Differential activation of catalase expression and activity by PPAR
agonists: Implications for astrocyte protection in anti-glioma therapy$
Nicholas K.H. Khooa,d,n,1, Sachin Hebbarc,1, Weiling Zhaoc, Steven A. Mooreb,
Frederick E. Domanna, Mike E. Robbinsc
aDepartment of Radiation Oncology, Free Radical and Radiation Biology, Holden Comprehensive Cancer Center, The University of Iowa, Iowa City, IA 52242,
bDepartment of Pathology, Holden Comprehensive Cancer Center, The University of Iowa, Iowa City, IA 52242, USA
cDepartment of Radiation Oncology, Comprehensive Cancer Center of Wake Forest University, Wake Forest University School of Medicine, Winston-Salem,
NC 27157, USA
dDepartment of Pharmacology and Chemical Biology, University of Pittsburgh, Pittsburgh, Pennsylvania 15261, USA
a r t i c l e i n f o
Received 7 December 2012
Received in revised form
20 December 2012
Accepted 21 December 2012
Primary rat astrocytes
C6 glioma cells
a b s t r a c t
Glioma survival is dismal, in part, due to an imbalance in antioxidant expression and activity. Peroxisome
proliferator-activated receptor (PPAR) agonists have antineoplastic properties which present new redox-
dependent targets for glioma anticancer therapies. Herein, we demonstrate that treatment of primary
cultures of normal rat astrocytes with PPAR agonists increased the expression of catalase mRNA protein, and
enzymatic activity. In contrast, these same agonists had no effect on catalase expression and activity in
malignant rat glioma cells. The increase in steady-state catalase mRNA observed in normal rat astrocytes was
due, in part, to de novo mRNA synthesis as opposed to increased catalase mRNA stability. Moreover,
pioglitazone-mediated induction of catalase activity in normal rat astrocytes was completely blocked by
transfection with a PPARg-dominant negative plasmid. These data suggest that defects in PPAR-mediated
signaling and gene expression may represent a block to normal catalase expression and induction in
malignant glioma. The ability of PPAR agonists to differentially increase catalase expression and activity in
normal astrocytes but not glioma cells suggests that these compounds might represent novel adjuvant
therapeutic agents for the treatment of gliomas.
& 2013 The Authors. Published by Elsevier B.V. All rights reserved.
Novel therapeutic approaches that selectively protect normal
brain cells and/or sensitize glioma cells to anti-cancer therapies
are urgently needed. Roughly 70,000 new cases of primary brain
tumors will be diagnosed in the US, the majority being grade
4 astrocytoma (glioblastoma) . Despite multimodality therapy
including surgery, radiation therapy and chemotherapy with
temozolomide, median survival remains only approximately 15
months  and the 5-year survival rate is 1% . This outcome
reflects both glioma cell resistance to therapy and the risk of
radiation-induced normal brain injury, which limits the total dose
that can be safely administered to the tumor . Recent data
suggest that progressive cognitive impairment occurs in ?50% of
brain tumor patients who are long-term survivors after treatment
with partial or whole-brain irradiation [5,6]. This negative prog-
nosis is, in part, due to radiation-induced oxidative tissue injury.
The central nervous system (CNS) is inherently susceptible to
oxidative stress. This is evidenced by the CNS being: (1) highly
active in oxidative metabolism, leading to a relatively high rate of
reactive oxygen species (ROS) production ; (2) relatively low in
the specific activity of the key antioxidant enzymes superoxide
dismutase (SOD), catalase and glutathione peroxidase (GPx)  in
oligodendrocytes, neurons and endothelial cells [9,10]; and
(3) rich in readily oxidizable polyunsaturated fatty acids (PUFAs)
such as docosahexaenoic acid and eicosapentaenoic acid .
These issues suggest that a lipid signaling-based therapeutic
strategy may be beneficial in treating CNS cancers. PUFAs and
oxidized lipids are ligands for peroxisome proliferator-activated
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2213-2317/$-see front matter & 2013 The Authors. Published by Elsevier B.V. All rights reserved.
$This is an open-access article distributed under the terms of the Creative
Commons Attribution License, which permits unrestricted use, distribution, and
reproduction in any medium, provided the original author and source are credited.
Abbreviations: 9cRA, 9 cis retinoic acid; ActD, actinomycin D; BCNU, 1,3-bis
(2-chloroethyl)-1-nitrosourea; CNS, central nervous system; DCF, dichlorofluor-
escein; GPx, glutathione peroxidase; GSH, glutathione; Pio, Pioglitazone; PPAR,
Peroxisome proliferator-activated receptor; PPARg-d.n, PPARg-dominant nega-
tive; PPRE, PPAR-response elements; PUFAs, polyunsaturated fatty acids; ROS,
reactive oxygen species; Rosi, Rosiglitazone; RXR, retinoid X receptor; shRNA,
short hairpin RNA; SOD, superoxide dismutase; TZDs, thiazolidinediones
nCorresponding author at: Department of Pharmacology and Chemical Biology,
E1314 BST, University of Pittsburgh School of Medicine, Pittsburgh, PA 15261, USA.
Tel.: þ1 412 648 9671; fax: þ1 412 648 2229.
E-mail addresses: email@example.com (N.K.H. Khoo),
firstname.lastname@example.org (S. Hebbar), email@example.com (W. Zhao),
firstname.lastname@example.org (S.A. Moore),
email@example.com (F.E. Domann).
Redox Biology 1 (2013) 70–79
receptors (PPARs) [12,13], which are members of the nuclear
hormone receptor (NHR) superfamily. PPARs are ligand-activated
transcription factors that heterodimerize with the retinoid X
receptor (RXR), bind to PPAR-response elements (PPRE), and
thereby regulate gene expression . Three isotypes of PPARs
have been identified and designated a, b/d, and g . PPARa is
activated by fibrates, a class of cholesterol-lowering drugs used
in the treatment of dyslipidemia, and functions by enhancing
b-oxidation in the liver . PPARg agonists, such as thiazolidi-
nediones (TZDs), are best known for controlling hyperglycemia
and treating type 2 diabetes mellitus, in part, through the
regulation of adipocyte differentiation and lipid storage/metabo-
lism . PPARb/d is expressed in tissues controlling lipid
metabolism and acts as a lipid sensor, thus representing a
molecular target for treating metabolic syndrome . Data
supports the concept that PPARg mediates anti-proliferative and
pro-apoptotic activities in tumor cells .
Studies have shown that activation of PPARg has antineoplas-
tic effects through redox-dependent mechanisms. The PPARg
agonists Rosiglitazone (Rosi) and Pioglitazone (Pio) decreased cell
viability in both human (U87 and A172) and rat (C6) malignant
glioma cells whereas normal rat astrocytes were not affected
[19,20]. This cytotoxic effect was spared by N-acetylcysteine
suggesting reactive oxygen species (ROS) are playing an integral
role. PPARg agonists decreased glutathione (GSH) levels in C6 rat
glioma cells and increased dichlorofluorescein (DCF) fluorescence
in both the primary rat astrocytes and C6 rat glioma cells .
Moreover, Pio treatment significantly reduced tumor volume in a
rat glioma model by decreasing proliferation, suppressing MMP9
induction, and inhibiting tumor cell invasion .
