Oxygen causes cell death in the developing brain

Ursula Felderhoff-Mueser,a,1 Petra Bittigau,b,c,1 Marco Sifringer,b,c,1 Bozena Jarosz,d

Elzbieta Korobowicz,d Lieselotte Mahler,a Turid Piening,a Axel Moysich,a Tilman Grune,c

Friederike Thor,e Rolf Heumann,e Christoph Bqhrer,a and Chrysanthy Ikonomidoub,c,*

aDepartment of Neonatology, Humboldt University Berlin, Charite, Campus Virchow Klinikum, 13353 Berlin, GermanybDepartment of Pediatric Neurology, Humboldt University Berlin, Charite, Campus Virchow Klinikum, 13353 Berlin, GermanycNeuroscience Research Center, Humboldt University Berlin, Charite, 10117 Berlin, GermanydDepartment of Clinical Pathology, Medical University of Lublin, PolandeMolecular Neurobiochemistry, Ruhr University, Bochum, Germany

Received 16 April 2004; revised 5 July 2004; accepted 30 July 2004

Substantial neurologic morbidity occurs in survivors of premature

birth. Premature infants are exposed to partial oxygen pressures that

are fourfold higher compared to intrauterine conditions, even if no

supplemental oxygen is administered. Here we report that short

exposures to nonphysiologic oxygen levels can trigger apoptotic

neurodegeneration in the brains of infant rodents. Vulnerability to

oxygen neurotoxicity is confined to the first 2 weeks of life, a period

characterized by rapid growth, which in humans expands from the

sixth month of pregnancy to the third year of life.

Oxygen caused oxidative stress, decreased expression of neurotro-

phins, and inactivation of survival signaling proteins Ras, extracellular

signal-regulated kinase (ERK 1/2), and protein kinase B (Akt). The

synRas-transgenic mice overexpressing constitutively activated Ras

and phosphorylated kinases ERK1/2 in the brain were protected

against oxygen neurotoxicity. Our findings reveal a mechanism that

could potentially damage the developing brain of human premature

neonates.

D 2004 Elsevier Inc. All rights reserved.

Keywords: Apoptosis; Development; Infant rat; Oxidative stress; Survival;

Oxygen

Introduction

Advances in neonatal intensive care have markedly improved

survival of premature and term infants. Unfortunately, neurologic

morbidity did not decrease at the same pace. A substantial

proportion of very low birth weight infant survivors have neuro-

logic deficits which affect motor and cognitive function (Hack et

al., 2002; Ment et al., 2000; Wood et al., 2000; Vohr et al., 2000).

Very common are speech and language difficulties, attention deficit

hyperactivity disorder, and dyslexia. Brain imaging studies of

survivors of premature birth have demonstrated that motor deficits

correlate with white matter damage whereas cognitive deficits

correlate with decreased volume of grey matter structures

(Abernethy et al., 2002; Ajayi-Obe et al., 2000; Nosarti et al.,

2002). These findings suggest that neuronal loss, occurring in the

brains of premature infants postnatally, is partly responsible for

their neurologic morbidity. Obvious catastrophic events, such as

intracerebral bleeding or parenchymal infarction, only partly

explain neurologic disability. An increasing body of evidence

casts serious doubts on the primary role of hypoxia and ischemia in

injury to the developing brain, particularly in preterm infants, while

recent work has emphasized the role of intrauterine and neonatal

infections (Murphy et al., 1995; Taylor et al., 1999). Still, in many

cases an obvious cause of brain damage is missing.

In recent years, we learned about silent triggers of cell death in

the developing brain. It has been reported that compounds that are

used as sedatives (Ikonomidou et al., 1999, 2000), anesthetics

(Jevtovic-Todorovic et al., 2003), or anticonvulsants (Bittigau et al.,

2002) in neonatal intensive care units and which alter physiologic

synaptic activity, such as antagonists at N-methyl-d-aspartate

(NMDA) receptors (ketamine, nitric oxide), agonists at GABAA

receptors (barbiturates, benzodiazepines, anesthetics), and sodium

channel blockers (phenytoin, valproate), can cause massive

apoptotic neurodegeneration in infant rats and mice. This neuro-

toxic effect in rodents is strictly confined to a developmental period

0969-9961/$ - see front matter D 2004 Elsevier Inc. All rights reserved.

doi:10.1016/j.nbd.2004.07.019

Abbreviations: Akt, protein kinase B; BDNF, brain-derived neuro-

trophic factor; ERK, extracellular signal-regulated kinase; GSH, reduced

glutathione; GSSG, oxidized glutathione; MAPK, mitogen-activated

protein kinase; MDA, malondialdehyde; NGF, nerve growth factor; Nv,

numerical densities; NT-3, neurotrophin 3; PI3-kinase, phosphatidylinosi-

tol-3 kinase.

* Corresponding author. Department of Pediatric Neurology, Humboldt

University Berlin, Charite, Children’s Hospital, Campus Virchow, Augus-

tenburger Platz 1, D-13353 Berlin, Germany. Fax: +49 30 450566920.

E-mail address: hrissanthi.ikonomidou@charite.de (C. Ikonomidou).1 These three authors contributed equally to this work.

Available online on ScienceDirect (www.sciencedirect.com.)

www.elsevier.com/locate/ynbdi

Neurobiology of Disease 17 (2004) 273–282

characterized by rapid brain growth, which starts prenatally in

humans and expands to several years after birth (Dobbing and

Sands, 1979; Ikonomidou et al., 1999; 2000; Olney et al., 2002).

This comparison points towards the likely possibility that human

infants may be susceptible to and may sustain iatrogenic brain

damage from treatments that are considered safe in older patients.

Such mechanisms could potentially silently lead to diffuse brain

injury in infancy and result in cognitive and motor impairment that

will become evident later in life.

Here we present evidence that oxygen, which is being used in

resuscitation and treatment of respiratory distress syndrome,

pulmonary hypertension, and in the context of cardiac surgery in

infants, is a powerful trigger for apoptotic neuronal death in the

developing brain.

Materials and methods

Animal experiments

All animal experiments were performed in accordance to the

guidelines of the Humboldt University in Berlin, Germany.

Wistar rat pups (Bundesinstitut fqr gesundheitlichen Verbrau-

cherschutz und Veterin7rmedizin BgVV, Berlin, Germany), ages 0–

14 days, weighing 4–49 g, or 7-day-old mice (synRas and wild

type) were placed together with their mothers into a chamber and

exposed to a 40%, 60%, or 80% oxygen/air environment for 2–72

h. Mothers were switched every 24 h to prevent adult respiratory

lung disease. Littermates kept in room air served as controls.

Animals were sacrificed at 2, 6, 12, 24, 48, 72 h following oxygen

exposure.

Each experimental group consisted of 6–10 pups. Exposure to

80% oxygen over 12 h was performed with synRas-transgenic

mice. The synRas-transgenic mice were first generated on a B6

background and were then crossed back to NMRI background over

30 times, as previously described (Heumann et al., 2000).

