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The Journal of Neuroscience, January 1, 2003, 23(1):213-222
SRC-1 Null Mice Exhibit Moderate Motor Dysfunction and Delayed
Development of Cerebellar Purkinje Cells
Eijun
Nishihara,
Hiromi
Yoshida-Komiya,
Chi-Shing
Chan,
Lan
Liao,
Ronald L.
Davis,
Bert W.
O'Malley, and
Jianming
Xu
Department of Molecular and Cellular Biology, Baylor College of
Medicine, Houston, Texas 77030
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ABSTRACT |
Hormones and nuclear receptors (NRs) play important roles in brain
development and function. The recently identified steroid receptor
coactivator (SRC) family contains three homologous members that can
enhance transcriptional activities of NRs and certain non-NR
transcription factors. To study the role of SRC-1 in brain development
and function, we examined the spatial and temporal expression patterns
of SRC-1 and characterized the phenotypes of brain development and
function in SRC-1 knock-out
(SRC-1 /)
mice. In the adult mouse brain, SRC-1 is highly expressed in the
olfactory bulb, hippocampus, piriform cortex, amygdala, hypothalamus, cerebellum, and brainstem. Multiple behavioral tests revealed that
SRC-1/
mice exhibit normal hippocampal function but moderate motor
dysfunction. The behavior phenotypes correlate with the spatial
distribution of the SRC family members. In most brain structures where
SRC-1 is expressed, SRC-2 is expressed at lower levels; however, SRC-3 mRNA is detectable only in the hippocampus. In the adult cerebellum, Purkinje cells (PCs) preferentially express SRC-1 over SRC-2, but SRC-2
mRNA is slightly elevated in the
SRC-1/
PCs. During embryonic development, SRC-1 is expressed in the cerebellar
primordium. SRC-2 is expressed in PCs after postnatal day (P) 10. Time
course analysis revealed that the precursors of
SRC-1/ PCs
were generated ~2 d later than wild-type precursor cells. A further
delay in
SRC-1/ PC
maturation was detected at the neonatal stage. The morphology and
number of
SRC-1/ PCs
were equivalent to wild type by P10; this timing correlated with the
early expression of SRC-2 in the
SRC-1/
PCs. These results demonstrate that the relative levels of SRC expression are region specific, and the degree of overlapping expression may influence their functional redundancy. Disruption of
SRC-1 specifically delays the PC development and maturation in early
stages and results in moderate motor dysfunction in adulthood.
Key words:
coactivator; nuclear receptor; gene expression; animal model; motor function; Purkinje cell
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Introduction |
Steroid, thyroid, and retinoid
hormones play important roles in brain development and function
(McEwen, 1994 ; Lopes da Silva and Burbach, 1995). The gonadal steroids
affect sexual differentiation of the brain and regulate reproductive
behaviors by influencing neuroendocrine functions (Tetel, 2000; McEwen,
2001). The adrenal steroids are required for survival of the pyramidal
neurons in the dentate gyrus, but the stress-induced increase of
glucocorticoids is believed to produce atrophy of dendrites of CA3
pyramidal neurons in the hippocampus (McEwen, 1994). In addition,
hypothyroidism is accompanied by a significant reduction in Purkinje
cell dendritic arborization and synaptogenesis (Koibuchi and Chin,
2000). In the brain, most hormonal actions are mediated by their
cognate nuclear receptors (NRs), which are ligand-regulated
transcription factors (Tsai and O'Malley, 1994). Therefore, disruption
of either estrogen or progesterone receptors in female mice completely
diminishes their sexual receptivity (Mani et al., 1997; Rissman et al.,
1997). Furthermore, the glucocorticoid receptor is essential for the regulation of hypothalamus-pituitary-adrenal axis and
stress response (Reichardt et al., 2000). In addition, severe defects
in brain development and function have been observed in mice lacking
individual orphan NRs, including retinoid-related orphan receptor  (ROR), chicken ovalbumin upstream promoter transcription factor 1, nuclear-related receptor 1, and rev-erbA (Zetterstrom et al., 1997;
Steinmayr et al., 1998; Zhou et al., 1999; Chomez et al., 2000).
The transcriptional activities of NRs are mediated by interaction with
several classes of coactivators in a ligand-dependent manner (Glass and
Rosenfeld, 2000; McKenna and O'Malley, 2002; Xu and O'Malley, 2002).
