compartment.
A transcription factor who gained a central role in the
O2-dependent modulation of gene expression is hypoxia-
inducible factor 1 (HIF-1) (9, 10). HIF-1 is composed of the O2-
and redox-sensitive ?-subunit and the constitutively expressed
HIF-1? (ARNT) subunit. Although other HIF isoforms appear
to exist (5), HIF-1 is considered to be the major regulator of
many physiologically important genes, including plasminogen
activator inhibitor 1 (PAI-1), vascular endothelial growth factor
(VEGF), and heme oxygenase 1 (HO-1) (11). Thus, it appears
likely that the redox-sensitive HIF-1? may be a target within an
O2-sensing system involving ROS and a Fenton reaction. To
characterize the ROS and redox-sensitive part of the oxygen-
sensing pathway further, it was the aim of this study to identify the
compartment in which the Fenton reaction takes place and to
investigate whether a compartment-specific inhibition of ?OH gen-
eration can interfere with the HIF-1-dependent gene expression.
Materials and Methods
All biochemicals and enzymes used were of analytical grade and
obtained from commercial suppliers.
Cell Culture. HepG2 cells were kept in a normoxic atmosphere of
for 24 h. At 24 h, medium was changed and culture was continued at
normoxia (16% O2) or hypoxia (8% O2) (87% N2?5% CO2, vol).
Plasmid Constructs. The pGL3-EPO-hypoxia response element
(HRE) luciferase plasmid, containing three EPO-HREs, was de-
scribed (12). For pDsRed-endoplasmic reticulum (ER), DsRed
the NheI and BglII sites of pECFP-ER (Clontech). The pDsRed-
Golgi and pDsRed-Mito constructs were constructed by replacing
the BamHI and NotI enhanced cyan fluorescent protein (ECFP)
fragment with pDsRed-N1 in pECFP-Golgi and pECFP-Mito.
The plasmid pECFP-Peroxi was obtained from Clontech.
The pGL3PAI-766, containing the rat PAI-1 promoter from
?766 to ?33, as well as pGL3PAI-766M2, were described (13).
The plasmids GFP-HIF1? (14), pG5E1B-LUC (15), pSG424
(19), Gal4-HIF1?-transactivation domain N (TADN), and
HIF1?-TADC, as well as the Gal4-HIF1?-TADCM construct
Abbreviations: ?OH, hydroxyl radical; HIF-1, hypoxia-inducible factor 1; DHR, dihydrorho-
damine; ER, endoplasmic reticulum; VHL, von Hippel–Lindau; TADx; transactivation do-
ECFP, enhanced cyan fluorescent protein; RH, rhodamine; GM, Golgi membrane; 2P-CLSM,
two-photon laser confocal microscopy.
†Q.L. and U.B.-P. contributed equally to this work.
§Deceased May 10, 2002.
¶To whom correspondence should be addressed. E-mail: email@example.com.
© 2004 by The National Academy of Sciences of the USA
March 23, 2004 ?
vol. 101 ?
with an exchange of cysteine 800 into serine, were described (16,
17). The Gal4-HIF1?-TADNM construct, in which proline P564
was changed to alanine, was constructed by using the Quick-
Change mutagenesis kit (Promega).
The plasmid pGEX-TADN was constructed by ligation of a
219-bp PCR fragment from rat HIF-1?, encompassing amino acids
Biosciences), allowing generation of a GST fusion protein. The
into Escherichia coli BL21(DE3), and protein expression was in-
duced by using 0.1 mM isopropyl ?-D-thiogalactoside for 4 h. After
MgCl2?1 mM DTT?10% glycerol), the clarified sonicates were
incubated with glutathione Sepharose 4B (Amersham Biosciences)
for 1 h at 4°C. After four washes in PBS, the GST-TADN and
GST proteins were eluted in 50 mM Tris?HCl?10 mM reduced
glutathione (pH 8.0). The integrity was assessed by SDS?PAGE,
followed by Coomassie blue staining.