The cytotoxic effect of PPARg agonists on glioma cells is
partially mediated by enhanced redox reactions. Yet, normal rat
astrocytes, which are the most prevalent cell type in the CNS,
comprising more than 50% of brain volume and outnumbering
neurons approximately 9:1 , appear to be resistant to PPARg-
mediated cytotoxicity . Catalase gene expression and enzy-
matic activity was the only antioxidant enzyme significantly
increased by PUFAs . In light of this, it is hypothesized that
catalase is responsible for the protective effects following PPAR
agonist treatment of primary cultures of normal rat astrocytes. To
investigate this putative role of PPARs in catalase regulation, we
treated primary cultures of normal rat astrocytes and C6 rat
glioma cells with PPAR agonists. We report herein that PPAR
agonists increased catalase gene expression and enzymatic activ-
ity in primary cultures of normal rat astrocytes. In contrast, PPAR
agonists failed to increase catalase expression in glioma cells. To
gain insight into a potential mechanism, Cos-1 cells were trans-
fected with a PPARg-dominant negative (PPARg-d.n.) plasmid,
revealing that PPARg plays a significant role in the regulation of
catalase expression. The ability of PPAR agonists to selectively
upregulate catalase expression in normal astrocytes but not
glioma cells suggests that these compounds might represent
novel adjuvant therapeutic agents for the treatment of gliomas.
Tissue culture materials were obtained from the following man-
ufacturers: Dulbecco’s Modified Eagle’s Medium (DMEM/F12), Mini-
streptomycin (Gibco, Grand Island, NY); fetal bovine serum (Hyclone;
Logan, Utah); L-glutamine and gentamycin (Amersham; Arlington
Heights, IL). Pioglitazone, Troglitazone, WY-14,643, and 9 cis Retinoic
Acid (9cRA) were purchased from BioMol (Plymouth Meeting, PA);
Rosiglitazone was purchased from Cayman Chemical (Ann Arbor, MI).
The PPARg agonists CP-086,325-02 (CP086; Darglitazone) and CP-
096,125 (CP096) were kindly donated by Michael Gibbs at Pfizer
(Groton, CT). The PPARa specific agonist GW3276 and GR-259662
were kind gifts from Tim Willson at Glaxo Wellcome (Research
Triangle Park, NC).
Primary rat astrocytes were isolated from 1 to 2 day old
Sprague-Dawley rat pups . The cells were cultured in MEM
containing 10% FBS, 2 mM L-glutamine, 6 g/L glucose (33 mM)
and 50 mg/mL gentamycin. The cultures were maintained at 37 1C
in a humidified atmosphere of 5% CO2. Astrocytes were purified
from other glial cells by shaking the flasks before each media
change. The media was changed every fourth day and the cells
were used within a week of reaching confluence. Immunofluor-
escence staining with GFAP was carried out to characterize the
cell purity (498% astrocytes).
The rat malignant glioma cell line C6 was grown in DMEM/F12
containing 10% FBS, 50 IU/mL penicillin, and 50 IU/mL streptomycin.
Cells were maintained at 37 1C as a monolayer in 75 cm2tissue
culture flasks in a humidified atmosphere containing 5% CO2. Stock
cultures were passaged twice weekly by trypsinization. For experi-
ments, the cultures were between passage 12 and 20, and the
incubations started when the cells were roughly 40% confluent.
Cloning of promoter deletion constructs
Total genomic DNA from primary rat astrocytes or rat brain
microvessel endothelial cells (RBMECs) was isolated as previously
reported . Rat catalase promoter deletion constructs were
generated by PCR using the following primers: -1046, 50-ACAGCC-
(3641–3660); -938, 50-ATTGATTAAAAT-
GAAAAATAAGCGAC-30(3751–3776); and -207, 50-CTCCTTCCAA
TCCTGTCCC-30(4481–4499). The number represents the location
of the rat catalase promoter construct with respect to the
translational start site. The common downstream primer, 50-
CAGATGAAGCAGTGGAAGGA-30(4719–4738), was used with all
the primers listed above. The numbers inside the parentheses
indicate the location of primers with reference to the published
sequence beginning with the first nucleotide of exon 1 [GenBank
accession # M25669].
PCR was performed using Taq DNA polymerase (Perkin-Elmer;
Emeryville CA). The PCR conditions consisted of 94 1C for 4 min,
followed by 35 cycles of 94 1C for 1 min, 58 1C for 50 s, and 72 1C
for 1.5 min, followed by a final extension at 72 1C for 5 min. The
expected sizes of the PCR products were verified by 1 kb and
100 bp DNA markers (Promega; Madison, WI) on a 1% TAE gel.
The PCR products were ligated into the pCR2.1 TA-TOPO cloning
vector and transformed into competent cells as described by the
manufacturer (Invitrogen; Carlsbad, CA). DNA was isolated using
the gel extraction kit (Qiagen), digested with EcoRI (Holden
Cancer Center; University of Iowa) and the identity was verified
by sequencing (DNA Core Facility; University of Iowa). The
constructs were engineered by subcloning the promoter deletion
fragments from pCR2.1 (Invitrogen) into a luciferase-based plas-
mid pGL3-basic (Promega). The constructs were verified by
sequencing (DNA Core Facility; The University of Iowa).
The tandem tripeat of the catalase PPRE (DR1x3) was synthe-
sized using an oligonucleotide consisting of TAATCAAGGT-
GAAAGTTGAGAAG with KpnI and XhoI restriction sites on its 50-
and 30-ends, respectively. The oligonucleotide was restricted with
KpnI and XhoI, gel isolated, and cloned into pGL3-basic. The
DR1x3 was verified by sequencing.
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
The full length RXRa was synthesized as previously reported .
PPARa expression vector was synthesized from mouse liver RNA by
PCR using the following primers: 50-GTGGCTGGTCAAGTTCGG-30
(upstream) and 50-CTCGGAGGTCCCTGAACAG-30(downstream) for
PPARa, [GenBank Acc# X57638]. Mouse liver was dounce homo-
genized and total RNA was isolated. Total RNA was reversed
transcribed using the Superscript II Reverse Transcription kit (Roche)
according to the manufacturer’s specifications. The cDNA generated
was PCR amplified under the conditions of 94 1C for 4 min, followed
by 35 cycles of 94 1C for 1 min, 62.5 1C for 1 min, and 72 1C for
1.5 min, followed by a final extension at 72 1C for 5 min. The expected
size of the PCR product was verified on a 1% low melting point TAE
gel. A single band of the expected size of the PPARa PCR product was
isolated using the gel extraction kit (Qiagen) and cloned into the
pTargeT mammalian expression vector (Promega) according to the
manufacturer’s directions. The PPARa expression vector was verified
Transfection and reporter luciferase assay
C6 rat glioma and Cos-1 cells were transiently transfected
using SuperFect Reagent (Qiagen; Valencia, CA) according to the
manufacturer’s directions. Primary rat astrocytes were transiently
transfected with Effectene (Qiagen). Cells were transfected with
0.25–1 mg of rat catalase reporter deletion constructs or the
tandem tripeat of the catalase PPRE (DR1x3) and co-transfected
with a plasmid containing 0.5–3 mg CMV SPORT-b-galactosidase
(Invitrogen) for 18–24 h depending on cell type. In a subset of
experiments, cells were additionally transiently transfected with
0.1–1 mg of pTargeT, mRXRa, mPPARg2, and mPPARa expression
vectors. Following transfection, cells were rinsed and treated with
PPAR and/or RXR agonists for 24 h. The cells were then rinsed
with phosphate buffered saline (PBS), lysed with Passive Lysis
Buffer (Promega), and luciferase activity was measured using
Luciferase Assay System (Promega) according to manufacturer’s
instructions. To control for transfection efficiency, b-galactosidase
activity was measured at 420 nm using o-nitrophenyl b-d-galac-
topyranoside (Sigma; St. Louis, MO) as substrate.
