Impaired neurological development in premature infants frequently arises from periventricular white matter injury (PWMI), a condition
associated with myelination abnormalities. Recently, exposure to hyperoxia was reported to disrupt myelin formation in neonatal rats. To
exposed to 48 h of 80% oxygen from postnatal day 6 (P6) to P8. Myelin basic protein expression and CC1?oligodendroglia decreased after
affect survival or proliferation of astrocytes in vivo, but modified GFAP and glutamate-aspartate transporter expression. The rate of [3H]-D-
nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7-sulfonamide treatment. Our analysis reveals a role for altered glutamate homeostasis in
The most common pathology observed in preterm infants is
periventricular white matter injury (PWMI), a disorder that has
been associated with myelination disturbances (Cheong et al.,
(Back, 2006; Deng et al., 2008; Khwaja and Volpe, 2008), and
neuronal/axonal disease (Pierson et al., 2007; Volpe, 2009).
Therefore, children born preterm are more likely to develop fu-
ture motor and cognitive deficits (Aylward, 2002; Bhutta et al.,
2002; Wocadlo and Rieger, 2008).
Causes associated with PWMI include perinatal infection and
use of high oxygen in preterm infants can also lead to poor neuro-
birth, preterm infants experience a several-fold increase in arterial
oxygen tension to 65–80 mmHg (Hoffmann, 2002), even without
In an experimental rat model, exposure to 80% oxygen at
postnatal day 7 (P7) resulted in cell death in the gray matter and
subcortical white matter (WM) that is not observed with expo-
sure at later postnatal ages (Felderhoff-Mueser et al., 2004). Al-
though cellular changes were not characterized in these studies,
cell death in the brain was associated with increased intracranial
expression of proinflammatory cytokines (Felderhoff-Mueser et
80% oxygen caused caspase-dependent cell death in cultured
O4?O1?preoligodendrocytes (pre-OLs), but not in mature
O4?O1?MBP?oligodendrocytes (Gerstner et al., 2008). These
findings indicate that high levels of oxygen cause oligodendroglial
Glutamate-mediated excitotoxicity has been shown to cause
extensive damage to the developing brain in animal models of
TheJournalofNeuroscience,March16,2011 • 31(11):4327–4344 • 4327
hypoxia/ischemia (Silverstein et al., 1986) and trauma (Bittigau
et al., 1999). In addition, the overactivation of non-NMDA
glutamate receptors (GluRs) leads to oligodendroglial cell
death (Follett et al., 2000; Deng et al., 2004), decreased oligo-
dendrocyte progenitor cell (OPC) proliferation, and attenu-
ated oligodendroglia lineage progression (Gallo et al., 1996;
Yuan et al., 1998). Vesicular release of glutamate within the
developing WM occurs from both myelinated (Bezzi et al.,
al., 2004). Furthermore, the uptake of glutamate through
membrane transporters is critical for its extracellular clear-
ance (Bergles et al., 1999; Danbolt, 2001). Glial transporters,
particularly in astrocytes, are believed to perform the majority
2005), and altered transport has been demonstrated in experi-
mental models of neural dysfunction (Zugno et al., 2007) and
Down syndrome (Begni et al., 2003).
In this study, we sought to characterize hyperoxia-induced
cellular changes within the developing WM and investigate the
role of astrocytes and astrocyte-mediated glutamate uptake as
potential contributors to oligodendroglial damage and altered
protein (GFAP) promoter were generated on an FVB/N background
(from Dr. Frank Kirchhoff, Max Planck Institute of Experimental Med-
of GFAP-positive astrocytes, depending on age and brain region (Weh-
ner et al., 2003). In our studies on external capsule (EC) and cingulum
(CG), we found an overlap of glutamine synthetase (GS) immunostain-
ing and EGFP expression in 50% of astrocytes at P8 and 80% at P12.
Transgenic mice expressing EGFP under the control of the 2–3-cyclic
nucleotide 3-phosphodiesterase (CNP) promoter have been described
previously (Yuan et al., 2002). CNP-EGFP expression is detected in cells
of the oligodendroglial lineage at early embryonic stages of develop-
ment, and this expression is maintained throughout brain matura-
tion. In accordance, we detected GFP?oligodendroglia within the
subcortical white matter and cortex of CNP-EGFP transgenic mice
throughout development. No obvious differences were observed in
brain or white matter size within either type of transgenic mouse. All
procedures were performed according to the Institutional Animal
Care and Use Committee, Children’s National Medical Center, and
National Institutes of Health guidelines.
mice, and CNP-EGFP transgenic mice were subjected to hyperoxia and
subsequently used for immunohistology, with only C57BL/6 wild-type
and female neonatal mice were divided into hyperoxia and control
in a chamber containing 80% O2for 6 or 48 h. The control pups of each
litter were kept in room air with a second lactating mother. The mothers
of the two groups were replaced after 24 h to prevent oxygen-induced
acute lung injury (Taglialatela et al., 1998). During recovery in room air,
until being killed. The pups appeared normal and did not suffer weight
loss (control, P7, 4.14 ? 0.24 g; P8, 5.31 ? 0.26 g; P12, 6.83 ? 0.29 g;
hyperoxia, P7, 4.26 ? 0.21 g; P8, 5.22 ? 0.11 g; P12, 6.76 ? 0.31 g) or
changes in body temperature (control, P7, 30.65 ? 1.11°C; P8, 30.07 ?
0.37°C; P12, 31.10 ? 0.49°C; n ? 6 for each time point) during or after
the hyperoxia exposure. Body temperature was monitored by using a
Micro Therma 2T device with a neonatal mouse (RET-4) rectal probe
abolic panel were obtained in both experimental groups at P8. Animals ex-
sample. pH, pO2, pCO2, Na?, K?, Ca2?, Cl?, glucose, and lactate were
Bromodeoxyuridine injections. Acute bromodeoxyuridine (BrdU) in-
intraperitoneal injection of 10 mg/kg BrdU in both hyperoxia-exposed
and litter-matched control mice 2 h before they were killed. For BrdU
analysis at P8, pups undergoing hyperoxia exposure where administered
the BrdU injection within the oxygen chamber to prevent room-air ex-
posure. During further analysis of BrdU incorporation after recovery in
of BrdU 2 h before they were anaesthetized and subjected to transcardial
recovery in room air from P8 to P12, both experimental groups were
subjected to a cumulative (pulse-chase) BrdU protocol where the ani-
mals were administered repeat intraperitoneal injections of 10 mg/kg
ing for 4 d until P12. At P12, all mice were killed and transcardially
tized following National Institutes of Health guidelines and transcardi-
ally perfused with PBS and then 4% paraformaldehyde (PFA). Brains
were dissected out and postfixed with 4% PFA overnight at 4°C. Fixed
brains were preserved in 10% glycerol in PBS. For tissue sections, brains
were rinsed in 1? PBS and then frozen in tissue freezing medium (Tri-
angle Biomedical Sciences) and mounted on a Microm HM400 mi-
crotome (Microm International). Sections were cut (30–40 ?m) and
stored in a 1? PBS, 0.05% sodium azide solution. For immunohisto-
chemistry, sections were blocked at room temperature for at least 1 h in
in 1? PBS). Polyclonal rabbit antibody to NG2 chondroitin sulfate pro-
teoglycan (Millipore Bioscience Research Reagents) was diluted 1:400.