Genotype of mice was determined by PCR as described (Heumann

et al., 2000).

To determine blood oxygen content during hyperoxia, infant

pups were placed into the hyperoxia chamber for 10 min, were then

taken out and anesthetized with ether, placed back into the

hyperoxia chamber for 3 min, and subsequently a left cardiac

puncture was performed subcutaneously using a 32-gauge hypo-

dermic needle. One hundred microliters of arterial blood was

obtained and immediately subjected to analysis of pH, paO2

(partial pressure of oxygen in mm Hg), paCO2, and bicarbonate.

Histology

For histological analysis of the brains, animals received an

overdose of intraperitoneal chloral hydrate and were transcardially

perfused with heparinized 0.1 M PBS, pH 7.4, followed by 4%

paraformaldehyde in cacodylate buffer, pH 7.4. Brains were

postfixed for 3 days at 48C and processed for DeOlmos silver

staining, terminal deoxynucleotide transferase-mediated dUTP

nick end labeling (TUNEL), or immunohistochemistry.

For light microscopy studies of plastic sections, pups were also

perfused transcardially with 1% paraformaldehyde and 1.5%

glutaraldehyde in pyrophosphate buffer, pH 7.4.

For DeOlmos cupric silver staining, brains were embedded in

agar, coronal sections of 70 Am thickness were serially cut on a

vibratome (Leica VT 1000 S, Nugloch, Germany) and processed

for staining with silver nitrate and cupric nitrate according to the

method of DeOlmos and Ingram (1971). Degenerating cells were

identified by a distinct dark appearance due to the silver

impregnation.

TUNEL staining was performed using the ApopTag Peroxidase

kit (S 7100, Oncor Appligene, Heidelberg, Germany) according to

the manufacturer’s instructions. Briefly, after pretreatment with

proteinase K and quenching of endogenous peroxidase, sections

were incubated first in equilibration buffer followed by working

strength TdT enzyme (incorporating digoxigenin labeled dUTP

nucleotides to free 3V-OH DNA termini, 1 h at 378C). Sectionswere incubated in stop/wash buffer (30 min, 378C), then with anti-

digoxigenin-peroxidase conjugate (30 min) followed by DAB

substrate (Sigma, Deisenhofen, Germany), and lightly counter-

stained with methylgreen.

For hematoxylin and eosin (H&E) staining, brain tissue was

paraffin embedded and subsequently cut into 5-Am sections.

Following deparaffinization, H&E staining was performed accord-

ing to standard protocols.

For plastic embedding, brains were sliced transversely into

70 Am serial sections. These sections were osmicated overnight

(1% osmium tetroxide), dehydrated in graded ethanols, cleared in

toluene, and embedded in araldite. Thin sections, 1 Am thick, were

cut at selected rostrocaudal levels of the brain, using glass knives

(1/2 in. wide). These sections were heat dried on glass slides and

stained with azure II and methylene blue for evaluation by light

microscopy. Ultrathin sections were cut and viewed by trans-

mission electron microscopy (Zeiss, EM 900).

For immunohistochemical staining, brains were embedded in

paraffin, coronal sections, 5 Am thick, were cut on a Microtome

HM 360 (Microm International GmbH, Walldorf, Germany) and

mounted onto 3-aminopropyltrietoxylane-coated (Sigma) glass

slides. Sections were microwaved in 10 mM citrate buffer, pH

6.5, at 750 W. Endogenous peroxidase activity was blocked

with 0.6% v/v hydrogen peroxide (15 min), sections were

incubated with normal goat serum (20 min) and left overnight at

48C with cleaved caspase 3 antibody (1:100, Cell Signaling,

New England Biolabs GmbH, Frankfurt, Germany). Negative

controls were performed by including the peptide immunogen as

a competitor of antibody binding according to the manufactur-

er’s instructions. Sections were then treated with goat anti-rabbit

IgG. After detection with the ABC kit (Vector Laboratories,

Peterborough, UK), positive cells were visualized with diami-

nobenzidine (DAB, Sigma) and counterstained with hematox-

ylin–eosin.

Quantification of cell death in different brain regions

Following oxygen exposure, degenerating cells were deter-

mined in sections (70 Am) stained by the DeOlmos cupric silver

method (rats) or in TUNEL-stained sections (12 Am; mice) in the

frontal, parietal, cingulate, retrosplenial cortex, caudate nucleus

(mediodorsal part), nucleus accumbens, corpus callosum and

adjacent white matter, thalamus (laterodorsal, mediodorsal, ventral

nuclei), hippocampal dentate gyrus, subiculum, and hypothalamus

by means of a stereological dissector (West and Gundersen, 1990),

estimating mean numerical densities (Nv) of degenerating cells

(cells/mm3). An unbiased counting frame (0.05 � 0.05 mm,

dissector height 0.07 or 0.012 mm) and a high aperture objective

were used for the sampling. The Nv for each brain region was

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282274

determined with 8–10 dissectors. To assess overall severity of cell

death and enable comparisons among treatment groups, a scoring

system was created as follows: numerical densities of degenerated

cells in 13 regions were determined. These values were added to

give a cumulative severity score for degeneration within each

brain.

RT-PCR

Animals (n = 4–6/group) were decapitated, brains were

immediately removed, and tissue was microdissected from the

cingulate and parietal cortex, the corpus callosum, the striatum, and

the thalamus, subsequently snap frozen in liquid nitrogen and

stored at �808C until analysis.

Total cellular RNA was isolated from snap frozen tissue by

acidic phenol/chloroform extraction (Peqlab, Erlangen, Germany)

and DNase treated; 500 ng of RNA was reverse transcribed with

Moloney murine leukemia virus reverse transcriptase (Promega,

Madison, USA) in 25 Al of reaction mixture. The resulting cDNA

(2 Al) was amplified by polymerase chain reaction. The following

oligonucleotide primers for the PCR reactions were used: BDNF:

sense primer 5V-CGACGTCCCTGGCTGGACACTTTT-3V, anti-

sense primer 5V-AGTAAGGGCCCGAACATACGATTGG-3V,GenBank sequence D10938; NT-3: sense primer 5V-GGTCA-GAATTCCAGCCGATGATTGC-3V, antisense primer 5V-CAGCGCCAGCCTACGAGTTTGTTGT-3V, GenBank sequence

M34643; NGF 5V-sense primer 5V-CTGAACCAATAGC-

TGCCCGTGTGAC-3V, antisense primer 5V-GGCAGCCT-

GTTTGTCGTCTGTTGTC-3V, GenBank sequence M36589;

internal standard g-actin: sense primer 5V-CCCTAAGGC-

CAACCGTGAAAAGATG-3V, antisense primer 5V-GAACCGCT-CATTGCCGATAGTGATG-3V, GenBank sequence V01217.

cDNA was amplified in 30–32 cycles, consisting of denaturing

over 30 s at 948C, annealing over 45 s at 568C, and primer

extension over 45 s at 728C. Amplified cDNA was subjected to

polyacrylamide gel electrophoresis, subsequent silver staining

(Lohmann et al., 1995), and densitometric analysis.