Protein sample preparation
Normal rat astrocyte and C6 glioma cell cultures were incu-
bated in the presence of the various PPAR and RXR agonists for
48 h and were confluent at the time of harvest. Cells were rinsed
in PBS twice and harvested by scraping and centrifugation. The
cell pellets were then stored at ?20 1C. Upon use, the cell pellets
were resuspended in 50 mM potassium phosphate buffer (pH 7.8)
and then sonicated on ice with four bursts of 20 seconds each
using a Vibra Cell sonicator (Sonics and Materials, Inc.) with a cup
horn at full power. Total protein concentrations were determined
by the Bradford method  using BSA as a standard. For nuclear
protein extraction, cells were rinsed in PBS and nuclear protein
was then isolated as described previously .
Catalase activity assay
Catalase enzymatic activity was determined by a modified
version of the method described by Beers and Sizer . Briefly,
300 mg of primary rat astrocyte and C6 glioma extract were added
to 30 mM H2O2in 50 mM of potassium phosphate buffer (pH 7.8),
and the consumption of H2O2was measured at 240 nm for 120 s
at 15 s intervals. Catalase activity was expressed in K units per g
protein per second (K/g/s).
Western blot analysis
Western blot analysis was performed using anti-IgG catalase
antibody (Athens BioTech.; Athens, GA), anti-PPAR a antibodies
(Santa Cruz, Santa Cruz, CA), anti-PPAR g antibodies (Wak-
Chemie; Sulzbacherstasse, Germany and Santa Cruz), anti-RXRa
antibody and anti-Actin antibody (Santa Cruz). Briefly, 10–20 mg
of whole cell lysate or nuclear protein were separated by poly-
acrylamide gel electrophoresis (PAGE) on a 10–12% gel by a
modified version described previously . Proteins were trans-
ferred to a nitrocellulose membrane (Schleicher and Schuell;
Keene, NH) for 1 h at 100 V and then blocked with 5% dry milk
in TBST (0.02 M Tris/0.15 M NaCl buffer, pH 7.45 and 0.05% Tween
20). The gel was stained by Gel Code (Pierce Chem.; Rockford, IL)
as a control for loading variation. Membranes were incubated
with catalase antibody (1:1000 dilution), PPARa antibodies
(1:500 dilution), PPARg antibodies (1:500 dilution) or Actin anti-
body (1:400) for 2–3 h, rinsed with TBST and then incubated with
a horseradish peroxidase-conjugated secondary antibody (Sigma;
St. Louis, MO) for 1 h. Bands were visualized by ECL chemilumi-
nescence (Amersham) and exposed to film.
Generation of a rat catalase partial cDNA probe
Total mRNA was extracted according to the directions pro-
vided by the manufacturer (Tel-Test B Inc.; Friendswood, TX).
Briefly, the cells were homogenized into RNA-STAT-60 and then
the total RNA was extracted into chloroform. The RNA was
removed, placed into a fresh tube, and allowed to precipitate
with isopropanol. Following the precipitation, the total RNA was
washed in 75% ethanol and quantified using UV spectroscopy.
Total RNA from primary rat astrocytes was used to synthesize the
catalase cDNA by RT-PCR using the following primers: 50-
TCTGG-30. These primers were located in exons 2 and 3 [GenBank
Accession #M25670 and M25671, respectively]. The predicted
product was 228 bp in length. The RT-PCR reactions were carried
out using one-step RT-PCR kit (Qiagen; Santa Clarita, CA) in 50 mL
reaction solution containing 1 mg total RNA, 10 mL of 10x PCR
buffer, 2 mL of 10 mM dNTP, 1 mM of sense and antisense primers,
8 units of RNase inhibitor (Promega; Madison, WI), and 2 mL
enzyme mixture. The following PCR conditions were used: 94 1C
for 5 min, then 35 cycles of 94 1C for 1 min, 64 1C for 30 s, and
72 1C for 1 min, followed by a final extension step at 72 1C for
5 min. The 228 bp PCR products of catalase were cloned into the
pCR2.1 TA-TOPO cloning vector and transformed into competent
cells as described by the manufacturer (Invitrogen; Carlsbad, CA,
USA). DNA was isolated (Promega) and sequenced by the DNA
Core at the University of Iowa.
RNA isolation and Northern blot analysis
Total mRNA was extracted as described above. Five to 10 mg of
RNA were resolved by electrophoresis in a 1% agarose gel contain-
ing 2.2% formaldehyde using a running buffer of 20 mM MOPS,
5 mM sodium acetate, and 1 mM EDTA at pH 7. RNA was
transferred to a Nytran membrane using a TurboBlotter transfer
system (Schleicher and Schuell) for 2 h and then UV cross-linked
(Stratagene; LaJolla, CA). The catalase partial cDNA probe
described above was labeled with [a32P]dCTP (NEN Life Science
Products; Boston, MA) using a random primed labeling kit
according to the manufacturer’s instructions (Roche Diagnostics;
Mannheim, Germany). Membranes were prehybridized with Per-
fectHyb Plus Hybridization Buffer (Sigma; St. Louis, MO) for 1 h
and then hybridized with the same buffer for 6–8 h at 68 1C.
Membranes were washed and scanned using a Typhoon 8600
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
PhosphoImager (Molecular Dynamics; Piscataway, NJ) and densi-
tometry was performed using ImageQuant 5.1. Membranes were
stripped and reprobed with a radiolabeled cyclophilin cDNA
probe (Ambion; Austin, TX).
Electrophoretic mobility shift assay (EMSA)
The following top and bottom strand complementary oligonu-
cleotides were hybridized to generate a double stranded DNA
probe of the same sequence as the PPRE found in the rat catalase
GATCCTTCTCAACTTTCACCT TGATTA-30. The oligonucleotide was
fill-in end-labeled with Klenow DNA polymerase, a32P-dCTP, and
dNTPs for 5 min at 37 1C. One to 10 mg of nuclear protein, gel shift
buffer [10 mM Tris (pH 7.5), 4% glycerol, 50 mM NaCl, 1 mM
MgCl2, 0.5 mM EDTA, 0.5 mM dithiothreitol (DTT)], poly dIdC
(Pharmacia, Piscataway, NJ), and the32P labeled probe described
above were incubated at room temperature for 15–30 minutes.
Following the incubation, antibodies were added for 1.5 h. Anti-
bodies were purchased from Santa Cruz Biotechnology, Inc.: anti-
PPARa (H-98X), anti-PPARg (H-100X), anti-RXRa (D-20X) and
normal rabbit IgG (sc2027). The bound DNA complexes with
antibody were separated from free probe by PAGE on a 5% native
gel between 15–20 mA in 1x TBE. Gels were exposed to X-ray film
at ?80 1C.
Data were expressed as the mean7SEM. Both Student’s t-test
and ANOVA were used in the comparison of different groups.
Within ANOVA, the weighted least squares method was used to
adjust for unequal variance among the groupings . Dunnit’s
multiple comparison procedure was performed to determine
PPARa, PPARg and RXR agonists upregulate catalase activity in
normal rat astrocytes but not in rat malignant glioma cells.