The first and best characterized steroid receptor coactivator (SRC)
family contains three homologous members including SRC-1 (Onate et al.,
1995), SRC-2 (also known as GRIP1 and TIF2) (Hong et al.,
1996; Voegel et al., 1996), and SRC-3 (also known as p/CIP, RAC3, AIB1,
ACTR, and TRAM-1) (Anzick et al., 1997; Chen et al., 1997; Li et
al., 1997; Takeshita et al., 1997; Torchia et al., 1997;
Suen et al., 1998). Biochemical experiments have revealed that SRC
coactivators function in several ways. These include recruitment of
histone acetyltransferases such as CREB-binding protein (CBP),
adenovirus E1A-binding protein p300, and p300/CBP-associated factor
(p/CAF) (Torchia et al., 1997; Li et al., 2000), histone
methyltransferases such as coactivator-associated arginine
methyltransferase-1 and protein arginine
N-methyltransferase-1 for chromatin remodeling (Koh et al.,
2001), interaction with other coactivators such as
activating signal cointegrator 2 (ASC-2), steroid receptor RNA
activator (SRA), and RNA-binding DEAD-box proteins (p72/p68) (Lanz et
al., 1999; Lee et al., 1999; Watanabe et al., 2001), and contact with
certain general transcription factors such as transcription factor IIB
and TATA-binding protein (Takeshita et al., 1996).
To study the in vivo function of SRC-1, we generated SRC-1
null
(SRC-1/)
mice by gene targeting. We found that although
SRC-1/
mice had normal body size, their hormone target tissues such as the
uterus, prostate, testis, and mammary gland exhibited decreased growth
and development in response to steroid hormones (Xu et al., 1998). A
partial resistance to thyroid hormone (TH) was also observed in
SRC-1/
mice (Weiss et al., 1999). These results validated SRC-1 as an in
vivo NR coactivator. However, the role of SRC-1 in brain
development and function remained to be characterized.
In this study, we compared the spatiotemporal expression patterns of
all three SRC coactivators in brains of wild-type (WT) and
SRC-1/
mice and characterized the role of SRC-1 in brain development and
function. We show that SRC coactivators are preferentially expressed in
particular brain regions and disruption of SRC-1 causes delayed
development of Purkinje cells at the neonatal stage and moderate motor
dysfunction in adulthood.
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Materials and Methods |
Animals.
SRC-1/
mice were initially generated from embryonic stem (ES) cells
with a targeted deletion of ~9 kb of SRC-1 genomic sequence encoding
446 amino acids from Met-381 to Thr-826. The targeting event in ES
cells eliminated all functional domains of SRC-1 for transcriptional
activation, histone acetyltransferase activity, and interactions with
NRs and other coactivators such as CBP/p300 and p/CAF (Xu et al.,
1998). To achieve a homogenous genetic background, the initial SRC-1
mouse line with hybrid genetic background of C57BL/6J and 129/SVJ
stains was backcrossed to C57BL/6J for six generations. Subsequently,
WT and age-matched
SRC-1/
mice for this study were produced from heterozygous breeding pairs.
Mice were weaned at 3 weeks after birth and allowed free access to
water and lab chow. They were housed in an environment with a
controlled temperature of 19°C and 12 hr alternating darkness and
artificial light cycles. All mice used for behavioral tests were 15-20
weeks old. All parts of apparatus for behavioral tests were cleaned
with deionized water and 70% ethanol after each trial. Behavioral
tests were performed between 1 and 6 P.M. in a soundproof room in which
external noise was greatly reduced (~45 dB at 500 Hz). The test room
was equipped with an air cleaning and air conditioning system. All
animal experiments were performed according to approved protocols at
Baylor College of Medicine.
Hanging wire test. Eight WT and 11 SRC-1/
mice were tested on their ability to hang from a screen with wire bars
(10 × 18 cm area, 1 mm in diameter spaced 1 cm apart). After mice
were placed on the bars, the screen was waved gently in the air three
times to force the mice to grip the wires. The screen was turned upside down and latency to fall (maximum 60 sec) was measured. Mice that fell
in <10 sec were provided a second trial.
Rotarod test. Mice were placed on a rod (3 cm in diameter),
and the rod was accelerated from 4 to 40 rpm in 5 min. The time periods
that individual mice spent on the rod without falling were recorded.
Three trials with a 30 min intertrial interval were performed every day
for 3 d. Twenty WT and 20 age-matched SRC-1/
mice were tested.