Western blot

Snap-frozen tissue was homogenized in 1% SDS buffer (pH

7.3, containing 250 mM EDTA and 1 tablet protease inhibitor

Boehringer complete; Roche Diagnostics GmbH, Mannheim,

Germany) by trituration through a 21-G needle, heated to 908C,and centrifuged at 15,000 � g (10 min). Total cellular proteins

(30 Ag/lane cytosolic fraction) were separated on a 10%

polyacrylamide gel and transferred onto nitrocellulose membrane

(Hybond ECL, Amersham International, Bucks, UK). For

analysis of the extracellular signal regulated kinase (ERK1/2)

pathway, a rabbit polyclonal p44/42 ERK1/2 antibody (1:1000)

and a mouse monoclonal phospho-p44/42 ERK1/2 (Thr202/

Tyr204) E10 antibody (1:1000) (Cell Signaling, New England

Biolabs GmbH) were administered. The protein kinase B (Akt)

pathway was analyzed by using a rabbit polyclonal Akt

antibody (1:1000) and a rabbit polyclonal phospho-Akt

(Ser473) antibody (1:1000, Cell Signaling).

Secondary incubations were with HRP-linked anti-mouse or

anti-rabbit antibody. Positive signals were visualized using

enhanced chemiluminescence (Amersham International) and serial

exposures were made to radiographic film (Hyperfilm ECL,

Amersham International).

Ras pull down assay

For measurements of Ras-activity, five animals were used for

each experimental group. Tissues dissected from cortex were

Dounce homogenized on ice in lysis buffer [1% NP-40, 150 mM

NaCl, 50mMTris–HCl, 15mMMgCl2, 40mMNaF, 10mMEDTA,

0.1% SDS, 0.1% sodiumdesoxycholate, Complete Mini Protease

Inhibitor Cocktail EDTA free (Roche Diagnostics) at pH 7.4 and

centrifuged at 13 000� g (5 min, 48C)]. Precipitations were carriedout by adding 10 Al of bacterial lysate containing Ras binding

domain glutathione-S-transferase fusion protein to 10 Al glutathionesepharose (Amersham) and incubating for 1.5 h at 48C, washed threetimes with wash buffer (50 mM Tris–HCl, pH 7.4, 100 mM NaCl, 1

mM DTT) and once with fish buffer [50 mM Tris–HCl, pH 7.4, 100

mM NaCl, 2 mMMgCl2, 10% (v/v) Glycerin, 1% (v/v) NP-40]. Six

hundred micrograms protein was then incubated with the prepared

glutathione sepharose beads at 48C on a rotation wheel. The

precipitates were collected by centrifugation at 8000 � g (2 min,

48C) and washed three times with fish buffer. Finally, the

immunoprecipitates were dissolved in Laemmli buffer, boiled at

958C, and centrifuged 10 min at 13000 � g. Samples were then

subjected to gel electrophoresis and Western blot analysis using

monoclonal anti-Ras (1:10 000 in TBS-T, upstate Biotechnology).

As control, 15 Ag protein of the lysates was analyzed by gel

electrophoresis and immunoblotting using the monoclonal anti-Ras

antibody. Total Ras and GTP-bound Ras were semiquantified

densitometrically. The mean values were obtained by calculating

the ratios between GTP-bound and total Ras.

Malondialdehyde measurement

Oxidative stress parameters were determined after exposure of

the pups to 6 h of 80% oxygen and immediately upon end of the

exposure. Each group consisted of 10 animals. All tissue samples

were homogenized in 1 ml 1 mM BHT. Determination of the lipid

peroxidation product malondialdehyde (MDA) in brain homoge-

nates was performed according to Wong et al. (1987), with

modifications by Sommerburg et al. (1993). The supernatants were

boiled, in the presence of thiobarbituric acid, for 60 min and the

reaction then stopped by the cooling of the samples in an ice bath.

The neutralized samples were analyzed on an isocratic reversed-

phase HPLC system using a Supelcosil column (Supelco,

Taufkirchen, Germany; 150 � 4 mm LG-18-S, 5 AM) and a

potassium phosphate buffer/methanol eluent. Detection was

performed by fluorescence with an excitation from 525 nm and

an emission from 550 nm.

Protein carbonyl measurement

All tissue samples were homogenized in 1 ml 1 mM BHT and

lysed for 1 h (at 48C) and then centrifuged for 15 min at 14,000

rpm. The resulting supernatant was used to determine the protein

carbonyl content. Protein carbonyl content was determined on

the homogenized tissue supernatant with a concentration of

proteins between 0.5 and 5 mg/ml employing an ELISA method

(Buss et al., 1997), with the appropriate modifications steps

(Sitte et al., 1998). The detection system used was an anti-

dinitrophenyl rabbit IgG-antiserum (Sigma) as the primary

antibody and a monoclonal anti-rabbit IgG antibody peroxidase

conjugate (Sigma) as the secondary antibody. Development was

performed with o-phenylenediamine.

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282 275

Glutathione determination

For the determination of reduced glutathione (Beutler et al.,

1963) and oxidized glutathione (Hissin and Hilf, 1976), the organs

were homogenized and the homogenates were treated with a

mixture of metaphosphoric acid, EDTA, and NaCl. After centri-

fugation, aliquots were taken for neutralization with disodiumhy-

drogenphosphate followed by addition of DTNB. Reduced

glutathione (GSH) was determined after reaction with DTNB in

a spectrophotometer at 412 nm. For the determination of oxidized

glutathione (GSSG), the autoxidation of GSH was stopped by

addition of N-ethylmaleimide. After addition of sodium hydroxide,

GSSG was modified using o-phthalaldehyde. GSSG was deter-

mined at a spectrofluorometer (excitation: 350 nm, emission: 420

nm) using GSSG standards for quantification.

Statistical analyses

Values are presented as mean F SEM. Comparisons among

groups were made using analysis of variance (ANOVA) or

unpaired Student’s t test, as appropriate.

Results

Hyperoxia causes apoptotic cell death in the developing rat brain

To determine whether exposure to a high oxygen environment

may cause cell death in the developing brain, we initially exposed

7-day-old rats with their mothers to 80% oxygen over a period of

24 h. Blood gas analysis performed in four pups, as described in

Materials and methods, revealed a partial oxygen pressure in

arterial blood taken from the left cardiac ventricle of 182 F 15 mm

Hg and an oxygen saturation of 100%. Pups were killed and

transcardially perfused immediately after the end of oxygen

exposure. Histological evaluation of the brains by means of

DeOlmos cupric silver or TUNEL staining revealed a light pattern

of neurodegeneration attributable to programmed cell death in the

brains of pups exposed only to room air (Fig. 1B). Hyperoxia

increased numerical densities of degenerating cells in various brain

regions in comparison to unexposed littermates. Brain regions

affected were the caudate nucleus, nucleus accumbens, layers II

and IV of the frontal, parietal, cingulate, and retrosplenial cortices,

as well as white matter tracts within the forebrain (Figs. 1A, C, D

and 2A, B). The distribution pattern of hyperoxia-induced cell

death is schematically depicted in Fig. 2A, and the numerical

densities of degenerating cells in different brain regions are shown

in Fig. 2B. Immunohistochemical staining with an antibody against

the active P20 subunit of caspase 3 revealed robust activation of

caspase 3 in degenerating neurons (Figs. 1F and G). By H&E

staining, methylene blue/Azur II staining of plastic sections, and

electron microscopy, it was determined that changes within

degenerating cells were similar to those described in neurons

undergoing physiological cell death (Figs. 1E, H, and I).