HIF-TADN Hydroxylation and von Hippel–Lindau (VHL) Pull-Down As-
say. HepG2 cells homogenized at 4°C in 250 mM sucrose?50 mM
Tris?HCl (pH 7.5) were centrifuged at 1,000 ? g for 10 min, and
the supernatant was centrifuged at 3,000 ? g for 10 min. Again,
supernatant was pelleted at 18,000 ? g for 10 min and suspended
in 40 mM Tris?HCl (pH 7.5). Extracts (300 ?g?ml) were incu-
bated at 37°C for 30 min in 40 mM Tris?HCl (pH 7.5)?0.5 mM
DTT?50 ?M ammonium ferrous sulfate?1 mM ascorbate?2
mg/ml BSA?0.4 mg/ml catalase?50,000 dpm of 5-[14C]2-
oxoglutarate (Perkin–Elmer)?0.1 mM unlabeled 2-oxoglutarate
with either 20 ?g of GST protein or GST-TADN protein in the
presence of various concentrations of dihydrorhodamine (DHR)
or RH. Radioactivity associated to succinate was determined as
described (18). The basal GST-dependent activity was sub-
tracted from the GST-TADN-dependent activity.
For VHL pull-down assay,35S-VHL was synthesized from
pCMV-HA-VHL by using [35S]methionine and the TNT-
coupled reticulocyte lysate system (Promega). The GST-TADN
was incubated either without extract or with cell extracts, as
described for the hydroxylation assay, either without or with 0.5
?M DHR or RH, respectively. The reaction products were
incubated at 4°C in 200 ?l of buffer (50 mM Tris?HCl, pH 8?120
mM NaCl?0.5% Nonidet P-40), supplemented with glutathione-
Sepharose beads and 50,000 dpm of [35S]-VHL. After 2 h, beads
were washed three times with cold buffer (20 mM Tris?HCl, pH
8?100 mM NaCl?1 mM EDTA?0.5% Nonidet P-40). Bound
proteins were eluted in 10 mM reduced glutathione and analyzed
by SDS?PAGE, followed by autoradiography.
RNA Preparation, Northern Blot Analysis, and Western Blot Analysis.
Isolation of RNA and Northern blot analysis for PAI-1 and
HO-1 were performed as described (19). Western blot analysis
was carried out as described (20). In brief, media or lysates from
or 7.5% SDS-polyacrylamide gel and, after electrophoresis and
blotting, probed with mouse mAb directed against PAI-1 (1:200
dilution; American Diagnostics, Pendleton, IN), HO-1 (1:500 di-
lution; Biomol, Plymouth Meeting, PA), or human HIF-1? (1:2000
dilution; BD Biosciences), or with a rabbit Ab directed against
Golgi membrane (GM; 1:2,000 dilution; Bioscience, Go ¨ttingen,
Germany). The enhanced chemiluminescence system (ECL;
Amersham Biosciences) was used for detection.
Cell Transfection and Luciferase Assay. We transfected ?4 ? 105
cells per 60-mm dish, as described (20). In brief, 2.5 ?g of
pGl3-EPO-HRE LUC, pGL3PAI-766, and pGL3PAI-796M2
was transfected. After 5 h, medium was changed and cells were
cultured under normoxia for 19 h. Then, medium was changed
again and cells were stimulated with 6 ?M DHR or rhodamine
(RH) and cultured further for 24 h under normoxia or hypoxia.
Transfection efficiency was controlled by cotransfection of 0.25 ?g
of Renilla Luciferase expression vector (pRLSV40; Promega).
To investigate HIF-1? transactivation, 2 ?g of pG5-E1B-LUC
was cotransfected with 500 ng of pSG424 or Gal4-TADN?
TADC. To investigate HIF-1? nuclear translocation, 2.5 ?g of
GFP-HIF1? was transfected, medium was changed after 18 h,
and cells were then cultured under normoxia for 30 h. Cells were
then stimulated with either 6 ?M DHR or 100 ?M PDTC.
Measurement of ?OH Generation by Two-Photon Laser Confocal Mi-
croscopy (2P-CLSM). Intracellular ?OH generation by the Fenton
reaction can be detected by irreversible conversion of nonfluo-
rescent DHR into fluorescent RH 123 (21).
Cells were transfected with 5 ?g of the pDsRed-ER, pDsRed-
Mito, pDsRed-Golgi, pECFP-Golgi, pECFP-ER, pECFP-Mito,
pDsRed-Peroxi, or pDsRed-Lyso gene constructs. Medium was
changed after 19 h, and cells were cultured for 24 h. Cells were
then transferred to a culture chamber on the microscope,
enabling observation at 37°C under a variable gas atmosphere.