Primary rat astrocytes treated with PPARa or PPARg agonists
(10 mM) for 48 h increased catalase activity compared to the
vehicle control (Fig. 1A). The maximal increase in catalase activity
was observed in primary cultures of rat astrocytes treated with
the PPARg agonist CP096 (5.8270.77 k/g/s), a value significantly
greater than that observed in primary cultures of rat astrocytes
treated with DMSO vehicle alone (2.7870.23 k/g/s, p40.0001).
There was no difference between vehicle control (0.02% vol/vol of
DMSO) and untreated confluent primary cultures of rat astro-
cytes. Significant increases in catalase activity were also observed
in astrocytes incubated with PPARg agonists rosiglitazone (Rosi),
troglitazone or RXR agonist (9cRA). Less pronounced increases
were observed following treatment of astrocytes with PPARa ago-
nists GW3276 or WY14643. In contrast, treating C6 glioma cells
with PPARa, PPARg or RXR agonists (10mM) for 48 h failed to
enhance catalase activity in C6 glioma cells (Fig. 1B). Moreover, C6
glioma cells (4.5470.09 k/g/s) expressed higher constitutive levels
of catalase activity as compared to the primary rat astrocytes
(2.7870.23 k/g/s, p¼0.002, t-test).
PPARa, PPARg and RXR agonists upregulate catalase mRNA and
protein expression in primary rat astrocytes but not in C6 glioma cells
Agonist treatment resulted in 1.5- to 2-fold increase in steady-
state catalase mRNA levels compared to the vehicle control (Fig. 2A).
Conversely, C6 glioma cells incubated with the same set of PPAR and
RXR agonists failed to alter catalase mRNA expression (Fig. 2B) over
the same time. There was no difference between untreated cells
compared to DMSO-treated cells for both primary rat astrocytes and
C6 rat glioma cells. PPARa agonist (GW3276) and RXR agonist (9cRA)
increased catalase protein in primary rat astrocytes 1.9- and 1.6-fold,
respectively (Fig. 2C). Additionally, catalase immunoreactive protein
levels were increased 1.4- and 1.5-fold following Rosi and troglita-
zone treatment, respectively (Fig. 2C). In contrast, malignant glioma
cells treated with the same agonists failed to increase catalase protein
levels (Fig. 2D).
PPAR agonist-mediated increase in catalase mRNA is not caused
increased mRNA stability
Changes in steady-state mRNA levels may be attributable to
alterations in the degradation rate of a transcript and/or changes
in the rate of transcription. The relative contribution of a post-
transcriptional mechanism in the PPAR agonist-mediated mod-
ulation of catalase mRNA levels was determined using the
transcriptioninhibitor actinomycinD (ActD). Primaryrat
Catalase Activity (K/g/second)
DMSOCP 096 GW 3276 WY
Fig. 1. PPARa and PPARg agonists increased catalase enzymatic activity in astrocytes
but not in glioma cells. Primary rat astrocytes (A) and C6 glioma cells (B) were
supplemented for 48 h with Rosi, Troglitazone, and CP096 (PPARg), GW3276 and WY-
14,643 (PPARa), 9cRA (RXR) and vehicle (DMSO) at concentration of 10mM. Protein
was then harvested and catalase activity was determined spectrophotometrically at
240 nm. For astrocytes, one-way analysis of variance (ANOVA) was used to compare
the groups. Within ANOVA, the weighted least squares method was used to adjust for
the heteroskedasticity (that is, the unequal variance among the groupings) . The
overall test of differences among the groups was statistically significant (po0.0001 by
ANOVA). Rosi, troglitazone, CP096 and 9cRA were significantly different from the
DMSO group by Dunnett’s multiple comparison procedure as indicated by
glioma cells, the overall test of differences among the groups was not statistically
significant (po0.30 by ANOVA). No groups were significantly different from the DMSO
group at the 0.05 level. There is a significant difference in the mean values for the
outcome variable between the astrocyte control (DMSO) vs C6 control (DMSO)
(p¼0.002) as indicated by #. Results are derived from at least six independent
experiments and data are expressed as mean7SEM.
n. For C6
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
astrocytes were supplemented with 10 mM CP096 (as it showed
a42-fold increase in catalase mRNA), Rosi, or DMSO for 24 h and
then treated with 5 mg/mL ActD. Cells were harvested at 0, 4, 8, 12
and 24 h time-intervals after addition of ActD. The amount of
catalase mRNA was normalized using the expression of cyclophi-
lin (Fig. 3A). The mRNA decay rates were comparable for both the
PPARg agonists and DMSO-treated cells, suggesting that the
PPAR-mediated increase in catalase was independent of transcript
stability (Fig. 3B).
PPAR agonists induce de novo transcription of catalase mRNA
We previously identified a PPRE in the rat catalase promoter
located at nucleotide (nt) ?1027 to ?1015 with respect to the
translation start site . To examine whether PPAR agonists led to
transcriptional transactivation of catalase gene expression, promoter
deletion constructs of the rat catalase promoter were transiently
transfected in to primary rat astrocytes. The promoter deletion
construct containing the PPRE (-1046) increased promoter activity
420-fold compared to empty vector control (Fig. 3C). The deletion of
the PPRE significantly diminished reporter activity following the
transfection of rat catalase deletion constructs (-938 and -207). The
luciferase activity was normalized to b-gal and then all the catalase
promoter deletion constructs were normalized to the activity of the
empty vector control (pGL3-basic). Additionally, transiently trans-
fected primary rat astrocytes had significantly increased promoter
activity with the rat catalase promoter deletion construct containing
the PPRE (-1046) with expression plasmids PPARg2 and RXRa
compared to empty vector (pTargeT) control (Fig. 3D, open bars).
Moreover, the treatment of 5mM Rosi for 24 h further increased
promoter activity compared to DMSO-treated primary rat astrocytes
transfected with PPARg2 and RXRa (Fig. 3D, dark bars). Taken
together, our results support the hypothesis that the PPAR-
mediated increase in steady-state catalase mRNA observed in primary
rat astrocytes is due to de novo catalase transcript rather than
increased catalase mRNA stability.
PPARg regulates catalase expression. To gain better insight into
mechanism, Cos-1 cells were used. Cos-1 cells exhibited low endo-
genous levels of RXRa protein determined by Western blot analysis.
In contrast, PPARa and PPARg protein were not detectable. Transient
transfection of mPPARg2 and RXRa expression vectors into Cos-1
cells resulted in a robust expression of PPARg and RXRa protein
Relative Fold Induction
Relative Fold Induction
Relative Fold Induction
Relative Fold Induction
Fig. 2. PPAR and RXR agonists increase catalase expression in astrocytes but not in glioma cells. Primary rat astrocytes and C6 rat glioma cells were supplemented for 48 h
with the following agonists (10 mM). Representative Northern blots for primary rat astrocytes (A) and C6 rat glioma cells (B) probed for catalase and cyclophillin.
Densitometric analysis was performed using Typhoon 8600 Phosphoimager and software package ImageQuant 5.1 (below blots). Representative Western blot for primary
rat astrocytes (C) and C6 glioma cells (D) probed for catalase. Quantification of catalase protein levels were determined (below). Northern and Western blots are
representatives of at least four independent experiments and data are expressed as mean7SEM. Treatments were significantly different from the DMSO group by
Dunnett’s multiple comparison procedure as indicated byn, po0.01.