antibody to GFAP (Abcam) were diluted 1:500. The monoclonal rabbit
Ki67 antibody (Vector Laboratories) was diluted 1:250. Monoclonal
mouse BrdU antibody was diluted 1:500 (Axyll). Tissue analyzed for
BrdU incorporation was pretreated with 2N HCl for 30 min followed by
0.1 M boric acid for 15 min at room temperature. Monoclonal rabbit
Olig2 antibody and the monoclonal mouse CC1 antibody were diluted
1:500 (Abcam; Calbiochem). Polyclonal guinea pig antibodies for
glutamate-aspartate transporter (GLAST)/excitatory amino acid trans-
porter 1 (EAAT1) (Millipore Life Science) and glutamate transporter-1
(GLT-1)/EAAT2 (Millipore Life Science) were diluted to 1:500. Poly-
clonal rabbit antibody for GS (Abcam) was diluted to 1:500. Brain sec-
solution every 10 min. All secondary antibodies used were from Jackson
anti-mouse IgG (1:200), FITC-conjugated goat anti-rabbit IgG (1:200),
FITC-conjugated donkey anti-chicken IgG (1:500); CY5-conjugated
goat anti-mouse IgG (1:500), CY5-conjugated goat anti-rabbit IgG (1:
500), CY5-conjugated goat anti-rat (1:500), and CY5-conjugated goat
anti-guinea pig; CY3/Rhodamine-conjugated goat anti-mouse IgG (1:
200), CY3/Rhodamine-conjugated goat anti-rabbit IgG (1:200), and
CY3/Rhodamine-conjugated goat anti-guinea pig (1:200). Incubation
was performed at room temperature for 1 h followed by three washes as
described above. Sections were then stained with 4?,6-diamidino-2-
phenylindole (DAPI) for 10 min and, after three washings with PBS,
mounted with Mowiol.
onic day 19 pregnant Sprague Dawley rats by mechanical dissociation
4328 • J.Neurosci.,March16,2011 • 31(11):4327–4344Schmitzetal.•HyperoxiaandWhiteMatterDevelopment
according to the method of McCarthy and de Vellis (1980) as described
previously (Gallo and Armstrong, 1995; Gallo et al., 1996). Mixed cul-
tures (7–10 d old) were shaken overnight to detach OPCs from the as-
trocyte monolayer. To minimize contamination by microglial cells, the
detached cell suspension was incubated in succession for 45 min each in
60 mm dishes. OPCs enriched by this method contained ?95% GD31
cells labeled by the LB1 monoclonal antibody (Levi et al., 1986; Curtis et
al., 1988), with ?0.05% GFAP?astrocytes and ?0.05% Ox42?micro-
glia. Attached astrocytes were passaged after overnight shaking and
transferred into T75 culture flasks (BD Falcon) at a density of 2 ? 106
cells per flask with 10 ml of DMEM (Invitrogen) containing 10% FCS,
which was changed after overnight incubation to remove nonattached
OPCs. Once cells became confluent, flasks were shaken overnight and
media containing nonattached OPCs and microglia was removed. For
immunocytochemistry, astrocyte layers were trypsinized and plated on
poly-lysine-coated (0.1 mg/ml) 25 mm coverslips in 35 mm dishes using
days. For Western blot protein analysis, astrocytes, after trypsinization,
of DMEM with 10% FCS until confluent. For subconfluent astrocyte
cultures, 1 ? 105cells in 2 ml of DMEM containing 10% FCS were
transferred to six-well plates and cultured for 48 h, and media were
changed the day after trypsinization. The six-well plates were then used
for experiments at either 80 or 21% O2.
Immunocytochemistry. Live staining for cell surface antigens with
A2B5, O4, and O1 antibodies (Bansal et al., 1989) was performed as
at room temperature for 1 h with primary antibodies diluted 1:10 in
DMEM followed by fluorescein-conjugated goat anti-mouse IgM for 45
(pH 7.3 in PBS) for 10 min at room temperature and washed in PBS.
Coverslips were then mounted in DAPI-containing Vectashield. For
double staining with GFAP, cells, after live staining, fixation, and wash-
ing, were blocked again in 10% NGS in DMEM containing 0.1% Triton
X-100 for permeabilization for 20 min at room temperature. Incubation
with GFAP mouse antibody (1:500; Sigma-Aldrich) followed for 1 h at
room temperature. After washing, the cells were incubated with Rhoda-
mine anti-mouse IgG antibodies (1:200; Jackson Immunoresearch Lab-
oratories). The cells were then washed and mounted in Vectashield with
Terminal deoxynucleotidyl transferase-mediated biotinylated UTP nick
end labeling assays. Terminal deoxynucleotidyl transferase-mediated bi-
otinylated UTP nick end labeling (TUNEL) assays were performed ac-
cording to the manufacturer’s directions (In Situ Cell Death Detection
Kit, Rhodamine; Roche Applied Science). TUNEL immunostaining was
performed on tissue obtained after hyperoxia or from animals kept in
room air (control). Tissue was then processed for NG2 immunolabeling
as described above (Immunofluorescence). Sections were then perme-
1? PBS, tissue was mounted on slides, allowed to dry, and placed in
Vectashield with DAPI. Cells grown on 25 mm coverslips were fixed in
paraformaldehyde for 10 min at room temperature and then rinsed in
PBS. Cells were then permeabilized for 2 min on ice before labeling with
humidified chamber under Parafilm on coverslips. After washing with
PBS, paraformaldehyde fixation was followed by treatment of cells with
0.07 NaOH in PBS for 10 min at room temperature. After washing, cells
were fixed again for 10 min and permeabilized in 0.1% Triton X-100 in
PBS for 1 min. After washing, cells were incubated with 10% goat serum
for 15 min, followed by monoclonal anti-GFAP antibodies (1:500;
Sigma-Aldrich) for 1 h at room temperature, followed by fluorescein-
conjugated goat anti-mouse IgG (1:200; Jackson Immunoresearch Lab-
oratories) for for 30 min at room temperature. Coverslips were then
mounted in DAPI-containing Vectashield.
3-(4,5-Dimethyl thiazol-2-yl)-2,5-diphenyl tetrazolium bromide assay.
DMEM containing 10% FCS, measurements of reduction of 3-(4,5-
dimethyl thiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) were
performed according to the manufacturer’s directions (TACS MTT As-
says; R & D Systems). MTT reagent (100 ?l) was added to each well of a
12-well plate containing 1 ml of growth media during the final 4 h of
incubation. Detergent reagent (300 ?l) was then added to each well to
solubilize the dark blue crystals overnight. Supernatants (250 ?l) were
finally transferred to 96-well plates and read on a Molecular Devices
ThermoMax 96-well plate reader, using a test wavelength of 570 nm.
Western blotting. Protein analysis was only conducted in WT mice.
Microdissected tissue of the WM, including corpus callosum (CC), cin-
cipitation assay (RIPA) buffer solution for protein extraction. For
30 min. Aliquots were then assayed for protein concentration using the
Pierce BCA kit with a 30 min incubation at 60°C before spectrophotom-
etry at 590 nm. Total proteins were equally loaded (10–40 ?g per lane)
on 4–20% mini precast Tris-glycine gels. The gels were transferred onto
polyvinylidene fluoride membranes at 4°C overnight and blocked in 4%
ies were diluted 1:500 to 1:2000 in 4% milk. Horseradish-peroxidase-
conjugated secondary antibodies (anti-rabbit and anti-mouse, BD
Biosciences PharMingen; anti-goat, Santa Cruz Biotechnology; anti-guinea
in TBST. Chemiluminescent detection was performed using ECL Plus (GE
ers’ directions. The antibodies used were as follows: monoclonal mouse
polyclonal rabbit GLAST/EAAT1, 1:250 (Abcam); polyclonal guinea pig
GLT-1/EAAT2, 1:250 (Millipore Bioscience Research Reagents); monoclo-
Microscopy and cell density measurements. A Zeiss LSM 510 confocal
laser scanning microscopic system was used for the analysis of fluores-
cence after immunohistochemical staining in wild-type, CNP-EGFP,
sion filter), Cy5 (647 nm excitation; 680/32 emission filter), and DAPI
(400 nm excitation). Data acquisition and processing were controlled by
LSM software. Analysis of immunofluoresence was performed on con-
taken from the CG and EC regions depicted in supplemental Figure 1
(available at www.jneurosci.org as supplemental material) within two to
three sections for each animal analyzed. Confocal Assistant 4.02 and
Image J (NIH) software were used to merge images for analysis. Merged
images were processed in Photoshop 7.0 with minimal manipulation of
contrast. Cells were counted in a blinded fashion and double or triple
labeled by analyzing the merged image for each confocal z-stack and
identifying positive immunofluorescence for each individual channel.