Morris water task and swim speed. A circular polypropylene
pool (120 cm in diameter) was filled with opaque water. In the hidden
platform version, a 10 cm-diameter platform was submerged 1 cm below
the water surface. Ten WT and 9 SRC-1/
mice were initially guided to the platform and allowed to remain for 1 min. Subsequently, they were placed in the pool and allowed to search
for the platform for 1 min using visible external cues. Each mouse was
allowed to stay on the platform for 20 sec before the mouse was carried
back to its home cage. Escape latency, the time to find the platform,
was recorded in each trial. Each trial was initiated from a different
quadrant of the pool, and four consecutive trials with 60 min
intervals were performed per day for 7 d. The latencies of the
four trials in the same day were averaged and used for statistical
analysis. Probe trial was performed after 7 d training with a
computerized video tracking device (HVS Image) to track mouse movement.
In the probe trial, the platform was removed, and the trained mice were
allowed to search for the platform for 1 min. The latencies to find the
platform position, the swim distances, and the swim speeds were
measured in four consecutive trials with 15 min intervals.
Real-time RT-PCR. Real-time RT-PCR was performed
with the One Step Master Mix reagent (Applied Biosystems, Foster City,
CA) and total RNA samples by using the ABI 7700 Sequence Detection System (Applied Biosystems). In the measurement of SRC-1 mRNA, the
sequences of the forward and reverse primers and the TaqMan probe were
tatctctccagcccatggtgt, caaagttcccttggttgttgc, and
tcatccacgttgccaccatcca. In the measurement of SRC-2 mRNA,
the forward and reverse primers and the TaqMan probe were
gcagcacaggaaatagccatagt, catggccctcgctgagg, and
accaacagttccctcaatgcactgcaa. In the measurement of SRC-3 mRNA, the
forward and reverse primers and the TaqMan probe were
agcaaaggccacaagaaactg, ggtcaaggaggaatggcctc, and
cagttactcacgtgctcctccgacgac. Parallel measurements of the 18S RNA were
performed for each sample to serve as endogenous controls. Relative
expression levels of SRC coactivators were determined by the ratios of
SRC mRNA levels to 18S RNA levels.
In situ hybridization. Mice were decapitated and their
brains were frozen immediately with O.C.T. compound in an
ethanol-dry-ice bath. Sections were cut at 12 µm on a cryostat and
thaw mounted on Superfrost Plus slides (VWR Scientific, West Chester,
PA). The sections were fixed in 4% paraformaldehyde as described
previously (Nishihara et al., 1998). To prepare DNA templates for
transcribing riboprobes, individual mouse cDNA fragments corresponding
to a 527 bp sequence of SRC-1 (nucleotides 3910-4436, GenBank U56920), a 446 bp sequence of SRC-2 (nucleotides 4017-4462, GenBank AF000582), and a 498 bp sequence of SRC-3 (nucleotides 3377-3874, GenBank AF000581) were subcloned into EcoRI/BamHI sites
of the Bluescript II SK plasmid, respectively. These plasmids were
linearized with either BamHI or EcoRI digestion.
The 35S-UTP-labeled antisense or sense
riboprobes were transcribed using T7 or T3 RNA polymerase,
respectively. Hybridization solution was made of 1 × 107 cpm/ml of labeled probe, 50%
formamide, 0.3 M NaCl, 10% dextran sulfate, 20 mM Tris-HCl, pH 8.0, 10 mM
sodium phosphate, pH 8.0, 5 mM EDTA, 1×
Denhardt's solution, and 500 µg/ml yeast tRNA. Each slide was
applied with 100 µl of hybridization solution. All slides were
coverslipped and incubated at 55°C for 16 hr in a humidified chamber. The slides with sections were washed twice at 65°C for 30 min in 2× SSC containing 50% formamide and 100 mM  -mercaptoethanol, and three times at 37°C
in 10 mM Tris-HCl, pH 7.5, containing 0.5 M NaCl and 5 mM EDTA.
Sections were digested with ribonuclease A (20 µg/ml) at 37°C for
30 min. Slides were dehydrated in an ethanol series and dried at room
temperature. Then, slides were dipped in the photographic emulsion
(Kodak NTB2), exposed at 4°C for 6 d, developed in Kodak D-19,
and fixed with Kodak Fix. These sections were counterstained with
hematoxylin, dehydrated through a graded series of ethanol, and coverslipped.
Paraffin sections were used when immunohistochemical staining was
required after in situ hybridization. Sections were dewaxed, rehydrated, and treated with proteinase K (20 µg/ml) in 50 mM Tris-HCl, pH 7.5, and 5 mM EDTA for 7.5 min before hybridization was
performed as described above. After hybridization, sections were
digested with ribonuclease A and kept in PBS until immunohistochemical staining. Autoradiography of the immunostained sections was performed as described above.