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
compared to the empty vector pTargeT control (Supplemental data
S1). To demonstrate binding and formation of the PPAR and RXR
heterodimer to the PPRE, Cos-1 cells were transiently transfected
with PPARg2 and RXRa and nuclear extracts were isolated for EMSAs.
The rat catalase PPRE (AGGTGA-a-AGTTGA) was end-labeled with32P
and incubated with nuclear extracts. The nuclear extracts isolated
from Cos-1 cells transfected with PPARg2 and RXRa exhibited
pronounced binding to the catalase PPRE compared to cells trans-
fected with the pTargeT vector alone (Fig. 4A). Supershift experiments
were performed to confirm specific PPAR binding to the catalase
PPRE. The incubation of anti-PPARg antibody resulted in a dose-
dependent supershift (Fig. 4A, lanes 6 and 9) in the binding of nuclear
extract isolated from Cos-1 cells transfected with PPARg2 and RXRa.
Although, the findings indicate that PPAR/RXRa heterodimers
present in the nuclear extract of transfected Cos-1 cells can bind to
the catalase PPRE, this does not demonstrate that the catalase PPRE is
functional. To confirm the functionality of the PPRE located in the rat
catalase promoter, Cos-1 cells were transfected with the catalase
promoter deletion construct containing the PPRE (-1046), PPARg2,
RXRa or pTargeT (empty vector control for the NHR expression
plasmids) and then incubated with Rosi and 9cRA. PPAR-RXR
heterodimers have been shown to synergistically activate reporter
genes when both receptors are activated by their respective agonists
[29,30]. A similar synergistic increase in reporter activity was
observed in Cos-1 cells transfected with the -1046 rat catalase
promoter deletion construct, PPARg2, RXRa and incubated with Rosi
and 9cRA, or both agonists together showed a 2.3-, 9.5- and 15-fold
increase, respectively (Fig. 4B). Moreover, Cos-1 cells transfected with
PPARg2, RXRa and a catalase promoter deletion construct (-938)
lacking the PPRE failed to increase promoter activity in response to
Rosi treatment (Fig. 4C). To determine whether the catalase PPRE
alone was sufficient to enhance PPAR-mediated promoter activity, we
constructed a promoter containing a tandem tripeat of the catalase
PPRE (DR1x3). Cos-1 cells were transfected with the DR1x3 or empty
vector (pGL3-promoter) in the presence of PPARg2/RXRa or pTargeT.
Cells transfected with the DR1x3 exhibited a 2-fold (#, p¼0.002)
induction in reporter activity compared with empty vector (Fig. 4D).
Moreover, Cos-1 cells transfected with DR1?3 containing PPARg2,
RXRa or both expression plasmids significantly increased promoter
activity 2.5-, 3.8- and 4.6-fold, respectively (&, po0.001). Rosi
treatment resulted in a significant increase in promoter activity only
in cells transfected with exogenous PPARg alone (6.2-fold,
Time with ActD (hours)
Catalase / Cyclophilin
Relative Fold Induction
Relative Fold Induction
Fig. 3. PPAR-induced catalase mRNA in astrocytes is not due to increased message stability. Primary rat astrocytes were supplemented for 24 h with 10 M CP096, Rosi,
or DMSO (vehicle control) and then treated up to an additional 24 h with 5 mg/mL ActD (transcription inhibitor). Total RNA was harvested at 0, 4, 8, 12, and 24 h post ActD
(A). Densitometric analysis was performed using Typhoon 8600 Phosphoimager and software package ImageQuant 5.1. Catalase mRNA was normalized to the
housekeeping gene, cyclophilin, within each time point. This Northern blot is a representative of at least three independent experiments and data are expressed as
mean7SEM (B). Primary rat astrocytes were transiently transfected with the indicated rat catalase promoter deletion constructs (-1046, -938, -207). The b-gal expression
vector was used to control for transfection efficiency. Luciferase activity was normalized to b-gal and then all the catalase promoter deletion constructs were compared to
the activity of the empty vector control (pGL3-basic) (C). Primary rat astrocytes were transiently transfected with the rat catalase promoter deletion construct containing
the PPRE (-1046), PPARg2, RXRa, or the empty vector (pTargeT) control and then treated with 5 mM Rosi for 24 h. Luciferase activity was normalized to b-gal and then all
the catalase promoter deletion constructs were normalized to the activity of primary rat astrocytes treated transfected with -1046 and pTargeT treated with vehicle
(DMSO) (D). Results of the luciferase promoter reporter assays shown represent mean7SEM of at least five independent experiments performed in triplicate.n, po0.01
compared to all groups; #, po0.01 compared to pTargeT treated with DMSO.
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
po0.0001) or in combination with RXRa (3.9-fold,
compared to their respective DMSO treated controls (Fig. 4D).
A similar response was observed in Cos-1 cells transfected with the
ACOX-PPRE, which was used as a positive control (not shown).
Pioglitazone (Pio)-mediated induction of catalase activity
is dependent on PPARg
Primary rat astrocytes were transfected with PPARg-d.n. and then
treated with a similar PPARg agonist Pio (10mM) for 48 h. Primary rat
astrocytes treated with Pio for 48 h resulted in a 1.8-fold increase in
catalase immunoreactive protein levels. However, primary rat astro-
cytes transfected with the PPARg-d.n. construct failed to exhibit a
Pio-mediated increase in catalase protein (Supplemental data S2).
More importantly, Pio (10mM) significantly increased catalase activ-
ity in primary rat astrocytes compared to vehicle alone (n, po0.001).
The Pio-mediated increase in catalase activity was completely abol-
ished in cells transfected with the PPARg-d.n. construct (Fig. 5A).
Pioglitazone treatment increased promoter activity 3-fold
(n, po0.01) in Cos-1 cells transfected with the rat catalase pro-
moter deletion construct containing the PPRE (-1046), PPARg2, and
RXRa. The Pio-mediated increase in promoter activity was ablated by
the co-transfection of PPARg-d.n. or with the incubation of the PPARg
specific inhibitor GR-259662 (Fig. 5B). Lastly, catalase enzymatic
activity was significantly increased in Cos-1 cells transfected with
PPARg2 and RXRa treated with Pio (10mM) for 24 h (Fig. 5C).