To determine what length of hyperoxia exposure is necessary to

cause apoptotic cell death in the developing rat brain, we exposed

7-day-old pups to 80% O2 for 2, 6, 12, 24, 48, or 72 h and killed

the pups at 24, 48, or 72 h after the beginning of hyperoxia.

Analysis of the brains by means of silver staining revealed that

after a 2-h exposure to hyperoxia, significant cell death occurred at

24 h. Following a 6- or 12-h exposure, the amount of apoptotic cell

death detected at 24 h increased significantly, following a 24-h

exposure no further enhancement was seen (Fig. 2C). After longer

exposure periods (48 and 72 h), the amount of cell death detected

in the brains upon the end of hyperoxia exposure decreased, most

likely because in the meantime apoptotic cells had been eliminated

and were not more amenable to detection by histological

techniques (Fig. 2C). An alternative explanation could be that a

tolerance mechanism to oxidative stress becomes active in the

brains of rat pups exposed to hyperoxia over longer time periods.

To determine how low oxygen concentrations can result in

apoptotic neurodegeneration in the developing rat brain, we

exposed 7-day-old rats to 40% or 60% oxygen over 12 h. Analysis

of the brains at 24 h after beginning of hyperoxia revealed that the

pups exposed to 40% oxygen had apoptotic scores similar to those

of littermates exposed to normoxia. Exposure to 60% oxygen

resulted in significant increase of apoptotic cell death in the brain

[apoptotic scores of 18,521 F 2351, n = 6 at normoxia; 23,592 F6351 at 40% O2, n = 6 (nonsignificant); 103,352 F 18,295, n = 6

at 60% O2 (P b 0.001, Student’s t test compared to normoxia);

152,359 F 22,851, n = 8 at 80% O2 (P b 0.001, Student’s t test

compared to normoxia)].

Oxygen-induced apoptotic cell death is age dependent

To determine whether the apoptotic response to hyperoxia may

differ as a function of developmental age, we subjected rats at

postnatal ages P0, P3, P7, P14, and P20 to 80% oxygen over a 24-h

period and perfused the animals immediately after end of the

exposure. These experiments revealed that there is a time window

between P0 and P14 when various neuronal populations are

vulnerable to hyperoxia-induced cell death. On P0, mainly

thalamic nuclei, caudate nucleus, putamen, hypothalamus, and

white matter tracts were affected. Most severe involvement of

cortical areas was seen at the age of 7 days (Fig. 2D) and the

highest overall vulnerability at the ages of 3 and 7 days. In 14-day-

old rat pups, some degenerating cells were detected within the

dentate gyrus following exposure to 80% oxygen over 24 h.

Oxidative stress induced by hyperoxia in the developing rat brain

Free radicals can cause cell death by an apoptotic mechanism.

To investigate whether hyperoxia may lead to production of free

radicals in the immature rat brain, we exposed 7-day-old rat pups to

80% O2 over 6 h and analyzed levels of reduced and oxidized

glutathione (GSH and GSSG), protein carbonyls, and the lipid

peroxidation product malondialdehyde (MDA) in brain tissue

taken from the left and right hemispheres. A significant increase of

GSSG and the GSSG/GSH ratio was detected in the brains of rats

exposed to hyperoxia (n = 10) compared with those exposed to

room air (n = 10). Protein carbonyls showed a trend towards

increase in the brains of rats exposed to hyperoxia, but this trend

did not reach significance (Fig. 3A). No significant change in

malondialdehyde levels was detected. These findings indicate that

oxidative stress is induced in the developing rat brain following

exposure to 80% oxygen.

To verify that hyperoxia-induced oxidative stress contributes to

apoptotic cell death in the developing rat brain, we exposed 7-day-

old rats to 12 h of hyperoxia (80% O2) and treated them before

placing them into the hyperoxia chamber (0 h) and upon end of the

hyperoxia exposure (12 h) with either N-acetylcysteine (400 mg/kg

ip) or vehicle. Quantification of degenerating cells at 24 h revealed

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282276

that N-acetylcysteine pretreated pups displayed significantly less

apoptotic neurodegeneration than vehicle-treated rats (Fig. 3B).

Hyperoxia leads to down-regulation of neurotrophins in the infant

rat brain

Neurotrophins provide trophic support to developing neurons

and their withdrawal may lead to neuronal death (Huang and

Reichardt, 2001; Lee et al., 2001). To explore potential mecha-

nisms involved in pathogenesis of apoptotic neurodegeneration in

the developing brain following exposure to hyperoxia, we tested

whether and how hyperoxia may affect expression of brain-derived

neurotrophic factor (BDNF), nerve growth factor (NGF), neuro-

trophin 3 (NT-3) and neurotrophin 4 (NT-4) in the cingulate/

retrosplenial cortex, caudate nucleus, and thalamus in P7 rats. For

this purpose, P7 rats were exposed to an 80% O2 environment over

2, 6, 12, or 24 h (n = 4/group) and sacrificed immediately after end

of the exposure. Hyperoxia triggered reduction in mRNA levels for

all four trophic factors, which was most prominent after 2–6 h of

exposure in all brain regions examined (Fig. 4).

Western blot analysis revealed that down-regulation of mRNA

levels for BDNF, NGF, NT-3, and NT-4 was associated with

decreased levels of the active, phosphorylated isoforms of the

serine–threonin kinase Akt (p-Akt, protein kinase B), and the

member of the mitogen-activated protein kinase (MAPK) extrac-

ellular signal-regulated protein kinase ERK1/2 (p-ERK1/2) (Fig.

Fig. 1. Morphological features of hyperoxia-induced brain damage in infant rats. Light microscopic overviews of histological sections and electron micrographs

depicting neurodegenerative changes in the brains of P8 rats who had been subjected to hyperoxia (80% O2) for 12 h on P7 and sacrificed at 24 h. In

micrographs A–D, silver-stained sections from thalamus (A and B), subiculum (C), and corpus callosum (D) from hyperoxic (A, C, and D) and normoxic (B)

rats are shown. Degenerating cells (small dark dots) are sparsely present in normoxic rat brain (B) but are abundantly present in hyperoxic brains (A, C, and D).