Exposure of primary astrocytes to hyperoxia. At the beginning of the
experiment, cultures were placed in six-well plates using 2 ml of DMEM
growth media per well as mentioned and transferred to a humidified
O2using a gas tank containing 80% O2, 5% CO2, and 15% N2(Roberts
L/min for 10 min, in accordance with the manufacturer’s instructions.
for control were kept under 21% O2, 5% CO2, and 15% N2at 37°C.
Astrocyte–OPC cocultures. Monolayers of confluent astrocytes were
obtained on coverslips, as described previously. Media were then re-
newed with DMEM without FCS containing 100 ?M dibutyryl (db)-
cAMP (Sigma-Aldrich) and exposed to a mixture of 80% O2, 5% CO2,
briefly before the OPC suspensions were prepared for plating. For con-
trol, dishes containing astrocytes on coverslips were kept at 21% O2, 5%
the mixed glia cell cultures, media were changed after shaking for 2 h.
After an overnight shake for 12–16 h, OPCs were harvested and trans-
Schmitzetal.•HyperoxiaandWhiteMatterDevelopmentJ.Neurosci.,March16,2011 • 31(11):4327–4344 • 4329
ferred to 15 ml tubes for centrifugation at 1500 rpm for 5 min. Resus-
pended cells were transferred to a 60 mm culture dish and incubated for
45 min at 37°C for attachment of microglia. OPCs were removed by
ml tube and centrifuged for 5 min at 1500 rpm.
ml of DMEM containing the OPCs to be plated. After incubation for 45
min at 37°C, the tube was centrifuged for 5 min at 1500 rpm and media
was aspirated. Pellets were resuspended in 2 ml of DMEM containing
incubation to allow OPC attachment, media was carefully aspirated and
replaced by 2 ml of DMEM without FCS for each dish/coverslip. For
glutamate challenge, 40 ?l of glutamate 50 mM was added to the dish to
obtain a final glutamate concentration of 1 mM/2 ml culture media.
For protection experiments, the non-NMDA-receptor antagonist 2,3-
100 ?M; Sigma-Aldrich) was added to the culture media 1 h before glu-
A2B5?/Dye?cells, and for 48 h for O4?/Dye?cells.
Measurement of [3H]-D-aspartic acid uptake by cultured astrocytes. Pri-
mary astrocytes cultures were grown on 24-well plates until confluent.
Media was changed to DMEM without FCS containing db-cAMP, and
hyperoxia experiments were performed as described previously. Plates,
after 48 h exposure to either 80 or 21% O2, were placed at the surface of
a 37°C water bath, and rinsed twice with 1 ml of preheated Krebs’ buffer
transportable analog D-aspartic acid, which is not metabolized and does
not interact with any form of glutamate receptors. [3H]-D-aspartic acid
(50 nM; PerkinElmer) was diluted with unlabeled D-aspartic acid to
uptake was stopped after 10 min by three rinses with ice-cold sodium-
free Krebs’ buffer in which NaCl was replaced by chloride of the same
and the radioactivity of 200 ?l of the lysate was determined by liquid
scintillation counting. A fraction of the lysate was also used for protein
determination. To correct for nonspecific [3H]-D-aspartic acid uptake,
we subtracted data obtained under conditions using Na-free Krebs’ buf-
fer. All data are expressed as the rate of Na-dependent uptake per milli-
gram of protein per minute. For conversion of scintillation counts into
uptake assays at all time points (P8, P12, P15, and P30) were performed
using WM tissue isolated by microdissection and a modified assay pro-
tocol (Weller et al., 2008). Brains were removed and placed in D1 solu-
tion (1? HBSS, containing 6% glucose, 15% sucrose, and 1% PenStrep)
removed and placed in 1 ml of homogenizing buffer (50 mM Tris, 0.3 M
sucrose, pH 7.4) on ice. After homogenization, suspension was centri-
fuged at 13,500 rpm for 10 min at 4°C. Pellet was then resuspended in
KRH (Krebs’ Ringer’s HEPES solution). Duplicate 200 ?l assays were
performed for each experimental condition. Assays containing 100 ?M
dihydrokainic acid (DHK; Sigma-Aldrich) and 75 ?M DL-threo-b-
benzyloxyaspartic acid (TBOA); Tocris Bioscience) were incubated at
37°C for 10 min. After each respective agent was added to the sample, 50
nM [3H]-D-aspartic acid (PerkinElmer) was placed in each sample and
allowed to incubate for 5 min at 37°C. Preparations were then centri-
ing the pellet in sodium-free KRH minus Na (sodium-free Krebs’
solution containing the same molar concentration of “normal” Krebs’
solution but with choline chloride in place of sodium chloride) twice to
stop additional sodium-dependent [3H]-D-aspartic acid uptake. Cells in
added to Fischer Scientific ScintiVerse scintillation fluid and allowed to
Bradford assay using the remaining 50 ?l of lysate. [3H]-D-aspartic acid
uptake was determined using a Beckman LS 6500 scintillation counter
and calculated as picomoles per milligram of protein per minute. Values
obtained from samples containing TBOA were subtracted from the total
and non-GLT values to show sodium-dependent [3H]-D-aspartic acid
uptake for each respective sample in all experiments.
Magnetic resonance imaging and diffusion tensor imaging. Magnetic
resonance imaging (MRI) was performed using a 7 tesla rodent scanner
(Pharmascan 70/16; Bruker BioSpin) with a 16 cm horizontal bore mag-
net and a 9 cm (inner diameter) shielded gradient with an H-resonance
frequency of 300 MHz and a maximum gradient strength of 300 mT/m.
For imaging, a1H-phased-array surface coil for mouse head and a1H-
radio frequency-volume resonator (72 mm) for transmission were used.
Data acquisition and image processing were performed with the Bruker
software Paravision 4.0. During the examinations, mice were anesthe-
tized with 2.0–1.0% isoflurane (Forene; Abbot) delivered in a O2/N2O
ing and Gating System; SA Instruments). To ensure physiological body
temperature, animals were placed on a heated circulating water blanket.
To localize the CC, we first used a T2-weighted two-dimensional turbo
spin-echo sequence [repetition time (TR), 4200 ms; echo time (TE), 36
cm and a matrix size of 256 ? 256 with 20 slices at 0.5 mm to cover the
whole brain. For subsequent diffusion tensor imaging (DTI), a 1 mm
slice with an FOV of 2.60 ? 2.60 cm and a matrix size of 128 ? 128 was
placed over the CC. Imaging parameters are as follows: DTI echo-planar
s/mm2; diffusion duration, ? ? 4 ms; diffusion separation, ? ? 20 ms;
TR, 3000 ms, TE, 34 ms; one average. If there were movement artifacts,
the whole data set of the animal was excluded from analysis. On a pixel-
by-pixel basis, fractional anisotropy (FA) as well as radial and axial ap-
parent diffusion coefficients (ADCs; determined perpendicularly and
CC fibers. In addition, directionally encoded color (DEC) maps were
used to represent anisotropy in three directional manners in coronal
images: red for lateral–medial, blue for anterior–posterior, green for in–
out (Pajevic and Pierpaoli, 1999). Given the axonal directory in the CC,
test) of FA and ADC values for control versus hyperoxia were calculated
using GraphPad 5.0 software.