Cells were treated with 30 ?M DHR (Molecular Probes) for 5
min under normoxic or hypoxic conditions, and they were then
washed without altering the pO2, as controlled by a polaro-
graphic pO2catheter electrode (Licox, Kiel, Germany). The RH
123 fluorescence was recorded by using 2P-CLSM (8), and an
850-nm excitation wavelength was applied to colocalize RH 123
fluorescence (emission 530 nm) and ECFP fluorescence (emis-
sion 470 nm). We used an 875-nm excitation for RH 123
fluorescence and DsRed fluorescence (emission 580 nm). Flu-
orescence was registered by a photo multiplier, digitized, and
visualized by the EZ 2000 software (Version 2.1.4; Coord Au-
tomatisering, Hilversum, The Netherlands). Deconvolution,
data reconstruction, and calculation of isosurfaces were per-
formed on a Unix-based OCTANE workstation (Silicon Graphics,
Mountain View, CA), essentially as described (22).
Immunohistochemistry. HepG2 cells were placed on coverslips,
transfected with pECFP-ER, and exposed to various pO2dis-
tributions, as described above. Cells were fixed by ice-cold
methanol?acetone (1:1) for 10 min, blocked with 3% BSA in
PBS, and incubated with an anti-HIF-1? mAb (1:50 dilution;
Transduction Laboratories, Lexington, KY) and a rabbit Living
Colors Av peptide Ab (1:20 dilution; Clontech). Cells were washed
in PBS before incubation with an Alexa Fluor 568-conjugated goat
anti-mouse IgG secondary Ab (1:400 dilution; Molecular Probes)
and a Cy2-conjugated goat anti-rabbit IgG secondary Ab (1:200
cells were captured by using 2P-CLSM, as described above, and
images were visualized in false colors, as indicated.
Localization of ?OH-Mediated RH Fluorescence at the ER. Because
?OH-mediated conversion of DHR to fluorescent RH 123 is
localized in a perinuclear space (8), we aimed to identify this
subcellular compartment. HepG2 cells transfected with pD-
sRed-ER, pECFP-ER, pDsRed-Mito, pECFP-Mito, pECFP-
Golgi, pDsRed-Peroxi, and pDsRed-Lyso were treated with 30
?M DHR for 5 min in a microscope tissue-culture chamber
under normoxia. In a total of 20 experiments (pDsRed-ER, n ?
3; pECFP-ER, n ? 3; pDsRed-Mito, n ? 2; p5CFP-Mito, n ? 2;
pDsRed-Peroxy, n ? 2; pDsRed-Lyso, n ? 2), RH 123 fluores-
cence and ECFP or DsRed fluorescence were recorded simul-
taneously by 2P-CLSM without any crosstalk because of the
emission-filter settings. After deconvolution and reconstruction,
DsRed or ECFP fluorescence overlapped with RH 123 fluores-
cence, indicating the site of ?OH generation.
The 3D reconstruction revealed that RH 123 fluorescence
overlapped with DsRed or ECFP fluorescence when expressed in
Liu et al.PNAS ?
March 23, 2004 ?
vol. 101 ?
no. 12 ?
when the ECFP or DsRed proteins were expressed in the mito-
chondria, Golgi apparatus, peroxisomes, or lysosomes (Fig. 1).
To investigate whether ?OH generation at ER is pO2-dependent,
the pECFP-ER-transfected cells exposed for 1 h to hypoxia were
treated with DHR. Under hypoxia, RH 123 fluorescence was
reached within the microscope chamber (15 min after returning to
normoxia), a clear overlap of ?OH generation and ECFP fluores-
cence in the ER was detected (Fig. 2A). The pO2-dependent ?OH
generation was supported further by applying a photoreduction
approach (23). The photoreduction effect is caused by blue-light
illumination initiating a reduction of flavins and, concomitantly, an
activation of flavin-containing oxidases. Subsequently, these oxi-
dases reduce O2to H2O2, which is then degraded to ?OH by the
Fenton reaction. Thus, this light reaction, showing correct imaging
of intracellular oxygen radical kinetics, is a control for intracellular
?OH production. Furthermore, it indicates that O2absence is the
major trigger for reduced intracellular ?OH production under
even under hypoxia. The challenge of pECFP-ER transfected cells
RH 123 fluorescence at the ER. However, this increase was seen
predominantly under normoxia; under hypoxia, ?OH generation
was minimal (Fig. 2A). Although photoreduction can be seen also
in mitochondria because they contain FADH for generation of
?OH generation at ER under normal conditions (Fig. 2A) because
FADH reduction and ?OH generation in mitochondria occurred
only on photoreduction.