(F) Immunohistochemical staining for activated caspase 3 is shown in the laterodorsal thalamus of a rat subjected to hyperoxia. Many caspase-3-positive cells

are found (arrows). The magnified view of the framed area in G illustrates that immunopositive cells for activated caspase 3 show pycnotic changes in the

nuclei that are indicative of apoptosis. (E) Magnified view of a neuron from the parietal cortex of a rat subjected to hyperoxia (H&E staining). This neuron is

obviously degenerating and demonstrates nuclear fragmentation consistent with apoptosis. (H and I) Electron micrographs from cortical (H) and thalamic (I)

neurons in the brains of infant rats subjected to hyperoxia upon the end of a 12-h exposure. The neuron in H is in a middle stage of apoptosis and demonstrates

formation of spherical chromatin masses, intermixing of nucleoplasmic and cytoplasmic contents and condensation. The neuron in I is at a more advanced

stage. Magnifications: A and B �25; C �40; D and F �120; E �250; G �200; H and I original magnification �12,000.

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282 277

Fig. 2. Distribution pattern, time, and age dependency of hyperoxia-induced apoptosis in infant rats. (A) Schematic illustration of the distribution of apoptotic

cells in the brains of P7 rats subjected to 24 h of hyperoxia. The dotted areas are the ones affected. (B) Numerical densities of degenerating cells in 11 brain

regions in normoxic (white columns; n = 8) rats and rats subjected to 12 h (black columns; n = 8) or 48 h (shaded columns; n = 8) of hyperoxia. There are

significantly increased numerical densities of degenerating cells in all brain regions after both hyperoxic periods compared to normoxic rats. (C) Impact of the

duration of hyperoxia exposure on the severity of apoptotic cell death. P7 rats were exposed to 80% O2 for 2–72 h and sacrificed at either 24 h (applies to

2–24 h exposure periods) or upon end of the exposure (applies to 48 and 72 h exposures; n = 7–10/time point). The extent of apoptotic cell death was

quantified as described in methods. A 2-h exposure to an 80% O2 environment was sufficient to trigger significant apoptotic cell death compared to

normoxic littermates. Most severe degeneration was detected in the brains of rats subjected for 12 or 24 h to hyperoxia. No further increase could be

detected by this method after longer exposure times, most likely because apoptotic cells had already been eliminated. (D) Developmental vulnerability

profile to hyperoxia. P0–P14 rats (n = 7–10/group) were exposed to 80% O2 for 24 h and killed at the end of the exposure period. Vulnerability to

hyperoxia subsided by P14 in the rat (**P b 0.01; ***P b 0.001, Student’s t test, comparison hyperoxia group versus normoxia group).

Fig. 3. Hyperoxia induces oxidative stress in the infant rat brain. Protection with N-acetylcystein (N-Acc). Seven-day-old rats were exposed to 80% O2 over 6 h.

Upon end of the exposure, brain hemispheres were removed and shock frozen in liquid nitrogen. (A) Levels of reduced and oxidized glutathione (GSH and

GSSG), protein carbonyls, and the lipid peroxidation product malondialdehyde (MDA) were analyzed as described in Materials and methods. A significant

increase of GSSG and the GSSG/GSH ratio was detected in the brains of rats exposed to hyperoxia (n = 10) compared with those exposed to room air (n = 10).

Protein carbonyls showed a trend towards increase in the brains of rats exposed to hyperoxia. No significant change in malondialdehyde levels was detected

(*P b 0.05; **P b 0.01, Student’s t test, comparison hyperoxia group versus normoxia group). (B) P7 rats were treated with N-acetylcystein (N-Acc), 400 mg/

kg ip (n = 8), or vehicle (n = 8) before placement in the hyperoxia chamber (80% O2 for 12 h) and upon end of the exposure. Cell death was quantified at 24 h

on silver-stained sections as described in Materials and methods. Rats treated with N-acetylcystein had significantly lower apoptotic scores than vehicle treated

rats (**P b 0.01; Student’s t test).

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282278

4), which mediate intracellular signaling following activation of

receptor tyrosin kinases (Trk) by growth factors (Han and

Holtzman, 2000; Huang and Reichardt, 2001; Kaplan and Miller,

2000; Lee et al., 2001).

To confirm that impairment of the Ras-MAPK pathway is

causally involved in hyperoxia-induced cell death, we exposed

synRas-transgenic mice to 24 h of hyperoxia at the age of 6 days and

analyzed the brains after end of the exposure by means of

histological techniques for detecting cell death. Constitutively

activated V12-Ha-Ras is expressed selectively in neurons of

transgenic mice via a synapsin promotor (Heumann et al., 2000).

Ras-transgene protein expression increases postnatally and reaches a

four- to fivefold elevation at day 40 persisting at this level thereafter.

On P7, there are significantly 3.5-fold elevated levels of activated

V12-Ha-Ras in the cortex of transgenic mice compared to wild-type

controls (Figs. 5A and B). Toxic oxygen treatment suppressed Ras

activity by 50% in wild-type (wt) animals. In synRas mice,

hyperoxia-induced Ras attenuation was fully counteracted resulting

in still 2.5-fold elevated levels after treatment.

A corresponding activating phosphorylation of mitogen-acti-

vated protein kinase ERK1/2 is observed but no changes in the

activity of phosphatidylinositol-3 kinase (PI3-kinase), the phos-

phorylation of its target kinase Akt/PKB, or expression of the

antiapoptotic proteins Bcl-2 or Bcl-XL (Heumann et al., 2000).

Hyperoxia (80% O2 over 24 h) caused apoptotic cell death in the

brains of wild-type and synRas-transgenic mice with a distribution

pattern very similar to the one seen in P7 rats. Quantification of cell

death revealed that in the brains of synRas-transgenic mice, there

were significantly less degenerating cells compared to wild-type

age-matched littermates after 24 h of hyperoxia (Fig. 5C).

Discussion

Our results show that exposure of the developing rat and mouse

brain to high concentrations of oxygen for a period of hours during

a specific period of development causes an apoptotic neuro-

degenerative reaction that deletes large numbers of neurons from

several major regions of the developing forebrain. Hyperoxia-

induced cell death is disseminated in the brains and affects cortical

areas, the basal ganglia, hypothalamus, hippocampus, and white

matter tracts. Vulnerability to oxygen-induced cell death is age

dependent and maximal during the first week of life in the rat. This

developmental period coincides with the vulnerability period to the

proapoptotic effect of N-methyl-d-aspartate antagonists, GABAA

agonists, sodium channel blockers, and ethanol (Bittigau et al.,

2002; Ikonomidou et al., 1999, 2000; Jevtovic-Todorovic et al.,

2003).