To induce hyperoxia in the neonatal mouse, WT mice were ex-
fold increase (n ? 5; p ? 0.001) in average oxygen tension (oxy-
gen partial pressure) compared to control animals kept in room
4330 • J.Neurosci.,March16,2011 • 31(11):4327–4344 Schmitzetal.•HyperoxiaandWhiteMatterDevelopment
air. No change in any metabolic panel value was observed in the
hyperoxia group when compared to control (Table 1).
MBP is part of the major family of myelin proteins (Bau-
mann and Pham-Dinh, 2001; Back, 2006), and MBP expres-
sion marks the onset of developmental myelination. We
therefore sought to investigate the effect of hyperoxia on MBP
expression through immunofluorescence and Western blot
analyses in the developing WM. When compared to controls,
animals exposed to hyperoxia showed a marked decrease in
MBP immunofluorescence at P8, P10, and P12 (Fig. 1A). A
reduction in MBP protein expression immediately after expo-
sure at P8 (56.4% decrease; n ? 4; p ? 0.025), after 2 d of
recovery at P10 (95.3% decrease; n ? 3; p ? 0.005), and after
4 d of recovery at P12 (66.0% decrease; n ? 4; p ? 0.025)
confirmed these observations (Fig. 1B,C). To determine
whether hyperoxia caused a chronic disturbance in MBP ex-
pression, levels were measured after more prolonged periods
of recovery. Both immunostaining and Western blots revealed
P30 (Fig. 1A–C). These results indicate two distinct phases of
myelin regulation after hyperoxia exposure: an acute phase
from P8 to P12, where MBP is reduced, followed by a second-
ary phase during continued recovery that involves compensa-
tory MBP formation and the return of MBP to control levels at
P15 and P30.
to become myelinating oligodendrocytes:
(1) OPCs, (2) pre-OLs, (3) immature/pr-
emyelinating oligodendrocytes, and (4)
mature oligodendrocytes (Baumann and
Pham-Dinh, 2001; Back, 2006).
We characterized the oligodendroglia
population at P4, P6, and P8 in neonatal
mice kept in room air (control) to deter-
stages of oligodendroglial development at
these ages. We found that the number of
total oligodendroglia (Olig2?cells) and
creased throughout early postnatal devel-
opment in the WM (Table 2). The OPC
(NG2?O4?cells) population appeared
OL lineage in the first postnatal week of
WM development. However, as the WM
matures, the relative contribution of
NG2?O4?cells (percentage of Olig2?
cells) to the overall oligodendroglia pop-
ulation decreases over time. From P4 to
ing a pre-OL phenotype (NG2?O4?and
O4?NG2?cells) remained consistent
throughout this time period, even though
the actual number of pre-OLs increased
throughout WM development (Table 2).
Last, in agreement with our observation
that the developing WM contains a low
percentage of CC1?cells between P4 and
able in the WM at P4 and P6 (data not
findings of Gerstner et al. (2008), we observed changes in WM
MBP expression after hyperoxia exposure during the first post-
natal week of development. Therefore, during this critical devel-
opmental time window, the WM of the rodent is of an immature
state and an insult during this period is more likely to result in
Our next goal was to determine the cellular changes underly-
ing the observed hyperoxia-induced delay in MBP expression in
the developing WM. Consistent with the changes in MBP, new-
born mice exposed to 48 h of hyperoxia displayed a reduction in
the number of Olig2?cells (total oligodendroglia) and CC1?
the WM at P8 (Fig. 2A–E). At P10, the hyperoxia group only
showed a reduction in the number of CC1?cells, and no differ-
groups in either WM region of interest (Fig. 2F). Interestingly,
to hyperoxia, the number of CC1?oligodendroglia returned to
control levels in both WM regions at P12 (Fig. 2I–K). This anal-
glial population within the developing WM, followed by
expression after hyperoxia at P8, P10 and P12. MBP expression returned to control levels at P15 and P30. Values represent
standardized mean ratios of MBP and ?-actin. For each group and time point, n ? 3–6 brains. An unpaired t test comparing
Schmitzetal.•HyperoxiaandWhiteMatterDevelopment J.Neurosci.,March16,2011 • 31(11):4327–4344 • 4331
oligodendrocytes after recovery in room air. Changes in the
CC1?cell population are likely responsible for the observed bi-
phasic response in MBP expression after hyperoxia.
It is well established that OPCs and pre-OLs differentiate into
Numbers of cells at various stages of oligodendroglia development in the postnatal WM. The Olig2? (total oligodendroglia) population expands throughout early postnatal development. NG2?O4?cells (OPCs) appear to be the
CC1+ OLIG2+ CC1+
4332 • J.Neurosci.,March16,2011 • 31(11):4327–4344Schmitzetal.•HyperoxiaandWhiteMatterDevelopment
and Pham-Dinh, 2001; Nishiyama et al., 2009), and OPCs play a
al., 2005; Franklin and Ffrench-Constant, 2008). We analyzed
changes in WM NG2?cells, which may underlie the hyperoxia-
induced decrease in MBP expression and the subsequent recov-
ery of MBP in the WM at P15. We also analyzed total NG2?cell
proliferation in newborn WT mice using BrdU immunofluores-
cence. Immediately after 48 h of hyperoxia at P8, there was a
significant decrease in the number of NG2?cells found within
the EC and CG when compared to control values (Fig. 3A,B,I).
In addition, hyperoxia diminished the number of proliferating
NG2?cells (NG2?BrdU?) in both WM regions of interest (Fig.
3C,D,I). After 2 d of recovery in room air (P10), we found no
After 4 d of recovery at P12, we observed a 43% increase in the
number of NG2?cells and a 62% increase in NG2?BrdU?cells
within the CG of the hyperoxia-exposed animals compared with
control animals (Fig. 3E,H–K). No difference in the number of
NG2?or NG2?BrdU?cells was detected in the EC at this time
point (Fig. 3K).
to oligodendroglia (Nishiyama et al., 2009). Therefore, to verify
that changes in the NG2?cell population observed in WT mice
were cells of the oligodendroglial lineage, the number of NG2?
cells in CNP-EGFP transgenic mice exposed to hyperoxia and
room air was evaluated. In the CNP-EGFP mouse, cells of the
oligodendroglial lineage are labeled with green fluorescent pro-
tein (GFP), which is under the control of the CNP promoter
(Yuan et al., 2002). Notably, virtually all NG2?cells in the WM
and temporal cellular response similar to that observed in WT
mice was found in NG2?GFP?progenitors of the CNP-EGFP
transgenic mice. At P8, the number of NG2?GFP?cells was
mental Fig. 2A,B,E, available at www.jneurosci.org as supple-
mental material). After 4 d of recovery at P12, NG2?GFP?cells
not in the EC (supplemental Fig. 2C,E, available at www.jneurosci.
These results indicate that hyperoxia alters the growth of the
NG2?oligodendroglia population. NG2?cells are also likely
Time-dependent changes in white matter NG2?oligodendrocyte progenitor cells after hyperoxia. A, B, Confocal images of NG2?progenitors under control conditions or after
Schmitzetal.•HyperoxiaandWhiteMatterDevelopmentJ.Neurosci.,March16,2011 • 31(11):4327–4344 • 4333
involved in the repair of the developing
WM after hyperoxia. This is supported by
our observation that hyperoxia causes an
acute reduction in the number of NG2?
we found a greater number of NG2?and
NG2?BrdU?cells when compared to
restoration of the CC1?cell population af-
The immature rat brain, especially the
WM, is susceptible to hyperoxia-induced
with cells of the oligodendroglial lineage
exhibiting several maturation-dependent
factors that enhance susceptibility to oxi-
Volpe, 2008; Bradl and Lassmann, 2009).