Oxygen-Dependent ?OH Generation and Colocalization of HIF-1? with
the ER Under Normoxia. To investigate whether ?OH generation at
at the ER. For this examination, cells transfected with pECFP-ER
were incubated under normoxia (60 min) or hypoxia (60 min) and
immunostained with an anti-HIF-1? Ab. Under normoxia, the
fluorescence indicating ER overlapped with the fluorescence in-
dicating the presence of HIF-1?. Under hypoxia, in contrast, the
ER fluorescence and the HIF-1? fluorescence were dissociated
with all of the HIF-1? fluorescence within the nucleus (Fig. 2B).
Induction of HIF-1-Dependent Genes by DHR. To investigate whether
scavenging of ?OH by DHR at ER interferes with the O2signal
modulating HIF-1-dependent genes such as PAI-1 and HO-1,
cells were treated with DHR under normoxia and hypoxia, and
PAI-1 and HO-1 expression were measured.
Hypoxia induced PAI-1 and HO-1 mRNA expression. Simi-
larly, treatment with DHR induced PAI-1 and HO-1 mRNA
under normoxia. Treatment with DHR under hypoxia did not
further enhance mRNA expression of both genes. The DHR-
induced mRNA increase was followed by an increase in PAI-1
and HO-1 proteins (Fig. 3).
HIF-1 activates gene expression by binding to HREs. In cells
transfected with pGL3PAI-766, LUC activity was induced 2-fold
under hypoxia (Fig. 4). The treatment with DHR increased LUC
activity 3-fold under normoxia and 3.5-fold under hypoxia (Fig. 4).
and DHR-mediated induction of LUC activity (Fig. 4). Further-
pDsRed-ER (Right), pECFP-ER (Left), pDsRed-Mito (Right), pECFP-Mito (Left),
(Left) constructs were treated for 5 min with 30 ?M DHR, converted into fluo-
visualized by 2P-CLSM. The various subcellular compartments are shown in red,
Insets show images (?20 magnification) of the areas with ?OH generation
(arrows). Dimensions of the x, y, and z axes are given in ?m.
location of HIF-1?. (A) HepG2 cells transfected with ECFP-ER or ECFP-Mito
constructs were cultured under hypoxia for 60 min and treated for 5 min with
again by 2P-CLSM. Insets show ?OH generation mediated by photoreduction.
fluorescence, indicating subcellular compartments, is shown in red. (B) For de-
tection of endogenous HIF-1? in pECFP-ER transfected HepG2 cells, cells were
exposed to normoxia (60 min) or hypoxia (60 min) and immunostained with an
anti-HIF-1? Ab and a Living Colors Av peptide Ab. HIF-1? (yellow-orange) and
ER (red) are shown. Dimensions of the x, y, and z axes are given in ?m.
Hypoxia-mediated inhibition of ?OH generation at the ER and trans-
www.pnas.org?cgi?doi?10.1073?pnas.0400265101Liu et al.
DHR induced LUC activity 2-fold. DHR treatment under hypoxia
induced LUC activity 4-fold. Treatment with RH 123, the final
product of DHR conversion, had no effect on LUC activity,
indicating the specificity of the DHR effect (Fig. 4).
Induction of HIF-1? Protein and Nuclear Translocation of HIF-1? by
DHR. It was next examined whether the ?OH scavenger DHR
could induce HIF-1? protein levels. Hypoxia induced HIF-1?
protein levels in HepG2 cells. Treatment with DHR under
normoxia also increased HIF-1?, whereas RH 123 had no effect
on HIF-1? protein levels (Fig. 5).
To investigate whether the mimicry of hypoxia by DHR influ-
ences HIF-1? nuclear translocation, cells were transfected with an
expression vector for GFP-tagged HIF-1?. Nuclear accumulation
of GFP-HIF-1? was observed after treatment with either hypoxia
or DHR. In addition, PDTC, which can act as radical scavenger,
also induced nuclear accumulation of HIF-1? (Fig. 6).
Induction of HIF-1? Transactivation by DHR. There are two TADs
present in HIF-1?, TADN (N-terminal; amino acids 531–575)
and TADC (C-terminal; amino acids 786–826). To investigate
fected with pG5-E1B-LUC containing five Gal4-responsive ele-
ments and constructs expressing fusion proteins of the Gal4-DNA
binding domain (Gal4) and either TADN or TADC.
vector neither hypoxia, DHR, nor RH 123 enhanced LUC activity.