Fig. 4. Impact of hyperoxia on survival promoting proteins. P7 rat pups were subjected to 80% O2 or normoxia over 2–24 h. Brain tissue from the thalamus was

dissected at the time points indicated. Decreased density of the BDNF, NGF, NT-3, and NT-4 specific bands, detected by PCR, is evident at 2, 6, 12, and 24 h

after the beginning of the hyperoxia (A). The density of the g-actin band is not affected. Immunoblotting was performed with anti-phospho-ERK 1/2, anti-

phospho-Akt, ERK 1/2, or Akt antibodies. There is a decrease in the levels of p-ERK 1/2 and p-Akt after hyperoxia, most evident at 12 h, whereas ERK 1/2 and

Akt levels (phosphorylation independent) remain unaffected. (B) Densitometric quantification of mRNA levels for BDNF, NGF, NT-3, and NT-4 in the

thalamus of P7 rats. Values represent mean normalized ratios of the neurotrophin bands to h-actin (n = 3–4/point F SEM). Analysis of variance (ANOVA)

revealed a significant effect of treatment with hyperoxia on BDNF [ F(1,15) = 24.34, P b 0.0001], NGF [ F(1,15) = 25.9, P b 0.0001], NT-3 [ F(1,15) = 5.438,

P b 0.0066], and NT-4 [ F(1,15) = 2.866, P b 0.0276] levels. (C) Densitometric quantification of protein levels for p-ERK 1/2 and p-Akt in the thalamus of P7

rats, analyzed by Western blotting. Values represent mean normalized ratios of the densities of the p-ERK1/2 and p-Akt bands compared to the density at 0 h

(n = 4/point F SEM). There was a significant effect of hyperoxia on p-ERK1/2 [ F(1,15) = 23.37, P b 0.0001] and p-Akt [ F(1,15) = 15.49, P b 0.0001]

levels (ANOVA). ERK1/2 and Akt levels were not affected.

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282 279

Our results reveal two mechanisms that are responsible for

hyperoxia-induced apoptotic death in the developing rodent brain.

The first one is oxidative stress. Hyperoxia-induced cell death is

associated with production of reactive oxygen species, as

evidenced by increased concentrations of oxidized glutathione

(GSSG) and a significantly elevated GSSG/GSH ratio.

These findings are in accordance with a previous report by

Taglialatela et al. (1998), who described that long exposure to 95%

oxygen over a period of 5 days, alone or in combination with a

glutathione synthesis inhibitor, increased DNA fragmentation in

the forebrain and lead to impairment of glutathione homeostasis. In

contrast to the study by Taglialatela et al. (1998), we show here that

much shorter exposures to lower levels of oxygen are sufficient to

compromise antioxidant protection systems in the developing brain

and lead to cell death.

It is well known that free radicals can lead to direct DNA

damage as well as damage of mitochondrial membranes, release of

cytochrome c into the cytoplasm, and activation of caspases

(Filomeni et al., 2003; Love, 1999; Yuan and Yarkner, 2000).

Furthermore, GSSG was shown to cause apoptosis in U937 cells

by activation of the p38 MAPK pathway (Filomeni et al., 2003).

Since N-acetylcystein ameliorated hyperoxia-induced apoptotic

cell death, we conclude that oxidative stress constitutes a

contributing factor.

Furthermore, we present evidence that hyperoxia exposure

leads to changes in gene expression and phosphorylation of

proteins that control neuronal survival during development.

Hyperoxia depressed synthesis of BDNF, NGF, NT-3, and NT-4

and reduced levels of the active forms of Ras, ERK1/2, and Akt in

a time-dependent fashion. Such changes reflect impairment of

survival promoting signals (Heumann, 1994) and an imbalance

between neuroprotective and neurodestructive mechanisms in the

brain, which, during a developmental period of ongoing physio-

logical elimination of brain cells, will likely promote apoptotic

death. The synRas-transgenic mice, which overexpress activated

Ras that counteracts hyperoxia-induced reduction in Ras activity

and display higher levels of phosphorylated ERK1/2 in the cortex,

were less susceptible to hyperoxia-induced cell death in the brain.

Since protection from hyperoxia-induced apoptosis was not

complete in synRas-transgenic mice, it is likely that mechanisms

other than reduced transcription of growth factors and impairment

of the extracellular signal regulated kinase (ERK1/2) pathway are

also contributing to hyperoxia-induced apoptotic cell death.

It has been known for decades that hyperoxia may induce

damage in immature lungs and retina. Oxygen toxicity has been

implicated in the pathogenesis of the neonatal diseases broncho-

pulmonary dysplasia and retinopathy of prematurity (Gibson et al.,

1990; Saugstad, 2001). Apoptosis has been identified as one form

of cell death occurring in the context of hyperoxia-induced organ

injury (Buckley et al., 1999; Okoye et al., 2003; Ray et al., 2003;

Yamada et al., 1999). Growth factors, activation of ERK1/2, and

activation of Akt can ameliorate hyperoxia-induced apoptotic cell

death in lungs and retina (Buckley et al., 1999; Lu et al., 2001;

Okoye et al., 2003; Ray et al., 2003; Yamada et al., 1999). A

Fig. 5. The synRas-transgenic mice are less sensitive to hyperoxia-induced apoptosis in the brain. Six-day-old synRas-transgenic and wild-type (wt) mice were

exposed to 80% O2 or normoxia for 24 h. After treatment, dissected brain samples were lysed and Ras activities were determined as described in the Materials

and methods section. (A) Blot of the GTP-Ras signals using anti-Ras in wt and synRas-transgenic mice. (B) Values represent mean ratios between GTP-bound

Ras to total Ras (4 wt, 4 synRas animals, 9 determinations,F SEM) normalized to 100% of untreated wt samples. Note the significant decrease in the levels of

activated Ras in the cortex of wt mice exposed to hyperoxia as compared to untreated controls (**P b 0.01). In synRas mice, the levels of GTP-Ras were 3.5-

fold elevated over wt. Despite hyperoxia-induced decreases in synRas mice, their levels of GTP-Ras exceeded those of wt mice after hyperoxia (***P b 0.001,

Student’s t test). (C) The synRas-transgenic and wild-type (wt) mice were exposed to hyperoxia for 24 h and killed upon end of the exposure. Quantification of

cell death in the brains was performed on TUNEL-stained sections as described in Materials and methods. Hyperoxia triggered significant apoptotic cell death

compared to unexposed littermates (dotted line) in both transgenic and wt mice. The synRas-transgenic mice developed significantly less degeneration

compared to wild-type mice after exposure to hyperoxia (n = 9/group). The level of physiological apoptosis was not different in wt and transgenic mice. Mean

apoptotic score for physiological apoptosis in wt and transgenic mice is depicted by the dotted line (*P b 0.05, Student’s t test).

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282280

pathogenetic role of hyperoxia-induced oxidative stress has also

been described (Lu et al., 2001). Thus, themechanisms we identified

as contributing to hyperoxia-induced apoptotic cell death in the

developing rodent brain are similar to those reported in other organs.