Recent in vitro studies revealed immature
oligodendroglia to be most vulnerable to
caspase-dependent apoptosis, whereas
(Gerstner et al., 2008). To assess overall
oping WM in vivo, we first analyzed the
ECandCGregions together for the pres-
ence of cleaved caspase-3? immunopo-
(TUNEL?) cells. In wild-type mice, hy-
peroxia caused a significant increase in
the number of Casp3?cells at P6 (after
6 h of hyperoxia), P8, and P12, with the
peak of cell death occurring at P8 (Fig.
4A). Hyperoxia-induced apoptosis was
virtually undetectable at P15 and P30
To verify our results obtained in WT
mice, we also analyzed the CNP-EGFP
transgenic mouse and found that these
mice when exposed to hyperoxia exhib-
ited a similar pattern of apoptotic cell
death in the WM. We observed a signif-
ter 4 d recovery at P12, when compared
to controls at these time points (Fig.
To identify the developmental stage of oligodendroglia sus-
genic mice were first immunostained for Casp3 and CC1. No
oxia exposure, indicating oligodendroglia undergoing cell death to
munohistochemical analysis was performed using TUNEL and an-
number of TUNEL?and NG2?O4?TUNEL?cells in the hyper-
or NG2?O4?TUNEL?was found between the two experimental
groups (Fig. 4F). At P12, we also observed a significant increase in
TUNEL?and NG2?O4?TUNEL?cells in the hyperoxia group
with very little cell death in control animals (Fig. 4I). Again, no
difference was found when comparing the numbers of O4?
NG2?TUNEL?and NG2?O4?TUNEL?cells in each group at
In summary, hyperoxia causes apoptosis in the OPC popula-
tion (NG2?cells) of the developing WM. This increased OPC
cell death, both during and immediately after hyperoxia expo-
sure, may in part explain the changes in (1) Olig2?/CC1?cell
numbers and (2) MBP expression.
no difference observed at P15 and P30. B, C, Images of activated Casp3?cells in P8 CNP-EGFP mice under control
conditions or after hyperoxia show more apoptosis in pups exposed to hyperoxia. Arrowheads indicate Casp3?GFP?
positive cells. D, Numbers of Casp3?GFP?cells in CNP-EGFP mice within both regions combined at P8 and P12 showed
cells to be negative for CC1, indicating the cells undergoing apoptosis were immature, not mature oligodendroglia. F,
G, H, Confocal images of TUNEL?and NG2?O4?TUNEL?cells in control and hyperoxia exposed animals. Arrowheads
indicate double immunopositive cells (NG2?TUNEL?). I, Quantification of TUNEL?, NG2?O4?TUNEL?,
O4?NG2?TUNEL?, and NG2?O4?TUNEL?cells at P12. J, Representative image from a P8 hyperoxia-exposed animal
showing several NG2?O4?TUNEL?cells. Scale bars, 50 ?m. Data are shown as mean ? SD (n ? 3–6 brains for each
Hyperoxia causes cell death of NG2?oligodendroglia in the developing white matter. A, Quantification of
4334 • J.Neurosci.,March16,2011 • 31(11):4327–4344Schmitzetal.•HyperoxiaandWhiteMatterDevelopment
The generation of mature, myelinating oligodendrocytes occurs
throughout development and is also crucial in WM repair after
oligodendrogenesis during the recovery period from P8 to P12, the
generation of newly formed oligodendrocytes was determined
through analysis of BrdU and CC1 double
immunopositive cells after cumulative
erated BrdU?CC1?oligodendrocytes was
increased in the hyperoxia-exposed group
cell populations between P8 and P12 re-
vealed that hyperoxia caused an 8.41-fold
average increase in the total number of
CC1?cells in the two WM regions of in-
terest compared to a 4.01-fold increase in
controls ( p ? 0.05; one-way ANOVA;
Holm-Sidak). This indicates that more
CC1?oligodendroglia were generated in
oxia over this time period.
Together with our OPC observa-
tions described previously, these findings
indicate that during the recovery phase,
restoration of the NG2?population con-
droglia in the WM at later time points.
The generation of new CC1?cells is then
pression observed at P15 and P30.
We hypothesized that the delay in oligo-
dendroglia development could produce
termine whether chronic disturbances
could be detected in the WM of mice ex-
posed to hyperoxia, FA was measured by
DTI. The measurements of diffusivity by
DTI using MRI represents a technique of
high sensitivity for changes of myelina-
tion and WM organization both during
development and after injury (Basser and
Pierpaoli, 1996; Inder et al., 1999). In ad-
dition, FA is more sensitive than conven-
injury (Counsell et al., 2003; Huppi and
Dubois, 2006). FA in the white matter in-
been associated with myelination in the
2006). In a hypoxia model of WM dam-
age, where neonatal mice were exposed
from P3 to P11, the CC was established as
a reliable region to determine FA de-
creases at age P45 to P51, and FA in the CC was markedly de-
low FA in the CC significantly correlated to poor neurological out-
come assessed by neurodevelopmental tests (Chahboune et al.,
DEC maps to represent anisotropy in the lateral–medial direc-
Increased oligodendrogenesis after hyperoxia exposure throughout the immature white matter. Six-day-old (P6)
o r t o
s i n
A l a
o i t c
region of CC that was selected for DTI measurements. C1, D1, FA obtained by DTI in control and hyperoxia-exposed animals,
Schmitzetal.•HyperoxiaandWhiteMatterDevelopmentJ.Neurosci.,March16,2011 • 31(11):4327–4344 • 4335
tional manners of the CC. In the anisotropy
in P30 animals that had been exposed to hy-
peroxia from P6 to P8 (Fig. 6B,D1,F1,F2,I)
(mean, 0.450; 25th–75th percentile, 0.433–
0.471) compared to litter-matched control
animals always kept in room air (Fig.
6A,C1,E1,E2,I) (mean, 0.580; 25th–75th
percentile, 0.524–0.601; n ? 6; Mann–
Whitney rank test; p ? 0.005). This signifi-
cant decrease of FA in the CC was
hyperoxia, 0.525; 25th–75th percentile,
0.495–545; mean control, 0.623; 25th–75th
percentile, 0.580–0.680; n ? 6; p ? 0.005).
To distinguish between diffusivity that
is parallel or perpendicular to the direc-
tory of CC fibers, we determined radial
(perpendicular) and axial (parallel)
ADCs. The radial ADC was increased af-
ter hyperoxia both at P30 (median,
0.615 ? 10?3mm2/s; range, 0.441–
0.809) and at P60 (median, 0.715 ?
10?3mm2/s; range, 0.528–0.734) com-
pared to controls (P30 median, 0.420 ?
10?3mm2/s; range, 0.420–0.612; P60
median, 0.465 ? 10?3mm2/s; range,
0.411–0.491; t test, p ? 0.01 and p ?
0.05, respectively; n ? 6–7). The axial
1.154 ? 10?3mm2/s after hyperoxia at
10?3mm2/s, respectively, at P60. Both
the FA and ADC data indicate that, de-
spite the restoration of the WM oligo-
dendroglia population and subsequent
recovery of MBP expression by P15, neonatal exposure to hy-
peroxia produces long-term damage to the WM.
In the course of characterizing hyperoxia-induced changes
within the oligodendroglial lineage, we considered that changes
in the functional properties of other WM cell types could inter-
fere with the survival or growth of oligodendroglia. Astrocytes
Kuroda and Shimamoto, 1991; Levine et al., 1999) and repair
(Davies et al., 1997; Li et al., 2008). We hypothesized that astro-
cytes may contribute to hyperoxia-induced WM damage, and
therefore analyzed the properties of WM astrocytes to determine
their reactivity in response to hyperoxia.