Cotransfection of pG5-E1B-LUC and Gal4-HIF1?-TADN en-
hanced LUC activity 200-fold (Fig. 7A). Hypoxia increased LUC
activity in Gal4-HIF1?-TADN cotransfected cells. Exposure to
DHR mimicked hypoxia, enhancing LUC activity 350-fold under
normoxia, whereas RH 123 had no effect (Fig. 7A).
Cotransfection of pG5-E1B-LUC and Gal4-HIF1?-TADNM
encoding a fusion protein with a hydroxylation-resistant muta-
tion (P564A), which thus inhibits binding of VHL and subse-
quent degradation, led to an enhancement of LUC activity and
a loss in the induction by hypoxia as well as by DHR (Fig. 7A).
PAI-1 promoter and EPO-HRE luciferase gene constructs. Cells were trans-
PAI-1 promoter construct mutated at the HRE-2 site (pGL3PAI766-M2), or a
luciferase construct containing three EPO-HREs in front of the SV40 promoter
24 h under normoxia (16% O2) or hypoxia (8% O2). In each experiment, the
was set at 100%, respectively. In pGL3PAI766-M2, the wild-type sequence is
shown on the upper strand, and mutated bases are given in lowercase letters.
t test for paired values was performed to determine significant difference.*,
16% O2vs. 8% O2.**, 16% O2vs. 16% O2? DHR, or 16% O2vs. 8% O2? DHR;
P ? 0.05.
Mimicry of hypoxia by DHR in cells transfected with HIF-1-dependent
either 6 ?M DHR or 6 ?M RH and incubated for 4 h under normoxia (16% O2)
or hypoxia (8% O2). (A) The HIF-1? protein level under normoxia was set at
100%. Values are given as mean ? SEM of three independent experiments.
Student’s t test for paired values was performed to determine significant
difference.*, HIF-1? 16% O2vs. HIF-1? 8% O2.**, HIF-1? 16% O2vs. HIF-1?
of protein to Western blot analysis with an Ab directed against HIF-1? or GM.
Induction of HIF-1? protein levels by DHR. Cells were treated with
Left shows PAI-1 and HO-1 mRNA, as measured by Northern blotting. The RNA
level under normoxia was set at 100%. Right shows the PAI-1 and HO-1 protein
levels, as measured by Western blotting. The protein level under normoxia was
Student’s t test for paired values was performed to determine significant differ-
ence.*, 16% O2 vs. 8% O2.**, 16% O2 vs. 16% O2 ? DHR; P ? 0.05. (B)
20 ?g of RNA was hybridized to PAI-1, HO-1, and ?-actin antisense RNA probes.
the whole-cell extract with an Ab directed against PAI-1, HO-1, and GM.
Liu et al.PNAS ?
March 23, 2004 ?
vol. 101 ?
no. 12 ?
HIF-1?–VHL interaction because of reduction of prolyl hydrox-
ylase activity by Fe redox chemistry. To investigate this possi-
bility, VHL pull-down assays were performed. Binding of radio-
active VHL to GST-TADN was reduced when extracts from
DHR-treated cells were used to hydroxylate the GST-TADN
protein. By contrast, use of cell extracts from RH 123-treated
cells did not disturb HIF-1?–VHL interaction (Fig. 7B).
Furthermore, the total prolyl hydroxylase activity present in the
cell extracts was measured by the formation of succinate from
2-oxoglutarate. It was found that addition of DHR to the assay had
an inhibitory effect on prolyl hydroxylase activity, whereas RH 123
did not influence prolyl hydroxylase activity (Fig. 7C).
Moreover, when Gal4-HIF1?-TADC was cotransfected with
pG5-E1B-LUC, the transactivation was lower than transactivation
with the TADN. Again, however, hypoxia and DHR enhanced
LUC activity, whereas RH 123 had no effect (Fig. 7A). The
cotransfection of Gal4-HIF1?-TADC, in which the critical site for
redox modification cysteine 800 was rendered nonfunctional, dis-
played a loss in induction of LUC activity by hypoxia as well as by
In this study, the usage of gene constructs allowing the cell
compartment-specific expression of the fluorescent proteins
ECFP or DsRed together with the ?OH scavenger DHR enabled
the in vivo localization of an H2O2-using Fenton reaction within
the ER. Scavenging of ?OH generation at the ER by DHR
interfered with O2signaling, leading to an enhanced activation
of HIF-1, HIF-1-dependent PAI-1, and HO-1 gene expression.