The vulnerability period to the proapoptotic effect of hyperoxia

coincides with the brain growth spurt period, which in the rat spans

the first two postnatal weeks of life. In humans, the comparable

period begins in the third trimester of gestation and extends to

several years after birth (Rice et al., 1981). The injury caused by

exposure to high levels of oxygen in the infant rat and mouse brain

does not resemble the bclassic large cystic lesionsQ of other animal

models of periventricular leukomalacia and white matter injury

(Redecker et al., 1998; Rice et al., 1981) since it is more diffuse. In

an early magnetic resonance imaging study in premature infants,

diffuse brain lesions (bdiffuse excessive high signal intensityQ)have been detected, which do not develop into cystic periven-

tricular leukomalacia later on (Maalouf et al., 1999). Interestingly,

in a recent study, hyperoxia could be identified as one risk factor

for abnormal neurologic outcome of preterm infants (Collins et al.,

2001). Thus, the clinical data in combination with the experimental

evidence presented here suggest that apoptotic neurodegeneration

triggered by a nonphysiological, high oxygen environment during

a critical stage of development may partly account for cognitive

and also motor impairment of premature infants. A complicating

factor to be considered is the fact that, in contrast to the infant rat,

premature human infants are already subjected to unphysiologi-

cally high oxygen concentrations compared to intrauterine con-

ditions when exposed just to ambient air. Magnetic resonance

spectroscopy studies of the brain demonstrate that cerebral

metabolism is different in preterm infants and that cerebral lactate

can be detected under normal conditions (Penrice et al., 1997;

Robertson et al., 2000). In addition, antioxidant defense mecha-

nisms in the brain of premature infants are very immature (Nishida

et al., 1994). Thus, the preterm human brain may be even more

susceptible than the rodent brain to mechanisms capable of

inducing oxidative stress, such as hyperoxia. We recommend that

every effort should be made in neonatal medicine to limit exposure

of these young patients to high oxygen concentrations.

Acknowledgments

This work was supported by grants from the Bundesministe-

rium fqr Bildung und Forschung (BMBF) 01ZZ0101 and

01GZ0305 and a grant from the Humboldt University.

References

Abernethy, L.J., Palaniappan, M., Cooke, R.W.I., 2002. Quantitative

magnetic resonance imaging in the brain in survivors of very low birth

weight. Arch. Dis. Child. 87, 279–283.

Ajayi-Obe, M., Saeed, N., Cowan, F.M., Rutherford, M.A., Edwards, A.D.,

2000. Reduced development of cerebral cortex in extremely preterm

infants. Lancet 356, 1162–1163.

Beutler, E., Duron, O., Kelly, B.M., 1963. Improved method for the

determination of blood glutathione. J. Lab. Clin. Med. 61, 882–888.

Bittigau, P., Sifringer, M., Genz, K., Reith, E., Pospischil, D., Govindarajalu,

S., Dzietko, M., Pesditschek, S., Mai, I., Dikranian, K., Olney, J.W.,

Ikonomidou, C., 2002. Antiepileptic drugs and apoptotic neurodege-

neration in the developing brain. Proc. Natl. Acad. Sci. U. S. A. 99,

15089–15094.

Buckley, S., Driscoll, B., Barsky, L., Weinberg, K., Anderson, K.,

Warburton, D., 1999. ERK activation protects against DNA damage

and apoptosis on hyperoxic rat AEC2. Am. J. Physiol., L159–L166.

Buss, H., Chan, T.P., Sluis, K.B., Domigan, N.M., Winterbourne, C.C.,

1997. Protein carbonyl measurement by a sensitive ELISA method.

Free Radical Biol. Med. 23, 361–366.

Collins, M.P., Lorenz, J.M., Jetton, J.R., Paneth, N., 2001. Hypocapnia and

other ventilation-related risk factors for cerebral palsy in low birth

weight infants. Pediatr. Res. 50, 712–719.

DeOlmos, J.S., Ingram, W.R., 1971. An improved cupric-silver method

for impregnation of axonal and terminal degeneration. Brain Res. 33,

523–529.

Dobbing, J., Sands, J., 1979. Comparative aspects of the brain growth spurt.

Early Hum. Dev. 3, 79–83.

Filomeni, G., Rotilio, G., Ciriolo, M.R., 2003. Glutathione disulfide

induces apoptosis in U937 cells by a redox-mediated p38 MAP kinase

pathway. FASEB J. 17, 64–66.

Gibson, D.L., Sheps, S.B., Uh, S.H., Schechter, M.T., McCormick, A.Q.,

1990. Retinopathy of prematurity-induced blindness: birth weight-

specific survival and the new epidemic. Pediatrics 86, 405–412.

Hack, M., Flannery, D.J., Schluchter, M., Cartar, L., Borawski, E., Klein,

N., 2002. Outcomes in young adulthood for very-low-birth-weight

infants. N. Engl. J. Med. 348, 149–157.

Han, B.H., Holtzman, D.M., 2000. BDNF protects the neonatal brain from

hypoxic-ischemic injury in vivo via the ERK pathway. J. Neurosci. 20,

5775–5781.

Heumann, R., 1994. Neurotrophin signalling. Curr. Opin. Neurobiol. 4,

668–679.

Heumann, R., Goemans, C., Bartsch, D., Lingenhohl, K., Waldmeier, P.C.,

Hengerer, B., Allegrini, P.R., Schellander, K., Wagner, E.F., Arendt, T.,

2000. Transgenic activation of ras in neurons promotes hypertrophy

and protects from lesion-induced degeneration. J. Cell Biol. 151,

1537–1548.

Hissin, P.J., Hilf, R., 1976. A fluorometric method for determination of

oxidized and reduced glutathione. Anal. Biochem. 74, 77–82.

Huang, E.J., Reichardt, L., 2001. Neurotrophins: roles in neuronal

development and function. Annu. Rev. Neurosci. 24, 677–736.

Ikonomidou, C., Bosch, F., Miksa, M., Vfckler, J., Bittigau, P., Dikranian,K., Tenkova, T., Turski, L., Olney, J.W., 1999. Blockade of NMDA

receptors and apoptotic neurodegeneration in the developing brain.

Science 283, 70–74.

Ikonomidou, C., Bittigau, P., Ishimaru, M.J., Wozniak, D.F., Koch, C.,

Genz, K., Price, M.T., Stefovska, V., Hfrster, F., Tenkova, T.,

Dikranian, K., Olney, J.W., 2000. Ethanol-induced apoptotic neuro-

degeneration and fetal alcohol syndrome. Science 287, 1056–1060.

Jevtovic-Todorovic, V., Hartman, R.E., Izumi, Y., Benshoff, N.D.,

Dikranian, K., Zorumski, C.F., Olney, J.W., Wozniak, D.F., 2003.

Early exposure to common anesthetic agents causes widespread

neurodegeneration in the developing rat brain and persistent learning

deficits. J. Neurosci. 23, 876–882.

Kaplan, D.R., Miller, F.D., 2000. Neurotrophin signal transduction in the

nervous system. Curr. Opin. Neurobiol. 10, 381–391.