Treatment of primary cultures of normal rat astrocytes with
PPARa, PPARg and RXR agonists increased steady-state levels of
3 x DR1
Fig. 4. PPARg plays a significant role in the regulation of catalase. Transfected PPARg and RXRa bind to the PPRE in the rat catalase promoter determined by EMSA. Cos-1
cells were transfected (supplemental S1), nuclear protein was isolated, incubated with
polyacylamide gel. Supershift assay were performed by incubating 2 or 4 mL anti-PPARg and -PPARa antibodies with reaction mixture (Santa Cruz) (A). Cos-1 cells
were transiently transfected with the catalase promoter deletion construct containing the PPRE (-1046), mPPARg2, mRXRa or pTargeT (empty vector control for the NHR
expression plasmids) and then incubated with Rosi and 9cRA for 24 h. Transfected cells were lysed, luciferase activity was measured, and then normalized to b-gal activity
(B). Results of the promoter reporter assays shown represent mean7SEM of at least five independent experiments performed in triplicate (n, po0.006 for comparison
between open bars vs shaded bars). Cos-1 cells were transiently transfected with b-gal, mPPARg2, mRXRa, and either the catalase promoter deletion construct containing
the PPRE (-1046) or lacking the PPRE (-938). Transfected cells were treated with Rosi for 24 h and promoter activity was determined. Results of the promoter reporter
assays shown represent mean7SEM of at least seven independent experiments performed in triplicate (n, po0.002 compared to DMSO; #, po0.001 compared to catalase
promoter construct lacking PPRE, -938) (C). Cos-1 cells were transiently transfected with the DR1x3 (inset) or empty vector (pGL3-promoter) in the presence of
mPPARg2þmRXRa or pTargeT. Transfected cells were treated with Rosi for 24 h and promoter activity was determined. Results of the promoter reporter assays shown
represent mean7SEM of at least four independent experiments performed in triplicate (#, po0.002 for comparison between DR1x3 vs empty for DMSO treatment; &,
po0.006 compared to respective empty vector treated with Rosi;nn, po0.0001 compared between Rosi vs DMSO;n, po0.05 compared between Rosi vs DMSO) (D).
32P-labeled catalase PPRE, and electrophoresed on a 5% nondenaturing
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
catalase mRNA, immunoreactive protein, and enzymatic activity.
In contrast, treatment of the C6 rat malignant glioma cell line
with a similar panel of PPAR and RXR agonists failed to increase
catalase mRNA, protein levels, and catalase enzymatic activity.
Promoter analysis and cell treatment with ActD indicated that the
increase in astrocyte catalase mRNA was a result of new mRNA
synthesis. In support of this, EMSA analysis revealed that Cos-1
cells transfected with PPARg and RXRa showed these transcrip-
tional regulatory proteins bound to the PPRE of the rat catalase
promoter and increased catalase promoter activity, as compared
to empty (pTargeT) control vector. Finally, the transfection of
PPARg-d.n. completely blocked Pio-induced catalase activity. In
aggregate, these studies reveal that PPARg is responsible for the
upregulation of catalase gene expression and enzymatic activity
in normal rat astrocytes treated with PPARg agonists.
Previous studies have demonstrated the presence of PPAR and
RXR isotypes in the rat CNS. The degree of expression and tissue
localization varies between these different lipid receptors. In the
adult rat brain, PPARa, PPARg and RXRa mRNA have all been
identified in cortical astrocytes and to a lesser extent in cerebellar
granule neurons . In the rat spinal cord, PPARa mRNA and
protein are expressed homogeneously in the gray matter, but in
the white matter are exclusively localized to the astrocytes;
PPARg was not detected in cervical, thoracic or lumbar segments
of the spinal cord . Herein, Western blot analysis revealed the
presence of PPARa and PPARg but not RXRa protein in normal rat
astrocytes, with the constitutive levels of expression for PPARa
and PPARg being markedly less than in C6 glioma cells. These
results are consistent with previous observations showing a
similar increase in constitutive expression of PPARg in human
glioma cells as compared to normal human astrocytes .
PPARa protein was expressed in Lipari human glioblastoma cell
line, but no comparison was made with levels in normal astrocytes at
that particular time . In a separate study, all PPAR isotypes and
RXRb were detected in rat astrocytes by immunoblotting . The
present inability to identify RXRa protein in primary cultures of
normal rat astrocytes confirms previous studies in which RXRa
expression could only be observed using RT-PCR . RXRa has been
detected in human and rat glioma cells and 20 primary cultures
established from biopsies from patients with glioblastomas multi-
forme . Thus, the constitutive expression of not only PPARg, but
also of PPARa and RXRa is greater in glioma cells than in normal cell
counterparts. This increased expression of PPARs and RXRs in glioma
cells compared to the astrocytes may account for the increased
catalase gene expression and enzymatic activity observed in glioma
cells (Figs. 1 and 2).
-1046 + PPARγ2 + RXRα
Catalase Activity (K/g/second)
Empty PPARγ+ RXR
Catalase Activity (K/g/second)
Fig. 5. Pioglitazone-mediated induction of catalase activity is dependent on PPARg. Primary rat astrocytes were transiently transfected with PPARg-d.n. or empty vector
and then treated with Pio (10 mM) for 48 h. Transfected cells were scrape harvested and catalase activity was determined spectrophotometrically at 240 nm. Results are
derived from at least three independent experiments and data are expressed as mean7SEM (n, po0.001 compared to all groups) (A). Cos-1 cells were transiently
transfected with mPPARg2, mRXRa, and either rat catalase promoter deletion construct containing the PPRE (-1046) or empty vector (pGL3-basic) and then treated with
Pio for 24 h. Additionally, Cos-1 cells were also transiently co-transfected (as described above) with PPARg-d.n. or co-treated with the PPARg specific antagonist GR-
259662 plus Pio for 24 h. Cells were lysed and promoter activity was determined. Results of the promoter reporter assays are mean7SEM of at least three independent
experiments performed in triplicate (n, po0.01 compared to all groups) (B). Catalase activity was measured using protein samples from the Cos-1 cells transfected with
mPPARg2 and mRXRa treated with 10 mM Pio for 24 h. Results are derived from at least three independent experiments and data are expressed as mean7SEM (n, po0.01
compared to all groups) (C).
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
Despite this increased expression of PPAR and RXR isotypes, C6
glioma cells incubated with PPAR and RXR agonists failed to
exhibit an increase in catalase expression and/or activity. This
failure to modulate catalase levels might reflect glioma cell
cytotoxicity. PPARg agonists are selectively cytotoxic to human
U87 and A172 glioma cells and to C6 glioma cells by not only
inhibiting proliferation but also by inducing apoptosis. In con-
trast, primary murine astrocytes were unaffected by PPARg
agonist treatment . However, this cytotoxic effect was only
observed in C6 cells incubated with concentrations ranging from
30 to 100 mM of the PPARg agonist ciglitazone. A similar study
was performed treating C6 glioma cells with ciglitazone and Rosi
for 48 h at a concentration of 10 mM and there was roughly 60%
and 35% reduction in cell viability determined by MTT assay,
respectively . We failed to detect significant levels of PPAR
and/or RXR agonist-mediated cytotoxicity at 48 h, assessed in
terms of esterase activity using the fluorescent dye Calcein AM in
primary rat astrocytes and C6 glioma cells (not shown). One
potential reason for the discrepancy at 48 h was that we treated
the cells with media containing 10% serum whereas the former
study was conducted in the absence of serum-containing med-
ium. Thus, the selective ability of PPAR and RXR agonists to
increase catalase expression in normal astrocytes but not in
glioma cells does not appear to reflect glioma cell cytotoxicity
after 48 h of treatment.
Most of the antineoplastic effects of PPARg agonists occur with
concentrations that typically are much higher than what is
required to bind to and activate PPARg suggesting PPARg-depen-
dent effects may be in part a consequence of off-target drug
effects. Moreover, the concentrations of PPARg agonists required
to induce cytotoxicity and antineoplastic effects is at least a
thousand times greater than drug concentrations used to treat
type 2 diabetes mellitus . PPARg agonist-dependent and -
independent effects inhibit cell proliferation, suppress inflamma-
tion, and block angiogenesis, thereby altering tumor microenvir-
onments . These actions can ultimately alter tumor cell
viability and metastatic potential. As noted previously, the pre-
sent experiments treated C6 glioma cells and astrocytes with
PPAR agonists in serum-containing culture conditions, a scenario
that best models in vivo conditions.