ROS as Messengers of the O2Signal: Localization and Kinetics. The
present study localized the Fenton reaction at the ER but not at
mitochondria or other intracellular compartments. This finding
is in line with a study showing a perinuclear Fenton reaction, as
ER (16). To localize the Fenton reaction, we used 2P-CLSM,
which is nonphototoxic, because of infrared light (23). Further-
more, wavelength tunability enabled optimal fluorescence exci-
tation without crosstalk of fluorescence, and maintenance of
physiological conditions in our microscope culture minimized
cell stress. The finding that DHR was fully convertible to RH 123
(which is irreversible) only under normoxia in combination with
5-s blue-light illumination renders short-term blue-light illumi-
nation an ideal proof of intracellular ROS turnover (Fig. 2A).
Under these conditions, hypoxia was accompanied by a decrease
in ROS levels, in agreement with earlier studies (24). However,
other studies (3, 25) have described enhanced ROS under
hypoxia and showed reversibility of normally irreversible ROS-
induced dye oxidation, suggesting that reversibility of the signal
was because of fluorescence-intensity changes of the oxidized
dye. Thus, the present study underlines the importance of ROS
as possible second messengers for O2signaling.
Redox State and Hypoxia-Regulated Gene Expression. Our study
place at the ER but not within mitochondria. This finding is
supported by the fact that, under physiological conditions, mito-
fected with GFP-HIF-1? and cultured under normoxia (16% O2). After 48 h,
?M PDTC under normoxia. (A) Green fluorescence indicates the accumulation
of GFP-HIF-1?. (B) Microscopic images of the same fields as shown in A; the
4?,6-diamidino-2-phenylindole (DAPI) fluorescence indicates the nucleus.
Induction of HIF-1? nuclear translocation by DHR. Cells were trans-
hydroxylase activity. (A) Cells were cotransfected with a luciferase reporter pG5-
E1B-LUC and various fusion gene constructs in which the Gal4 DNA binding
are shown in italics. After 24 h, transfected cells were treated with either 6 ?M
(8% O2). Values are given as mean ? SEM of four independent experiments.
Student’s t test for paired values was performed to determine significant differ-
ence.*, 16% O2(Control) vs. 8% O2.**, Control vs. DHR; P ? 0.05. (B) VHL
pull-down assay. The GST-TADN-HIF-1? protein was incubated either without
visualized. The input remains from directly loaded [35S]VHL. The two bands
represent the 213- and 160-aa VHL products. (C) Dose-dependent inhibition of
HIF prolyl hydroxylase activity by DHR. The GST-TADN-HIF-1? protein or the GST
protein were incubated with HepG2 cell extract, cofactors, and 5-[14C]2-
dependent activity, which was set at 100%, was determined by subtracting the
culture experiments, with each point measured in duplicate.
Induction of HIF-1? transactivation by DHR by inhibition of prolyl
www.pnas.org?cgi?doi?10.1073?pnas.0400265101 Liu et al.
chondria work at a membrane potential of about ?120 mV. Thus,
ROS generation in mitochondria can be measured only when this
potential decreases below ?120 mV, which occurs under stress-
induced increases of calcium ions or decreases of the ATP?ADP
ratio (26). Thus, this study also suggests that an operating respira-
tory chain is not involved in ROS generation (26). Further, the
relevance of the mitochondrial respiratory chain as oxygen sensor
was questioned in experiments with various cell lines defective in
components of the electron-transport chain. These studies showed
no difference in wild-type cells (27–29).
The H2O2levels are controlled mainly by glutathione perox-
idase in cytosol and mitochondria or by catalase in peroxisomes.
Because glutathione peroxidase (KM ?100 ?M) and catalase
(KM? 100 mM) (30) require relatively high H2O2concentra-
tions, it seems conceivable that lower concentrations may be
converted nonenzymatically in an ER-localized Fenton reaction.
The resulting ?OH can then directly or indirectly modify tran-
scription factors, such as HIF-1? (8). This proposal is in line with
this study and investigations showing that H2O2 destabilized
HIF-1? and HIF-2? (EPAS-1) (31, 32). Moreover, our study
shows a compartmentalization of the redox-dependent regula-
tion of HIF-1? because ?OH and ER fluorescence coincide with
HIF-1? under normoxia (Fig. 2B). Furthermore, scavenging of
as HIF-1 activity, which subsequently enhanced PAI-1 and HO-1
gene expression (Figs. 3 and 4). Thereby, it remains uncertain why
DHR and hypoxia were not additive in PAI-1 promoter regulation,
but this phenomenon may depend on a specific PAI-1 promoter
The localization of a substantial amount of HIF-1? to ER
under normoxia does not seem to be caused by an ER retention
sequence but by the interaction of HIF-1? with VHL, which is
retained at the ER by a 64-aa region (33). Because interaction with
HIF-1?, which did not colocalize with ER under normoxia, rep-
resents a part already associated with proteasomes.