Lee, R., Kermani, P., Teng, K.K., Hempstead, B.L., 2001. Regulation of

cell survival by secreted proneurotrophins. Science 294, 1945–1948.

Lohmann, J., Schickle, H., Bosch, T.C., 1995. REN display, a rapid and

efficient method for nonradioactive differential display and mRNA

isolation. Biotechniques 18, 200–202.

Love, S., 1999. Oxidative stress in brain ischemia. Brain Pathol. 9,

119–131.

Lu, Y., Parkyn, L., Otterbeit, L.E., Kureishi, Y., Walsh, K., Ray, A., Ray, P.,

2001. Activated Akt protects the lung from oxidant-induced injury and

delays death of mice. J. Exp. Med. 193, 545–549.

Maalouf, E.F., Duggan, P.J., Rutherford, M.A., Counsell, S.J., Fletcher,

A.M., Battin, M., Cowan, F., Edwards, A.D., 1999. Magnetic resonance

imaging of the brain in a cohort of extremely preterm infants. J. Pediatr.

135, 351–357.

Ment, L.R., Vohr, B., Allan, W., Katz, K.H., Schneider, K.C., Westerveld,

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282 281

P.A., Duncan, C.C., Makuch, R.W., 2000. Change in cognitive function

over time in very-low-birth-weight infants. JAMA 289, 705–711.

Murphy, D.J., Sellers, S., MacKenzie, I.Z., Yudkin, P.L., Johnson, A.M.,

1995. Case-control study of antenatal and intrapartum risk factors for

cerebral palsy in very preterm singleton babies. Lancet 346, 1449–1454.

Nishida, A., Misaki, Y., Kuruta, H., Takashima, S., 1994. Developmental

expression of copper, zinc-superoxide dismutase in human brain by

chemiluminescence. Brain Dev. 16, 40–43.

Nosarti, C., Al-Asady, M.H.S., Frangou, S., Stewart, A.L., Rifkin, L.,

Murray, R.-M., 2002. Adolescents who were born very preterm have

decreased brain volumes. Brain 125, 1616–1623.

Okoye, G., Zimmer, J., Sung, J., Gehlbach, P., Deering, T., Nambu, H.,

Hackett, S., Melia, M., Esumi, N., Zack, D.J., Campochiaro, P.A., 2003.

Increased expression of brain-derived neurotrophic factor preserves

retinal function and slows cell death from rhodopsin mutation or

oxidative damage. J. Neurosci. 23, 4164–4172.

Olney, J.W., Wozniak, D.F., Jevtovic-Todorovic, V., Farber, N.B., Bittigau,

P., Ikonomidou, C., 2002. Drug-induced apoptotic neurodegeneration in

the developing brain. Brain Pathol. 12, 488–498.

Penrice, J., Lorek, A., Cady, E.B., Amess, P.N., Wylezinska, M., Cooper,

C.E., D’Souza, P., Brown, G.C., Kirkbride, V., Edwards, A.D., 1997.

Proton magnetic resonance spectroscopy of the brain in normal preterm

and term infants, and early changes after perinatal hypoxia-ischemia.

Pediatr. Res. 40, 6–14.

Ray, P., Devaux, Y., Stolz, D.B., Yarlagaffa, M., Watkins, S.C., Lu, Y., Chen,

L., Yang, X.F., Ray, A., 2003. Inducible expression of keratinocyte

growth factor (KGF) in mice inhibits lung epithelial cell death induced

by hyperoxia. Proc. Natl. Acad. Sci. U. S. A. 100, 6098–6103.

Redecker, C., Hagemann, G., Witte, O.W., Marret, S., Evrard, P., Gressens,

P., 1998. Long-term evolution of excitotoxic cortical dysgenesis induced

in the developing rat brain. Brain Res. Dev. Brain Res. 109, 109–113.

Rice, J.E., Vannucci, R.C., Brierley, J.B., 1981. The influence of immaturity

on hypoxic-ischemic brain damage in the rat. Ann. Neurol. 9, 131–141.

Robertson, N.J., Kuintm, J., Counsell, T.J., Rutherford, T.A., Coutts, A.,

Cox, I.J., Edwards, A.D., 2000. Characterization of cerebral white

matter damage in preterm infants using 1H and 31P magnetic resonance

spectroscopy. J. Cereb. Blood Flow Metab. 20, 1446–1456.

Saugstad, O.D., 2001. Chronic lung disease: oxygen dogma revisited. Acta

Paediatr. 90, 113–115.

Sitte, N., Merker, K., Grune, T., 1998. Proteasome-dependent degrada-

tion of oxidized proteins in MRC-5 fibroblasts. FEBBS Lett. 440,

399–402.

Sommerburg, O., Grune, T., Klee, S., Ungemach, F.R., Siems, W.G., 1993.

Formation of 4-hydroxynonenal and further aldehydic mediators of

inflammation during bromotrochloromethane treatment of liver cells.

Mediat. Inflamm. 2, 27–31.

Taglialatela, G., Perez-Polo, J.R., Rassin, D.K., 1998. Induction of

apoptosis in the CNS during development by the combination of

hyperoxia and inhibition of glutathione synthesis. Free Radical Biol.

Med. 8, 936–942.

Taylor, D.L., Edwards, D., Mehmet, H., 1999. Oxidative metabolism,

apoptosis and perinatal brain injury. Brain Pathol. 9, 93–117.

Vohr, B.R., Wright, L.L., Dusick, A.M., Mele, L., Verter, J., Steichen, J.J.,

Simon, N.P., Wilson, D.C., Broyles, S., Bauer, C.R., 2000. Neuro-

developmental and functional outcomes of extremely low birth weight

infants in the National Institute of Child Health and Human

Development Neonatal Research Network 1993–1994. Pediatrics

105, 1216–1226.

West, M.J., Gundersen, H.J.G., 1990. Unbiased stereological estimation of

the number of neurons in the human hippocampus. J. Comp. Neurol.

296, 1–22.

Wong, S.H.J., Knight, J.A., Hopfer, S.M., Zaharia, O., Leach Jr., C.N.,

Sunderman Jr., F.W., 1987. Lipoperoxides in plasma as measured by

liquid-chromatographic separation of malondialdehyde-thiobarbituric

acid adduct. Clin. Chem. 33, 214–220.

Wood, N.S., Marlow, N., Costeloe, K., Gibson, A.T., Wilkinson, A.R.,

2000. Developmental disability after extremely preterm birth. N. Engl.

J. Med. 343, 378–384.

Yamada, H., Yamada, E., Hackett, S.F., Ozaki, H., Okamoto, N.,

Campochiaro, P.A., 1999. Hyperoxia causes decreased expression of

vascular endothelial growth factor and endothelial cell apoptosis in

adult retina. J. Cell Physiol. 179, 149–156.

Yuan, J., Yarkner, B.A., 2000. Apoptosis in the nervous system. Nature

407, 802–809.

U. Felderhoff-Mueser et al. / Neurobiology of Disease 17 (2004) 273–282282