Immunohistochemistry. Mice were anesthetized with Avertin
and perfused through the heart with 4% paraformaldehyde in PBS. The
brain was removed and kept in the same fixative for 16 hr at 4°C.
Samples were dehydrated in ethanol series, cleared in xylene, and
embedded in paraffin. Sections were cut at 7 µm. For SRC-1
immunostaining, sections were dewaxed, rehydrated, and boiled for 10 min in 10 mM citrate buffer, pH 6.0 (Nishihara et
al., 2000). After washing in PBS, endogenous peroxidase was blocked using 0.3% H2O2 in
methanol for 15 min. To reduce nonspecific binding of antibodies,
sections were washed in PBS again and preincubated with 500 µg/ml
goat IgG and 5% bovine serum albumin (BSA) in PBS for 60 min at room
temperature. Sections were incubated in a 1:400 dilution of the
polyclonal SRC-1 antibodies (Santa Cruz Biotechnology, Santa Cruz CA)
overnight at 4°C. After being washed in PBS with 0.075% Brij 35 (Sigma, St. Louis, MO), sections were further incubated with the
horseradish peroxidase (HRP)-goat anti-mouse IgG (1:500 dilution) for
2 hr at room temperature. Sections were washed again in PBS with
0.075% Brij 35. The sites of HRP were visualized by 3,3'-diaminobenzidine (DAB, Sigma). Finally, sections were
counterstained with methyl green.
Immunostaining of the calbindin and synaptophysin was performed using
the Histomouse kit (Zymed Laboratories, South San Francisco, CA).
Monoclonal antibodies against calbindin-D28k (Sigma) and synaptophysin
(Sigma) were diluted at 1:500 and 1:50, respectively. Sections were
treated with DAB and counterstained with methyl green.
BrdU labeling of the Purkinje cell precursors during embryonic
development was performed as described previously with slight modifications (Hayashi et al., 1988; Vogel et al., 2000). Briefly, successful mating between SRC-1 heterozygous males and females was
identified by checking coitus plugs in the morning. The day when the
coitus plug was found was defined as 0.5 d postcoitum (d.p.c.). On
10.5, 12.5, and 13.5 d.p.c., a single dose of BrdU (50 mg/kg body
weight) was injected intraperitoneally into the pregnant females.
BrdU labeling of Purkinje cells was examined on postnatal day (P) 5. All pups were genotyped by PCR as described previously (Xu et al.,
1998). Pups were decapitated, and their brains were dissected and fixed
in 10% formalin overnight at room temperature. Samples were embedded
in paraffin and sectioned as described above. Deparaffinized and
rehydrated sections were immersed in methanol containing 3%
H2O2 to inactivate
endogenous peroxidase. The sections were treated with 0.02% pepsin and
0.01N HCl in PBS for 60 min at 37°C and denatured with 2N HCl for 45 min at room temperature. These sections were neutralized in 0.1 M sodium borate for 45 min at room temperature and washed
with PBS. BrdU incorporation was detected using a monoclonal antibody
specific to BrdU (Sigma) at 1:50 dilution.
Cerebellar neuronal culture and immunocytochemistry.
Dissociated cerebellar neuronal cultures were prepared from the
cerebellum of mouse embryo on 18 d.p.c. according to the protocol
for Purkinje cell culture (Furuya et al., 1998). In brief, 5.0 × 106 cells/ml were plated onto glass
coverslips (12 mm in diameter; Becton Dickinson Labware, Mountain
View, CA) coated with poly-L-lysine and
cultured in DMEM/Ham's F-12 medium containing bovine insulin (10 µg/ml), BSA (100 µg/ml), gentamicin (10 µg/ml), glutamycine (200 µg/ml), human apotransferrin (100 µg/ml),
putrescine (100 nM), sodium selenite (30 nM), progesterone (20 nM),
tri-iodothyronine (T3) (0.5 ng/ml), and fetal bovine serum (10%). One
day after plating, the medium was changed to a serum-free medium. The
cultures were maintained at 37°C in a humidified incubator supplied
with 5% CO2. Half of the culture medium was
changed weekly with the same fresh medium.