The scavenging of ?OH by DHR acted mainly on TADN and
TADC (Fig. 7A). These domains appear to be affected mainly by
a new family of prolyl (34–37) and asparagyl (38) hydroxylases.
Especially for HIF-1?, hydroxylation of Pro-564 under normoxia
would enable VHL binding, which then targets HIF-1? for
proteasomal degradation (17). However, the activity of prolyl
hydroxylase depends on Fe2?, thus pointing to the modulation of
its activity by a redox cycle. This proposal was supported by the
Fe2?-dependent Fenton reaction inhibited HIF-1? prolyl hydrox-
ylase activity and binding of VHL to TADN (Fig. 7 B and C).
The HIF-1?-TADC appeared to be critically regulated by
recruitment of coactivators, such as CBP?p300 and SRC-1 (16,
39). Thereby, interaction of CBP?p300 and SRC-1 with TADC
depended on the redox state of cysteine 800, which can be
modified by Ref-1 (16, 31, 39). Because mutation of C-800
abolished DHR-mediated TADC activation, DHR seems to
affect the redox status of Cys-800, and thus the recruitment of
CBP?p300, directly. This mechanism may act together with the
factor-inhibiting HIF (FIH-1), which has been found to hydroxy-
late Asn-803 of HIF-1?-TADC under normoxia, thus inhibiting
binding of CBP?p300 (40). Similarly, the modification of certain
transcription factors, such as c-Jun or NF-?B (1).
In summary, the present study has shown that generation of
?OH in a Fenton reaction at the ER contributes to HIF-1?
regulation by either modulation of prolyl hydroxylase activity or
interference with redox-sensitive residues, such as Cys-800,
within the TADC. Thus, under normal physiological conditions,
the changes of the redox status within a cell may contribute to
an efficient and fast-responding O2-sensing system.
We thank Dr. Patrick Maxwell for the kind gift of the pCMV-HA-VHL
construct. This study was supported by Deutsche Forschungsgemein-
schaft Grants SFB 402, Teilprojekt A1, and GRK 335 (to T.K.);
Bundesministerium fu ¨r Bildung, Wissenschaft, Forschung und Tech-
nologie Grant 13N7447?5 (to H.A.); and grants from Silicon Graphics,
Nikon, and Newport (to H.A.).
1. Bunn, H. F. & Poyton, R. O. (1996) Physiol. Rev. 76, 839–885.
2. Gorlach, A., Holtermann, G., Jelkmann, W., Hancock, J. T., Jones, S. A., Jones,
O. T. & Acker, H. (1993) Biochem. J. 290, 771–776.
3. Chandel, N. S., McClintock, D. S., Feliciano, C. E., Wood, T. M., Melendez, J. A.,
Rodriguez, A. M. & Schumacker, P. T. (2000) J. Biol. Chem. 275, 25130–25138.
4. Agani, F. H., Pichiule, P., Chavez, J. C. & LaManna, J. C. (2000) J. Biol. Chem.
5. Wenger, R. H. (2002) FASEB J 16, 1151–1162.
6. Hildebrandt, W., Alexander, S., Bartsch, P. & Droge, W. (2002) Blood 99, 1552–1555.
7. Kietzmann, T., Fandrey, J. & Acker, H. (2000) News Physiol. Sci. 15, 202–208.
8. Kietzmann, T., Porwol, T., Zierold, K., Jungermann, K. & Acker, H. (1998)
Biochem. J. 335, 425–432.
9. Semenza, G. L. (2001) Pediatr. Res. 49, 614–617.
10. Wenger, R. H. & Gassmann, M. (1997) Biol. Chem. 378, 609–616.
11. Semenza, G. L. (1998) Curr. Opin. Genet. Dev. 8, 588–594.
12. Kietzmann, T., Cornesse, Y., Brechtel, K., Modaressi, S. & Jungermann, K.
(2001) Biochem. J. 354, 531–537.