Besides the differences in constitutive levels of catalase enzymatic
activity between primary cultures of normal rat astrocytes and rat
malignant glioma cell line C6 (Fig. 1), there was also a greater
expression of catalase mRNA and protein in the glioma cells com-
pared to the astrocytes. These results are consistent with previous
observations made in the 36B10 rat glioma cell line . The
relationship between this apparent increase in catalase activity in
these two rat glioma cell lines and the resistance of glioma cells to
redox-related anticancer therapies such as ionizing radiation remains
unclear. However, it is notable that a comparison of antioxidant
enzyme activities in 10 human and rat glioma cell lines with
sensitivity to 1,3-bis (2-chloroethyl)-1-nitrosourea (BCNU, a che-
motherapeutic and glutathione modulating drug used in the treat-
ment of brain cancer), revealed a significant positive correlation with
catalase activity alone . Additionally, catalase inhibition sensitizes
36B10 glioma cells to oxidative stress. Catalase was inhibited in
36B10 glioma cells by two methods: (1) biochemically, using
3-amino-1,2,4-triazole as a non-competitive inhibitor and (2) using
a catalase shRNA knockdown approach. Both methods reduced
catalase enzymatic activity by ?75% thereby increasing extracellular
H2O2and oxidative stress in the glioma cells. These catalase deficient
glioma cells had increased sensitivity to ionizing radiation, supporting
a regulatory role for catalase in the resistance of glioma cells to
oxidative stress .
The current data support the concept that the selective
manipulation of catalase gene expression and/or activity may
improve normal tissue survival during radiation therapy. Catalase
expression is regulated at the level of tissue-specific transcription,
post-transcription, and post-translation [41,42]. The identification
of functional PPRE in the 50-flanking region of the rat catalase
promoter (located between nt-1027 and -1015 with respect to the
translation start site) supports that there is a transcriptional
mechanism for regulation of catalase gene expression that could
be mediated by TZDs, a drug class presently used to treat
diabetes. This is reinforced by ActD (Fig. 3A), promoter-based
studies (Figs. 3–5) and EMSA (Fig. 4A) revealing that PPAR
agonist-induced upregulation of catalase expression in primary
rat astrocytes is due, at least in part, to agonist binding and
activation of PPAR/RXR heterodimers to the PPRE in the rat
increase in catalase expression and activity in astrocytes by PPAR
agonists would suggest that astrocytes have increased protection
from H2O2-induced damage. Experiments to test this would be of
significant value. Moreover, an in vivo experiment testing
whether this observed increase in catalase expression and activity
in astrocytes resulted in a potential adjuvant therapy in treating
malignant gliomas would be of great interest as most patients
with these tumors have a poor prognosis.
this observed differential
In summary, PPARa and PPARg agonists selectively increase
catalase expression in normal astrocytes but not in glioma cells.
The ability of PPARa and PPARg agonists to selectively upregulate
catalase expression in microvascular endothelial cells and astro-
cytes, which are both critical modulators of the response of the
brain to radiation-induced injury, is promising. Since PPAR
agonists did not increase catalase enzymatic activity or expres-
sion in glioma cells, a potential therapeutic strategy is revealed
for reducing the severity of normal brain injury in glioma
patients, without compromising tumor cell killing. These findings
offer the promise of reducing the severity of normal brain injury
in glioma patients without compromising tumor cell kill. This
concept is reinforced by the growing evidence that PPARg
agonists inhibit proliferation of human and rat glioma cells. Thus,
cancer therapeutic strategies utilizing PPAR agonists could pro-
tect normal tissue through the upregulation of catalase activity
and increase the susceptibility of glioma cells. These agonists
might have significant therapeutic impact as novel agents for
improving the outcome for what remains an aggressive and all
too often fatal neoplasm.
The authors would like to dedicate this work in memory of
Mike E. Robbins and Larry W. Oberley. This work was supported
by National Institutes of Health grants CA82722 (to M.E. Robbins),
CA73612 and CA66081 (to F.E. Domann) and NS24621 (to S.A.
Appendix A. Supporting information
Supplementary data associated with this article can be found in
the online version at http://dx.doi.org/10.1016/j.redox.2012.12.006.
 National Cancer Institute and Central Brain Tumor Registry, [/http://www.
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79
 R. Stupp, W.P. Mason, M.J. van den Bent, M. Weller, B. Fisher, M.J. Taphoorn, Download full-text
K. Belanger, A.A. Brandes, C. Marosi, U. Bogdahn, et al., Radiotherapy plus
concomitant and adjuvant temozolomide for glioblastoma, The New England
Journal of Medicine 352 (2005) 987–996.
 F.G. Davis, S. Freels, J. Grutsch, S. Barlas, S. Brem, Survival rates in patients
with primary malignant brain tumors stratified by patient age and tumor
histological type: an analysis based on surveillance, epidemiology, and end
results (SEER) data, 1973–1991, Journal of Neurosurgery 88 (1998) 1–10.
 T.E. Schultheiss, L.C. Stephens, Invited review: permanent radiation myelo-
pathy, British Journal of Radiology 65 (1992) 737–753.
 J.P. Imperato, N.A. Paleologos, N.A. Vick, Effects of treatment on long-term
survivors with malignant astrocytomas, Annals of Neurology 28 (1990)
 J.R. Crossen, D. Garwood, E. Glatstein, E.A. Neuwelt, Neurobehavioral sequelae
of cranial irradiation in adults: a review of radiation-induced encephalo-
pathy, Journal of Clinical Oncology 12 (1994) 627–642.
 D.D. Clarke, L. Sokoloff, Circulation and Energy Metabolism of the Brain,
Lippincott-Raven, Philadelphia, 1999.
 R. Dringen, J.M. Gutterer, J. Hirrlinger, Glutathione metabolism in brain
metabolic interaction between astrocytes and neurons in the defense against
reactive oxygen species, European Journal of Biochemistry 267 (2000)
 S. Peuchen, J.P. Bolanos, S.J. Heales, A. Almeida, M.R. Duchen, J.B. Clark,
Interrelationships between astrocyte function, oxidative stress and antiox-
idant status within the central nervous system, Progress in Neurobiology 52
 K.J. Smith, R. Kapoor, P.A. Felts, Demyelination: the role of reactive oxygen
and nitrogen species, Brain Pathology 9 (1999) 69–92.
 A. Spector, Essentiality of fatty acids, Lipids 34 (1999) S1–S3.
 M. Gottlicher, E. Widmark, Q. Li, J.A. Gustafsson, Fatty acids activate a chimera of
the clofibric acid-activated receptor and the glucocorticoid receptor, Proceedings
of the National Academy of Sciences USA 89 (1992) 4653–4657.
 S.A. Kliewer, S.S. Sundseth, S.A. Jones, P.F. Brown, G.B. Wisely, C.S. Koble,
P. Devchand, W. Wahli, T.M. Willson, J.M. Lenhard, J.M. Lehmann, Fatty Acids
and eicosanoids regulate gene expression through direct interactions with
peroxisome proliferator-activated receptors a and g, Proceedings of the National
Academy of Sciences USA 94 (1997) 4318–4323.