For immunostaining, cells were fixed with 4% paraformaldehyde in PBS
for 30 min at 4°C and treated with PBS containing 0.05% Triton
X-100, 500 µg/ml goat IgG, and 1% BSA for 30 min at room temperature. Cells were incubated with a monoclonal antibody against calbindin-D28k (Sigma) at 1:400 dilution for 60 min at room temperature and then with a biotinylated anti-mouse IgG. The biotinylated secondary
antibody was visualized by a streptavidin peroxidase conjugate
(Vectastain, Vector Laboratories, Burlingame, CA) and subsequent
addition of DAB.
Data analysis. Statistical comparisons between
two groups were made using unpaired Student's t tests.
ANOVA with repeated measure was used in the analysis of behavioral
experiments. Experimental observers were blinded to mouse genotypes in
all behavioral studies. All values are expressed as mean ± SEM.
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Results |
Expression of SRC-1 in the adult mouse brain
To determine potential regions of SRC-1 function in the brain, the
expression pattern of SRC-1 mRNA in the adult mouse brain was examined
by in situ hybridization. SRC-1 was expressed at high levels
in the olfactory bulb, piriform cortex, hippocampus (CA1, CA2, and CA3
regions and dentate gyrus), amygdala complex, hypothalamus (relatively
high in the paraventricular nucleus), cerebellum, and brainstem (Fig.
1A-H).
Areas with strong hybridization signals in the brainstem include the
facial nucleus (Fig.
1E,G,J), trigeminal nucleus (both motor and mesencephalic nuclei) (Fig. 1E,G), and hypogrossal nucleus
(Fig. 1F). In the cerebellum, SRC-1 was selectively
expressed in the Purkinje cell layer but was undetectable in the
granule and molecule layers (Fig.
1E,H,K). The
SRC-1 protein was specifically detected in nuclei of Purkinje cells by
immunohistochemical analysis (Fig. 1L). In addition,
moderate levels of SRC-1 mRNA were observed in the thalamus and
isocortex (Fig. 1B-D). These data were
substantiated by the absence of SRC-1 mRNA signal in SRC-1/
mice when the same antisense probe was used in
SRC-1/
mice (Fig. 1I) and when a sense control probe was
used in WT mice (data not shown). These results indicate that the
relative expression levels of SRC-1 are dependent on specific brain
regions, suggesting that SRC-1 may have a functional preference in
those brain regions with higher levels of SRC-1.
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Figure 1.
Expression of SRC-1 mRNA in the adult mouse brain.
In situ hybridization was performed with
35S-labeled SRC-1 antisense RNA probe. Coronal
(A-D, F,
L) and sagittal (E,
G-K) brain sections were prepared
from 10-week-old WT mice (A-H,
J, K). As a negative control,
in situ hybridization was performed under identical
conditions on the sagittal section (I)
prepared from the cerebellum region of
SRC-1/
mice of the same age. A-I and
J-L were taken under dark-field and
bright-field lighting conditions, respectively. J and
K were stained with hematoxylin after in
situ hybridization, and their images were taken at higher
magnifications from the corresponding areas indicated in
G and H. Arrows indicate
the facial nucleus in J and Purkinje cells in
K and L. In L, SRC-1
protein (brown color) was specifically detected in the
nuclei of Purkinje cells by immunostaining. Scale bars:
A-I, 600 µm;
J-L, 25 µm. OB,
Olfactory bulb; Amy, amygdala complex;
Pir, piriform cortex; PVH,
paraventricular nucleus of the hypothalamus; Arc,
arcuate nucleus; HC, hippocampus; P,
Purkinje cell layer; V, trigeminal nucleus;
VII, facial nucleus; XII, hypoglossal
nucleus.
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SRC-1/
mice exhibited a moderate motor dysfunction
To explore the possible role of SRC-1 in brain development, the
general brain structure and histology of the adult
SRC-1/
mice were compared with their sex- and age-matched WT mice by staining
serial sections of the entire brain with hematoxylin and eosin (H&E) or
Nissl. However, no gross morphological defect was observed in the adult
brains of
SRC-1/
mice (data not shown). To assess the role of SRC-1 in brain function, we subsequently performed a battery of behavioral tests to compare specific neural functions between WT and
SRC-1/
mice. Motor coordination, strength, and balance were assessed by
hanging wire and rotarod tests. In the hanging wire test,
SRC-1/
mice fell 29% earlier than WT mice (Fig.
2A). Statistical
analysis showed significant differences (p = 0.03) in latencies to fall between WT and
SRC-1/
mice. The average body weight of
SRC-1/
mice is similar to WT mice (Xu et al., 1998), so differential weights
cannot account for the difference.
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Figure 2.