13. Kietzmann, T., Roth, U. & Jungermann, K. (1999) Blood 94, 4177–4185.
14. Kallio, P. J., Okamoto, K., O’Brien, S., Carrero, P., Makino, Y., Tanaka, H. &
Poellinger, L. (1998) EMBO J. 17, 6573–6586.
15. Kruger, M., Schwaninger, M., Blume, R., Oetjen, E. & Knepel, W. (1997)
Naunyn Schmiedeberg’s Arch. Pharmacol. 356, 433–440.
16. Carrero, P., Okamoto, K., Coumailleau, P., O’Brien, S., Tanaka, H. &
Poellinger, L. (2000) Mol. Cell. Biol. 20, 402–415.
18. Baader, E., Tschank, G., Baringhaus, K. H., Burghard, H. & Gunzler, V. (1994)
Biochem. J. 300, 525–530.
20. Immenschuh, S., Hinke, V., Ohlmann, A., Gifhorn-Katz, S., Katz, N., Junger-
mann, K. & Kietzmann, T. (1998) Biochem. J. 334, 141–146.
21. Ehleben, W., Porwol, T., Fandrey, J., Kummer, W. & Acker, H. (1997) Kidney
Int. 51, 483–491.
22. Strohmaier, A. R., Porwol, T., Acker, H. & Spiess, E. (2000) Cells Tissues
Organs 167, 1–8.
23. Hockberger, P. E., Skimina, T. A., Centonze, V. E., Lavin, C., Chu, S., Dadras,
S., Reddy, J. K. & White, J. G. (1999) Proc. Natl. Acad. Sci. USA 96, 6255–6260.
24. Fandrey, J. & Genius, J. (2000) Adv. Exp. Med. Biol. 475, 153–159.
25. Chandel, N. S., Maltepe, E., Goldwasser, E., Mathieu, C. E., Simon, M. C. &
Schumacker, P. T. (1998) Proc. Natl. Acad. Sci. USA 95, 11715–11720.
27. Srinivas, V., Leshchinsky, I., Sang, N., King, M. P., Minchenko, A. & Caro, J.
(2001) J. Biol. Chem. 276, 21995–21998.
29. Searle, G. J., Hartness, M. E., Hoareau, R., Peers, C. & Kemp, P. J. (2002)
Biochem. Biophys. Res. Commun. 291, 332–337.
30. Chance, B., Sies, H. & Boveris, A. (1979) Physiol. Rev. 59, 527–605.
31. Huang, L. E., Arany, Z., Livingston, D. M. & Bunn, H. F. (1996) J. Biol. Chem.
32. Wiesener, M. S., Turley, H., Allen, W. E., Willam, C., Eckardt, K. U., Talks, K. L.,
33. Schoenfeld, A. R., Davidowitz, E. J. & Burk, R. D. (2001) Int. J. Cancer 91, 457–467.
34. Jaakkola, P., Mole, D. R., Tian, Y. M., Wilson, M. I., Gielbert, J., Gaskell, S. J.,
Kriegsheim, A., Hebestreit, H. F., Mukherji, M., Schofield, C. J., et al. (2001)
Science 292, 468–472.
35. Ivan, M., Kondo, K., Yang, H., Kim, W., Valiando, J., Ohh, M., Salic, A., Asara,
J. M., Lane, W. S. & Kaelin. (2001) Science 292, 464–468.
36. Epstein, A. R., Gleadle, J. M., McNeill, L. A., Hewitson, K. S., O’Rourke, J.,
Mole, D. R., Mukherji, M., Metzen, E., Wilson, M. I., Dhanda, A., et al. (2001)
Cell 107, 43–54.
37. Oehme, F., Ellinghaus, P., Kolkhof, P., Smith, T., Ramakrishnan, S., Hutter, J.,
Schramm, M. & Flamme, I. (2002) Biochem. Biophys. Res. Commun. 296, 343.
38. Lando, D., Peet, DJ., Whelan, DA., Gorman, J. & Whitelaw, M. L. (2002)
Science 295, 858–861.
39. Ema, M., Hirota, K., Mimura, J., Abe, H., Yodoi, J., Sogawa, K., Poellinger,
L. & Fujii-Kuriyama, Y. (1999) EMBO J. 18, 1905–1914.
40. Lando, D., Peet, D. J., Gorman, J. J., Whelan, D. A., Whitelaw, M. L. & Bruick,
R. K. (2002) Genes Dev. 16, 1466–1471.
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vol. 101 ?
no. 12 ?