Hyperoxia exposure caused a biphasic astrocytic response in
the developing WM. Immunofluorescence and Western blots
were first used to assess GFAP expression. At P8, immediately
after 48 h of hyperoxia, GFAP immunoreactivity was reduced in
in GFAP protein expression after hyperoxia (Fig. 7J,K). In con-
trast, at P12, the intensity of GFAP immunofluorescence was
elevated in the hyperoxia treated group compared to controls
(Fig. 7E–I). The increase of GFAP expression was confirmed by
Western blot analysis (Fig. 7J,K).
This initial decrease followed by an increase in GFAP expres-
model of transient global ischemia (Ouyang et al., 2007). The
similarity between the pattern of GFAP expression and that of
changes in the oligodendroglia lineage after hyperoxia suggests a
role for astrocyte dysfunction in WM damage.
To quantify the number of astrocytes in the WM under both
experimental conditions we performed coimmunolabeling for
GFAP and GS.
GFAP?processes of fibrous astrocytes are often at a distance
from the cell soma or nucleus, making identification and quan-
sion of GS, the enzyme responsible for the conversion of
glutamate and ammonia to glutamine, is highly confined to the
soma of all astroglia in the WM (Miyake and Kitamura, 1992).
at P8 or P12 (Fig. 8D,H). We also performed GS immunostain-
labeled with GFP expressed under the control of the human
GFAP promoter (Nolte et al., 2001) to asses astrocyte prolifera-
tion (Ki67) and apoptosis (Casp3) after hyperoxia. At all time
points evaluated, no Ki67?GFP?GS?astrocytes were observed
in either the hyperoxia or control group (supplemental Fig. 3,
immediately after 48 h of hyperoxia exposure at P8 and after 4 d of room air recovery at P12. A–H, Confocal images of GFAP
4336 • J.Neurosci.,March16,2011 • 31(11):4327–4344Schmitzetal.•HyperoxiaandWhiteMatterDevelopment
cell death was observed in GFP?GS?WM astrocytes (supple-
mental Fig. 4, available at www.jneurosci.org as supplemental
material), even at time points at which immature oligoden-
droglia displayed susceptibility to hyperoxia-induced apopto-
sis (Fig. 3).
GFAP?GS?astrocytes express high-affinity glutamate up-
take transporters; GLAST (EAAT-1), GLT-1 (EAAT-2), and
EAAC1 (EAAT-3) (Danbolt, 2001; Fukamachi et al., 2001; Mi-
ralles et al., 2001; Liu et al., 2006; Arranz et al., 2010) and the
is regulated during postnatal development throughout the ro-
dent brain (Furuta et al., 1997; Regan et al., 2007). GLAST and
GLT-1 are expressed on the astrocytic cell membrane, function
in vivo, and are responsible for the majority of astrocyte-
mediated glutamate uptake (Anderson and Swanson, 2000;
Danbolt, 2001). EAAC1 is thought to be primarily neuronal
(Danbolt, 2001), and the functional significance of EAAC1 in
glutamate uptake performed by astrocytes has yet to be fully
established. Even though cultured astrocytes do express
EAAC1, it was not found to be expressed at the cell surface
(Dallas et al., 2007; Arranz et al., 2010).
Intriguingly, recent evidence points to a functional interac-
tion between GFAP and GLAST (Sullivan et al., 2007), and both
2007) as well as ablation of GFAP have been associated with re-
duced glutamate clearance (Hughes et al., 2004). Modifications
in glutamate uptake have been shown to cause glutamate-
mediated excitotoxic cell death of immature oligodendroglia
(Matute et al., 2002, 2007; Deng et al., 2006), and activation of
oligodendroglia non-NMDA GluRs can lead to decreased OPC
proliferation and the inhibition of lineage progression (Gallo et
al., 1996; Yuan et al., 1998). Therefore, we reasoned that the
sociated with altered astrocyte-mediated glutamate uptake,
which may ultimately have detrimental effects on the oligoden-
droglia population of the immature WM.
To investigate a possible link between hyperoxia and altered
glutamate homeostasis in the developing WM, we analyzed the
HYPEROXIA (3)CONTROL (3)
HYPEROXIA (3) CONTROL (3)
expressing GLT-1 at P8. F, G, Immunolabeling with antibodies for GFAP, GS, GLT-1, and DAPI to analyze astrocyte numbers and the amount of astrocytes expressing GLT-1 in WT mice at P12.
Schmitzetal.•HyperoxiaandWhiteMatterDevelopment J.Neurosci.,March16,2011 • 31(11):4327–4344 • 4337
number of astrocytes expressing GLAST
number of GFAP?GS?GLAST?cells
and a 14.08% reduction in the astrocyte
population expressing GLAST was found
at P8 in the hyperoxia group, with no dif-
ference in either parameter detected at
P12 (Fig. 8A1–E). When quantifying the
number of astrocytes expressing GLT-1,
there was no statistically significant differ-
ence in total GFAP?GS?GLT-1?cells or
trol groups at either P8 or P12 (Fig. 8D–
I). In addition, the majority (?95% at P8
and 85% at P12) of cells expressing either
GLAST or GLT-1 were GFAP?GS?as-
both experimental groups, also displayed
a differential expression pattern, where
GLAST immunoreactivity could be ob-
served in the cell soma or astrocytic pro-
cesses or in both locations (Fig. 8C). In
contrast, GFAP?GS?GLT-1?cells ex-
hibited a more uniform distribution of
Protein expression in microdissected
WM tissue was also measured at both P8
WM of mice exposed previously to hyper-
oxia when compared to age-matched con-
trols (n ? 8; p ? 0.005), and GLAST
expression returned to control levels at P12
glutamate uptake by altering GLAST expression on GFAP?GS?
To determine whether hyperoxia directly affects astrocyte-
mediated sodium-dependent glutamate uptake during early
performed in tissue extracts of the WM and in culture. The
amount of total and non-GLT-1 mediated (100 ?M DHK was
used to selectively block GLT-1) sodium-dependent [3H]-D-
aspartic acid uptake was determined by analyzing WM taken at
P8, P12, P15, and P30. TBOA, a nontransportable agent that
selectively blocks all EAATs was also administered to a subset of
samples, at a concentration of 75 ?M. At P8, the rate of total
[3H]-D-aspartic acid uptake was 1.11 ? 0.13 pmol/mg protein ?
min?1in the control group decreased to 0.75 ? 0.24 pmol/mg
protein ? min?1in the hyperoxia group (Fig. 9A). Non-GLT-1-
mediated [3H]-D-aspartic acid uptake was also decreased from
0.83 ? 0.07 pmol/mg protein ? min?1in controls to 0.47 ? 0.16
pmol/mg protein ? min?1in the hyperoxia group (Fig. 9A) (n ?
7; p ? 0.001). A similar pattern was observed at P12, where total
[3H]-D-aspartic acid uptake and non-GLT-1 uptake were re-
control, total, 1.28 ? 0.11 pmol/mg protein ? min?1; non-GLT,
0.73 ? 0.04 pmol/mg protein ? min?1; hyperoxia, total, 0.68 ?
0.09 pmol/mg protein ? min?1; non-GLT, 0.23 ? 0.04 pmol/mg
protein ? min?1) (Fig. 9B). No change in total or non-GLT-1
[3H]-D-aspartic acid uptake was found between control and hy-
peroxia groups at either P15 (Fig. 9C) or P30 (data not shown).
There was a significant difference between total and non-GLT-
mediated (?DHK) [3H]-D-aspartic acid uptake in both control
and hyperoxia groups at P8 and P12 (Fig. 9A,B). Last, no differ-
when comparing hyperoxia and control groups (Fig. 9A–C).