Behavioral analyses. A, Hanging
wire test. WT (n = 8) and
SRC-1/
(n = 11) mice were placed on bars and turned upside
down. The latency to fall was measured. B, Rotarod test.
WT (n = 20) and
SRC-1/
(n = 20) mice were trained three times per day for
a period of 3 d. Latency to fall from the accelerating rotarod was
measured. C, Morris water task. WT
(n = 10) and
SRC-1/
(n = 9) mice were trained with four trials per day
for 7 d. Mean escape latency was measured in the hidden platform
version of the Morris water task. D, Swim speed. WT
(n = 9) and
SRC-1/
(n = 9) mice were allowed to swim in the pool for
60 sec. Mean swim speed was measured from four consecutive
trials.
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In the rotarod test, a total of nine trials were recorded (Fig.
2B).
SRC-1/
mice exhibited identical latencies to fall in the first and last trials
of this experiment. However, a shorter latency to fall was detected
consistently in
SRC-1/
mice from trials 2 thru 9. For trials 2 and 3, WT mice significantly improved their performance and stayed on the rotarod for ~200 sec
(compared with 150 sec in trial 1). In contrast,
SRC-1/
mice did not improve and remained on the rotarod for the same amount of
time as in trial 1 (150 sec). In addition, WT mice rapidly reached the
maximum value of their latency to fall after four trials. Although
SRC-1/
mice gradually improved their performance in the rotarod test, they did
not reach a maximum value of latency until the ninth trial (Fig.
2B). These results suggest that SRC-1 is required for
normal motor learning.
Because SRC-1 was highly expressed in regions of the
hippocampus and amygdala, we assayed the animals in several
different learning and memory tests. During training in the Morris
water maze (Morris et al., 1982), the
SRC-1/
mice had significantly delayed escape latencies compared with WT mice
(F(1,17) = 8.3; p = 0.01) (Fig. 2C). The increased escape latency could result
from an impairment of learning and memory, or from a dysfunction of
motor skills required by the test. To assess spatial memory after
training, the platform was removed from the pool after the 28th trial.
Both WT and
SRC-1/
mice searched for the missing platform selectively during this probe
trial in the quadrant where the platform was located previously (data
not shown). Tests of cued and contextual fear conditioning and passive
avoidance also failed to reveal any impairment of learning and memory
(data not shown).
We next analyzed swim speed for WT and
SRC-1/
mice in the absence of the platform.
SRC-1/
mice showed a significantly slower swim speed compared with WT mice
(14.9 ± 0.8 vs 19.5 ± 0.6 cm/sec; p = 0.0002) (Fig. 2D). Therefore, the relatively poor
performance of the null mice during training in the Morris water task
can be attributed to a slower swim speed from a motor impairment.
Because of the limitations of the above tests, we have recognized the
difficulties in distinguishing a motor deficit from muscle weakness. In
an attempt to search for answers, we extracted RNA samples from the
femoral muscle and quantitatively measured the expression levels of the
SRC family members by performing TaqMan Real-Time RT-PCR. The relative
amounts of SRC-1, SRC-2, and SRC-3 mRNAs in the skeletal muscle were
4.6:1.0:267. Therefore, the total number of SRC-1 mRNA molecules in the
skeletal muscle (~2.4 molecules per nanogram total RNA) was
~60-fold lower than the number of SRC-3 mRNA molecules (~144
molecules per nanogram total RNA). Real-Time RT-PCR is an extremely
sensitive technique providing very accurate measurements. When in
situ hybridization was used, SRC-1 mRNA could not be detected in
the skeletal muscle (data not shown). Furthermore, although the SRC-3
mRNA level was much higher than SRC-1 and SRC-2 in the skeletal muscle
when measured by real-time RT-PCR, the level of SRC-3 expression in the
muscle was still lower than the detectable threshold of X-gal staining in the reporter mice with a LacZ knock-in (Xu et al., 2000).
Thus, it can be concluded that SRC-1 is expressed at extremely low
levels in the skeletal muscle, and therefore it is very unlikely that disruption of SRC-1 will directly affect the muscle strength. Collectively, our results from the hanging wire test, rotarod test,
Morris water task, and gene expression analyses suggest that
SRC-1-deficient mice exhibit a moderate motor dysfunction.
Expression of other members of the SRC family in the brain
Because
SRC-1/
mice exhibit only a moderate motor dysfunction, we reasoned that other
members of the SRC family may provide an overlapping function in the
brain. To address this possibility, the spatial expression patterns of
all SRC members in the brain were compared by in situ
hybridization. We found that SRC-2 mRNA was coexpressed with SRC-1 at
lower levels in multiple brain structures, including the olfactory
bulb, piriform cortex, hippocampus (CA1, CA2, and CA3 regions and
dentate gyrus), amygdala complex, hypothalamus, and cerebellum (Fig.