 S.A. Kliewer, K. Umesono, D.J. Noonan, R.A. Heyman, R.M. Evans, Convergence
of 9-cis retinoic acid and peroxisome proliferator signalling pathways
through heterodimer formation of their receptors, Nature 358 (1992)
 A unified nomenclature system for the nuclear receptor superfamily. Cell 97
 M. Yoon, The role of PPAR[alpha] in lipid metabolism and obesity: Focusing
on the effects of estrogen on PPAR[alpha] actions, Pharmacological Research
60 (2009) 151–159.
 P. Tontonoz, B.M. Spiegelman, Fat and beyond: the diverse biology of PPARg,
Annual Review of Biochemistry 77 (2008) 289–312.
 S.M. Reilly, C.-H. Lee, PPAR[delta] as a therapeutic target in metabolic disease,
FEBS Letters 582 (2008) 26–31.
 C. Grommes, G.E. Landreth, M. Sastre, M. Beck, D.L. Feinstein, A.H. Jacobs,
U. Schlegel, M.T. Heneka, Inhibition of in vivo glioma growth and invasion by
peroxisome proliferator-activated receptor g agonist treatment, Molecular
Pharmacology 70 (2006) 1524–1533.
 J.M. Perez-Ortiz, P. Tranque, C.F. Vaquero, B. Domingo, F. Molina, S. Calvo,
J. Jordan, V. Cena, J. Llopis, Glitazones differentially regulate primary astro-
cyte and glioma cell survival, Journal of Biological Chemistry 279 (2004)
 D.L. Montgomery, Astrocytes: form, functions, and roles in disease, Veter-
inary Pathology 31 (1994) 145–167.
 S. Vartak, R. McCaw, C.S. Davis, M.E.C. Robbins, A.A. Spector, g-linolenic acid
(GLA) is cytotoxic to 36B10 malignant rat astrocytoma cells but not to
‘normal’ rat astroxytes, British Journal of Cancer 77 (1998) 1612–1620.
 M. Preuss, G.D. Girnun, C.J. Darby, N. Khoo, A.A. Spector, M.E. Robbins, Role of
antioxidant enzyme expression in the selective cytotoxic response of glioma
cells to gamma-linolenic acid supplementation, Free Radical Biology and
Medicine 28 (2000) 1143–1156.
 G.D. Girnun, F.E. Domann, S.A. Moore, M.E. Robbins, Identification of a
functional peroxisome proliferator-activated receptor response element in
the rat catalase promoter, Molecular Endocrinology 16 (2002) 2793–2801.
 M.M. Bradford, A rapid and sensitive method for the quantitation of
microgram quantities of protein utilizing the principle of protein-dye bind-
ing, Analytical Biochemistry 72 (1976) 248–254.
 N. Marx, U. Schonbeck, M.A. Lazar, P. Libby, J. Plutzky, Peroxisome
proliferator-activated receptor gamma activators inhibit gene expression
and migration in human vascular smooth muscle cells, Circulation Research
83 (1998) 1097–1103.
 R.F. Beers, I.W. Sizer, A spectrophotometric method for measuring the
breakdown of hydrogen peroxide by catalase, Journal of Biological Chemistry
195 (1952) 133–140.
 N.R. Draper, H. Smith, Applied Regression Analysis, John Wiley & Sons, Inc,
New York, 1998, pp. 221–229.
 J. DiRenzo, M. Soderstrom, R. Kurokawa, M.H. Ogliastro, M. Ricote, S. Ingrey,
A. Horlein, M.G. Rosenfeld, C.K. Glass, Peroxisome proliferator-activated
receptors and retinoic acid receptors differentially control the interactions
of retinoid X receptor heterodimers with ligands, coactivators, and corepres-
sors, Molecular and Cellular Biology 17 (1997) 2166–2176.
 I. Schulman, G. Shao, R. Heyman, Transactivation by retinoid X receptor-
peroxisome proliferator-activated receptor g (PPARg) heterodimers: inter-
molecular synergy requires only the PPARg hormone-dependent activation
function, Molecular Cell Biology 18 (1998) 3483–3494.
 J.O. Nwankwo, M.E. Robbins, Peroxisome proliferator-activated receptor-
gamma expression in human malignant and normal brain, breast and
prostate-derived cells, Prostaglandins, Leukotrienes and Essential Fatty Acids
64 (2001) 241–245.
 A. Cimini, L. Cristiano, A. Bernardo, S. Farioli-Vecchioli, S. Stefanini, M.P.
Ceru, Presence and inducibility of peroxisomes in a human glioblastoma cell
line, Biochimica Biophysica Acta 1474 (2000) 397–409.
 L. Cristiano, A. Bernardo, M.P. Ceru, Peroxisome proliferator-activated recep-
tors (PPARs) and peroxisomes in rat cortical and cerebellar astrocytes,
Journal of Neurocytology 30 (2001) 671–683.
 T.E. Cullingford, K. Bhakoo, S. Peuchen, C.T. Dolphin, R. Patel, J.B. Clark,
Distribution of mRNAs encoding the peroxisome proliferator-activated
receptor alpha, beta, and gamma and the retinoid X receptor alpha, beta,
and gamma in rat central nervous system, Journal of Neurochemistry 70
 H. Bouterfa, T. Picht, D. Ke, C. Herbold, E. Noll, P.M. Black, K. Roosen, C. Tonn,
Retinoids inhibit human glioma cell proliferation and migration in primary
cell cultures but not in established cell lines, Neurosurgery 46 (2000) 419.
 T. Zander, J.A. Kraus, C. Grommes, U. Schlegel, D. Feinstein, T. Klockgether,
G. Landreth, J. Koenigsknecht, M.T. Heneka, Induction of apoptosis in human
and rat glioma by agonists of the nuclear receptor PPARgamma, Journal of
Neurochemistry 81 (2002) 1052–1060.
 D. Panigrahy, S. Singer, L.Q. Shen, C.E. Butterfield, D.A. Freedman, E.J. Chen,
M.A. Moses, S. Kilroy, S. Duensing, C. Fletcher, et al., PPARgamma ligands
inhibit primary tumor growth and metastasis by inhibiting angiogenesis,
Journal of Clinical Investigation 110 (2002) 923–932.
 D. Panigrahy, S. Huang, M.W. Kieran, A. Kaipainen, PPARgamma as a
therapeutic target for tumor angiogenesis and metastasis, Cancer Biology
and Therapy 4 (2005) 687–693.
 W. Zhong, T. Yan, R. Lim, L.W. Oberley, Expression of superoxide dismutases,
catalase, and glutathione peroxidase in glioma cells, Free Radical Biology and
Medicine 27 (1999) 1334–1345.
 P.S. Smith, W. Zhao, D.R. Spitz, M.E. Robbins, Inhibiting catalase activity
sensitizes 36B10 rat glioma cells to oxidative stress, Free Radical Biology and
Medicine 42 (2007) 787–797.
 H. Van Remmen, M.D. Williams, H. Yang, C.A. Walter, A. Richardson, Analysis
of the transcriptional activity of the 50-flanking region of the rat catalase gene
in transiently transfected cells and in transgenic mice, Journal of Cell
Physiology 174 (1998) 18–26.
 C. Cao, Y. Leng, X. Liu, Y. Yi, P. Li, D. Kufe, Catalase is regulated by
ubiquitination and proteosomal degradation. Role of the c-Abl and Arg
tyrosine kinases, Biochemistry 42 (2003) 10348–10353.
N.K.H. Khoo et al. / Redox Biology 1 (2013) 70–79