Confluent cultures (treated with db-cAMP) exposed to 80%
oxygen for 48 h demonstrated a significantly reduced rate of
sodium-dependent [3H]-D-aspartic acid uptake, with a Vmax
ues at 100 ?M [3H]-D-aspartic acid (95% confidence interval; p ?
compared with controls, but this trend did not reach statistical sig-
nificance (hyperoxia, 6.5 ? 1.6 fmol/mg protein ? min?1; controls,
10.6 ? 2.8 fmol/mg protein ? min?1). Levels of GLAST protein ex-
hyperoxia. Western blot analysis revealed a 40% decrease in total
GLAST (Fig. 9E) (summation of monomeric 65 kDa, and multi-
meric 150 and 180 kDa bands) (Haugeto et al., 1996) expression in
cultured astrocytes kept at 21% oxygen [control, 2.26 ? 0.23
tio); data are mean ? SD; n ? 4; p ? 0.05, using an unpaired t test
Together, these data suggest that hyperoxia exposure de-
creases sodium-dependent glutamate uptake transporter activity
in astrocytes through a non-GLT-mediated mechanism. This is
in total and non-GLT-1-mediated (?DHK) [3H]-D-aspartic acid uptake when compared to control (n ? 7). B, Lower total and
non-GLT-1-mediated [3H]-D-aspartic acid uptake was also observed at P12 (n ? 4). C, No difference in total or non-GLT-1-
*p ? 0.001 was considered to be significant. D, Representative measurement of the maximum velocity of sodium-dependent
Astrocyte-mediated sodium-dependent [3H]-D-aspartic acid uptake is decreased after hyperoxia in tissue and in
4338 • J.Neurosci.,March16,2011 • 31(11):4327–4344 Schmitzetal.•HyperoxiaandWhiteMatterDevelopment
supported by our previous data showing GLAST to be the pri-
mary glutamate transporter present on astrocytes in the WM of
in both experimental groups at P8 and P12 (Fig. 9A,B) indicates
that the GLT-1-mediated component of sodium-dependent glu-
the observed decrease in [3H]-D-aspartic acid uptake may be
contributed largely by GLAST. This is in agreement with our
observation that there was no change in the number or percent-
age of astrocytes expressing GLT-1 (Fig. 8D–I) and no change in
astrocyte-mediated glutamate uptake/clearance has been impli-
cated in animal models in which oxidative stress is prominent
(Chen et al., 2005; Yeh et al., 2005). However, unlike our study,
in this model of brain injury in the adult rodent.
To analyze the role of astrocyte dysfunction in hyperoxia-
induced WM damage, we studied the effects of 80% oxygen on
purified rat astrocytes under various
culture conditions. In subconfluent as-
trocyte cultures, the numbers of GFAP?
peroxia (Fig. 10A). Immature vimentin?
astrocytes were also reduced in the
hyperoxia-exposed cultured astrocytes
(Fig. 10A). In addition, the number of
proliferating GFAP?and vimentin?as-
labeling, were decreased in hyperoxic
cultures (Fig. 10A–E).
It has been well described that cAMP
stimulation of cultured astrocytes can con-
vert flat polygonal shaped astroglia into
process-bearing stellate astrocytes (Favero
semble astrocytes in vivo with regard to
morphology and differentiation (Shapiro,
cultures not treated with db-cAMP, 48 h of
pared to cultures kept at 21% oxygen (Fig.
10F–H). Furthermore, in agreement with
the expectation that astrocyte proliferation
age of Ki67?GFAP?cells was reduced by
db-cAMP in both experimental groups.
However, hyperoxia and db-cAMP stimula-
tion in combination produced a further de-
in culture (Fig. 10F,I,J). No cell death, mea-
sured by TUNEL immunofluorescence, was
observed in cultured astrocytes (confluent or
or 21% oxygen (supplemental Fig. 5A,
available at www.jneurosci.org as supple-
mental material). In confluent cultures,
hyperoxia did not alter the numbers of
GFAP?cells despite a modest, but signif-
icant, reduction in Ki67 expression. The reason for this is un-
caused an upregulation of GFAP levels (supplemental Fig. 5B,C,
available at www.jneurosci.org as supplemental material). It
erties of astrocytes in vivo and in culture are observed. Both hy-
peroxia paradigms were designed to result in oxygen levels
twofold to fourfold above normal conditions. The oxygen levels
in vivo, and these extreme conditions are likely to underlie some
to hyperoxia. Nevertheless, the changes in glutamate transport
after hyperoxia are in agreement in both systems; therefore, this
mechanism of damage is being further investigated in culture.
ological glutamate challenge can inflict excitotoxic damage to
Cells / mm2
Cells / mm2
Hyperoxia decreases cell proliferation in primary cultured astrocytes. A–C, In subconfluent cultures of GFAP?
Schmitzetal.•HyperoxiaandWhiteMatterDevelopmentJ.Neurosci.,March16,2011 • 31(11):4327–4344 • 4339
surrounding cells (Matute et al., 2007).
The overactivation of non-NMDA GluRs
can cause decreased OPC proliferation
and inhibit oligodendroglia lineage pro-
gression (Gallo et al., 1996; Yuan et al.,
1998). We wanted to determine whether
the reduction in astrocyte-mediated glu-
tamate uptake induced by hyperoxia al-
tered the OPC (A2B5?cells) and/or O4?
oligodendroglia populations in the pres-
To test this, we established an astrocyte–
OPC coculture system in which dye-
labeled OPCs were added to primary
astrocytes exposed previously to hyper-
oxia or normoxia. After the presentation
of an exogenous glutamate challenge (1
mM), OPCs were immunostained and an-
cells were analyzed after 24 h, and O4?
cells after 48 h, in culture. Glutamate did
not significantly affect the number of
OPCs in cocultures prepared with nor-
moxic (control) astrocytes. (Fig. 11A–D,I).
However, glutamate challenge significantly
peroxia (Fig. 11A,B,E,F,I). Among the
experimental groups that contained as-
but did not receive a glutamate challenge,
or A2B5?Dye?cells (Fig. 11I). The toxic
effect of glutamate was prevented by ex-
posure to the non-NMDA-receptor an-
tagonist NBQX (100 ?M) 1 h before
glutamate challenge (Fig. 11, compare E,
G for A2B5; compare F, H for O4).