3) (data not shown). In contrast, SRC-3
mRNA was detected at low levels only in the CA1, CA2, and CA3 regions of the hippocampus (Fig. 3) (data not shown). These results demonstrate that all three SRC family members are coexpressed in the hippocampus, although their relative expression levels are different. SRC-1 and
SRC-2 have overlapping expression patterns in the adult mouse brain,
although in general, SRC-1 is expressed at higher levels.
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Figure 3.
Expression of SRC-2 and SRC-3 transcripts in the
hippocampus and cerebellum of
SRC-1/
mice. Expression of all SRC family members was analyzed by in
situ hybridization in the hippocampus (A, B, E, F, I,
J) and cerebellum (C, D, G, H, K, L).
Serial coronal sections were prepared from 10-week-old WT (A, C,
E, G, I, K) and
SRC-1/
(B, D, F, H, J, L) mice. Note that hybridization signals
in F, J, and H
(SRC-1/)
are stronger than those signals in E, I,
and G (WT), respectively. SRC-3
mRNA was undetectable in the cerebellum (K,
L). Arrows in C,
G, and H indicate the Purkinje cell
layer. Scale bar, 250 µm.
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Next, the question regarding whether the absence of the SRC-1 gene
product might alter the expression levels of SRC-2 and SRC-3 was
addressed. Interestingly, although SRC-2 was expressed at low levels in
the hippocampus and cerebellar Purkinje cells of WT mice, its
expression was observed at higher levels in those equivalent brain
regions of
SRC-1/
mice (Fig. 3, compare E, F and G,
H). In the hippocampus of
SRC-1/
mice, SRC-3 was also expressed at slightly higher levels than in WT
mice (Fig. 3, compare I, J); however, the
SRC-3 signal remained undetectable in the cerebellum of
SRC-1/
mice (Fig. 3K,L). Therefore, the
disruption of the SRC-1 gene results in an apparent upregulation of
other SRC family members in specific regions of the brain.
Delayed development of Purkinje cells in
SRC-1/
mice
Because the coexpression and upregulation of other SRC members in
the
SRC-1/
mouse brain may compensate for the loss of SRC-1 function during neural
development and correct early developmental defects at a later stage,
brain morphologies at earlier stages after birth including P0, P5, and
P10 were examined. Although no other abnormal structures in the brain
could be identified, the Purkinje cell layer in the cerebellum was
apparently missing at P0 on the
SRC-1/
cerebellar sections stained with H&E (data not shown). To confirm this
observation, immunostaining of calbindin, a Purkinje cell-specific marker, was performed (Anderson et al., 1998). In WT pups,
calbindin-positive Purkinje cells were detected at P0 and at all later
stages examined (Fig.
4A,B).
In contrast, the calbindin-positive cells were nearly undetectable at
P0 in
SRC-1/
mice (Fig. 4C). At P3, the calbindin-positive Purkinje cells in some
SRC-1/
pups became detectable on the midsagittal sections (data not shown). At
P5, the density and total number of Purkinje cells in
SRC-1/
pups were still significantly lower than those observed in WT pups. At
P10, the number of calbindin-positive Purkinje cells in
SRC-1/
mice finally reached comparable levels with their age-matched WT pups
(Fig. 4B,D). These observations
were further confirmed by the quantitative counting of
calbindin-positive Purkinje cells in the cerebellum of WT and
SRC-1/
mice at P0, P5, and P10 (Table 1). These
results indicate that Purkinje cell development is significantly
delayed at the neonatal stages in mice lacking SRC-1.
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Figure 4.
Delayed neonatal development of Purkinje cells in
SRC-1/
mice. Immunostaining with anti-calbindin antibody
(A-D) and with anti-synaptophysin antibody
(E-H) was performed on the midsagittal
cerebellar sections. Mouse ages and genotypes are as indicated. Note
that the calbindin-positive Purkinje cells in C and the
synaptophysin signal in G were undetectable. Scale bars,
50 µm.
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|
The development of synaptic connections of the Purkinje
cells was further examined by immunostaining of synaptophysin, an abundant glycoprotein localized in developing presynaptic termini (Leclerc et al., 1989). At P0, the immunostaining of synaptophysin was
undetectable in the cerebellum of
SRC-1/
mice but was clearly detected in Purkinje cells of wild-type mice (Fig.