Our results obtained in astroglia–OPC
coculture support the notion that hyper-
oxia diminishes extracellular glutamate
uptake by a mechanism likely to involve
reduced expression of GLAST, which in
turn could inhibit the growth or survival
perform in vivo time course analysis of the oligodendroglial lin-
eage after hyperoxia, (2) demonstrate recovery of WM oligoden-
effect on WM astrocyte phenotype and function, and (4) analyze
the long-term consequences of neonatal hyperoxia. We demon-
strate that hyperoxia exposure in newborn mice perturbs WM
development by decreasing proliferation and increasing apopto-
sis in NG2?oligodendroglia. Hyperoxia also alters astroglial
GFAP and GLAST expression in vivo and decreases [3H]-D-
aspartic acid uptake. Moreover, during recovery, animals ex-
posed previously to hyperoxia exhibited compensatory (1)
drocytes, and (3) MBP protein expression in the WM. However,
despite the number of oligodendroglia, MBP levels, and [3H]-D-
impairment of WM integrity detected by DTI persisted into
cause WM damage, hypomyelination, and altered WM integrity
(Scafidi et al., 2009; Li et al., 2010). Several studies have also
myelinating oligodendrocytes is thought to be a major contrib-
Control - GluControl + Glu
Hyperoxia + Glu
Hyperoxia + Glu + NBQX
Cell Number (% Control - Glu)
Control + Glu
Hyperoxia - Glu
Hyperoxia + Glu
Hyperoxia + Glu + NBQX
Control + Glu
Hyperoxia - Glu
Hyperoxia + Glu
Hyperoxia + Glu + NBQX
Control + Glu
Hyperoxia - Glu
Hyperoxia + Glu
Hyperoxia + Glu + NBQX
cocultures without glutamate challenge. C, D, Glutamate challenge does not significantly affect A2B5?cells and O4?cells in
non-NMDA-receptor antagonist NBQX (100 ?M) 1 h before glutamate challenge significantly attenuated the toxic effects on
A2B5?/Dye?(G) and (H) O4?/Dye?cells. I, Values were normalized with DAPI and expressed as a percentage of control
O4?Dye?). *p ? 0.01; **p ? 0.001 (unpaired student’s t test vs Control ? Glu);#p ? 0.01, unpaired student’s t test vs
Reduced astrocyte-mediated protection of OPCs during glutamate challenge is attenuated by NBQX. Astrocytes
4340 • J.Neurosci.,March16,2011 • 31(11):4327–4344 Schmitzetal.•HyperoxiaandWhiteMatterDevelopment
uting factor in chronic PWMI (Back et al., 2007; Khwaja and
Volpe, 2008), and oligodendroglia exhibit a maturation-
dependent vulnerability in vitro (Gerstner et al., 2006, 2008). We
found significantly more NG2?O4?TUNEL?cells after hyper-
oxia at P8, P10, and P12, without changes in the number of
O4?NG2?TUNEL?or NG2?O4?TUNEL?cells. These data
indicate that, in vivo, NG2?progenitor cells, and not O4?preo-
ligodendrocytes, are highly susceptible to hyperoxia-induced
OPCs are the primary source of oligodendrocytes in the de-
veloping brain (Baumann and Pham-Dinh, 2001; Nishiyama et
al., 2009) and in remyelinating WM lesions (Zhao et al., 2005;
Franklin and Ffrench-Constant, 2008). The preterm infant is
most vulnerable to PWMI when immature, premyelinating oli-
godendroglia, including OPCs, predominate in the developing
WM (Back et al., 2001). Our observations in the proliferating
NG2?cell population are consistent with other studies that in-
dicate that changes in oligodendroglia proliferation constitute
mechanisms of injury (Robinson et al., 2005; Yan and Rivkees,
injury (Amat et al., 1998; Segovia et al., 2008) in the developing
brain. Our finding that during recovery more NG2?and
local NG2?cell populations in the various WM regions respond
differentially to injury, or that targeted recruitment of migratory
OPCs into the cingulum occurs from germinal regions in close
proximity, such as the subventricular zone.
the NG2?cell population after hyperoxia is likely to be respon-
sible for the decrease in WM CC1?oligodendroglia observed at
P8 and P10. This is because of fewer immature oligodendroglia
being available to undergo maturation. The reduction in mature
oligodendrocytes is, at least in part, responsible for the decrease
To evaluate the long-term impact of neonatal hyperoxia on
adult mice (P60). Changes in FA have been associated with de-
myelination, axonal damage and behavioral abnormalities
(Chahboune et al., 2009; Xie et al., 2010), and preterm infants
exhibit a loss in WM volume and FA when compared to term
infants (Dudin et al., 2007). Our DTI measurements in the CC
demonstrate that neonatal hyperoxia during a critical develop-
diffusivity both at P30 and P60. These findings indicate that ab-
normalities in WM organization, including packing and align-
ment of WM fibers, can occur despite apparently normal
restored by P15. Further studies will determine the impact of
these long-term WM deficiencies on functional properties of the
Astroglial reactivity plays a role in injury of the developing
WM and was identified in ischemia (Biran et al., 2006), trauma
(Rostworowski et al., 1997), and infection (Rousset et al., 2006).
Although we observed no hyperoxia-induced changes in astro-
cyte number or cell death or proliferation in vivo, the astroglial
porter GLAST overlapped temporally with oligodendroglial
changes and occurred in two phases: (1) an initial decrease in
hyperoxia at P8 and (2) increased levels of GFAP and restored
GLAST expression after 4 d of recovery in room air at P12. The
transporters GLAST and GLT-1 are coexpressed by both astro-
cytes and oligodendroglia in vivo and in vitro (Domercq et al.,
2005; Regan et al., 2007), and GLT-1 has been shown recently to
be important for glutamate homeostasis in cultured oligoden-
droglia (DeSilva et al., 2009). However, evidence suggests that
only mature oligodendrocytes express functional glutamate up-
take transporters in vivo (De Biase et al., 2010; Kukley et al.,
2010), and as indicated by Table 2, the majority of the cells in the
WM at P6 display an immature phenotype. Although we cannot
rule out the contribution of oligodendroglial GLAST and GLT-1
in WM glutamate homeostasis, our data indicate that at P8 and
decrease in the number GFAP?GS?GLAST?cells and GLAST
protein levels in the WM after hyperoxia. No change in total
GLT-1 expression or the number of GFAP?GS?GLT-1?cells
was found after hyperoxia.
of total and non-GLT-1-mediated sodium-dependent [3H]-D-
aspartic acid uptake when compared with controls. Similar re-
ductions in uptake were observed in control and hyperoxia
groups with the application of the GLT-1 inhibitor DHK at P8
and P12 (Fig. 9), suggesting that hyperoxia did not alter levels of
GLT-1 activity in the WM. Together, these observations support
a non-GLT-1-mediated mechanism of injury.
Intriguingly, total and non-GLT-1-mediated [3H]-D-aspartic
P12 and did not return to control levels until P15. This was de-
spite a recovery in GLAST expression within the WM. Possible
explanations for the decrease at P12 include posttranslational
regulatory mechanisms such as (1) decreased cell surface expres-
sion of the transporter, (2) decreased Na,K-ATPase coupling
(Rose et al., 2009), (3) altered interactions with the extracellular
matrix (Ye and Sontheimer, 2002), and (4) oxidative stress-
status can regulate glutamate uptake kinetics directly through
changes in reactive cysteine residues in transporter structure,
which allows maximal uptake activity only in the reduced state
(Trotti et al., 1997).
Altered glutamate homeostasis has been implicated in many
neurological disorders including epilepsy, Alzheimer’s disease,
and Parkinson’s disease (Doble, 1999; Tilluex and Hermans,
2007; David et al., 2009). Glutamate has been shown to cause
liferation and apoptosis of NG2?OPCs (Gallo et al., 1996; Yuan
of axoglial junctions (Fu et al., 2009). In our glial coculture sys-
tem, astrocytes exposed to hyperoxia were unable to protect
A2B5?and O4?oligodendroglial cells against the toxicity of
glutamate challenge. This vulnerability was eliminated in the
presence of the non-NMDA antagonist NBQX. Therefore, it is
possible that reduced glutamate uptake/clearance leads to higher
amounts of glutamate in the extracellular space. This phenome-
non may occur in isolation or in combination with enhanced
cellular and/or axonal glutamate release throughout the WM.
The resulting environment likely contributes to changes in the
oligodendroglia population. Excitotoxicity has been associated
with AMPA/kainite glutamate receptor activation in oligoden-
droglia (Alberdi et al., 2002; Sanchez-Gomez et al., 2003), and
OPCs may exhibit enhanced vulnerability, because they express
or immature/mature oligodendrocytes (De Biase et al., 2010;
Kukley et al., 2010).
Schmitzetal.•HyperoxiaandWhiteMatterDevelopmentJ.Neurosci.,March16,2011 • 31(11):4327–4344 • 4341
In conclusion, our analysis shows that hyperoxia modulates
glial interactions through a novel mechanism involving altered
astrocyte GFAP and GLAST expression and decreased astrocyte-
mediated glutamate uptake. These changes in astrocyte proper-
delayed MBP expression. Despite apparent recovery in the glial
population and in MBP levels, the disruption in oligodendroglia
ability leads to long-term deficiencies in WM organization and in-
tegrity. A better understanding of the cellular and molecular events
following hyperoxia will help define the role of glutamate in im-
aimed at improving WM integrity and function after hyperoxia
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