4E,G). At P10, synaptophysin
immunostaining of the puncta around Purkinje cells and of the
connections with granule cells showed no gross differences between WT
and
SRC-1/
mice (Fig. 4F,H). This
observation is consistent with the results obtained from calbindin
staining described above. Collectively, these results prove that
disruption of the SRC-1 gene in the mouse brain delays Purkinje cell development.
Temporal expression of SRC coactivators during Purkinje
cell development
To define the temporal involvement of SRC coactivators in
origination and differentiation of Purkinje cells, the temporal expression patterns of the SRC family members during Purkinje cell
development were examined. SRC-1 mRNA was expressed in the cerebellar
primordium since 10.5 d.p.c. (Fig.
5A,B).
At P0, although the Purkinje cells had not lined into a clear cell
layer at this stage, high levels of SRC-1 mRNA were detected only in
the calbindin-positive Purkinje cells (Fig.
5C-E). At P10, SRC-1 expression was clearly detected in the Purkinje cell layer (Fig. 5F). In
contrast, SRC-2 mRNA was undetectable in Purkinje cells of WT mice at
P10 (Fig. 5G), whereas low levels of SRC-2 expression were
observed in the Purkinje cells of the adult brain in WT mice (Fig.
3G). Interestingly, in the Purkinje cells of
SRC-1/
mice, SRC-2 mRNA clearly became detectable at P10 (Fig.
5H), suggesting that SRC-2 was expressed earlier in
the Purkinje cells of
SRC-1/
mice. In addition, SRC-3 mRNA was undetectable in the Purkinje cells by
in situ hybridization. These results suggest that SRC-1 is
the major member of the SRC family to be involved in the Purkinje cell
development at prenatal and neonatal stages. The earlier expression of
SRC-2 in the Purkinje cells of
SRC-1/
mice at neonatal stage may replace SRC-1 to support Purkinje cell
differentiation.
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Figure 5.
Temporal expression of SRC-1 and SRC-2
mRNAs at early cerebellar developmental stages. Sagittal sections
through the cerebellar primordium at embryonic day (E) 10.5 (A, B) and the cerebellum at P0
(C-E) and P10 (F-H) were
prepared from WT (A-G) and
SRC-1/
pups (H). The expression of SRC-1
(A-F) and SRC-2 (G,
H) was examined by in situ
hybridization using 35S-labeled antisense RNA probes.
Images in A, C, and
F-H were photographed under dark-field
lighting. The bright-field images in B and
D were photographed from the same areas shown in
A and C, respectively.
Arrows in A and B indicate
the expression of SRC-1 in the cerebellar primordium. E
is a bright-field image of the premature cerebellum at P0 in higher
magnification, showing the colocalization of the immunostaining signals
of calbindin (brown color) and the in
situ hybridization signals of SRC-1 (small black
particles) in the Purkinje cells (arrows).
Arrows in F and H indicate
the expression of SRC-1 (F) and SRC-2
(H) in the Purkinje cell layer. Note that
SRC-2 expression was negative in WT pups at P10
(G) but positive in
SRC-1/
pups at the same stage (H). Scale bars:
A-D, 100 µm; E, 50 µm;
F-H, 300 µm.
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Birth of Purkinje cell precursors in
SRC-1/
mice
To search for the cellular mechanisms responsible for the delayed
development of Purkinje cells in SRC-1-deficient mice, several experiments were performed. First, programmed cell death was analyzed by performing terminal deoxynucleotidyl transferase-mediated
biotinylated UTP nick end labeling assays, and no increase of apoptotic
cells was detected in the cerebellum of
SRC-1/
mice at any developmental stage examined (data not shown). Second, the
differentiation potential of SRC-1-deficient Purkinje cells was
analyzed in an in vitro culture system (Furuya et al.,
1998). Cells were isolated from the cerebellum of 18-d-old embryos and treated with T3 and progesterone to induce differentiation in culture.
Successful differentiation of Purkinje cells in culture was verified by
calbindin immunostaining. After 6 d, large globular cell bodies of
Purkinje cells were observed in cultures derived from both WT
and SRC-1/ mice (Fig.
6A,D).
During 11-16 d in culture, both WT and SRC-1/ Purkinje
cells developed their typical mature morphology, exhibited as a large
globular cell body with thick apical dendrites and numerous branches
(Fig.
6B,C,E,F).
These results suggest that immature Purkinje cells from
SRC-1/
embryos can differentiate normally into mature Purkinje cells in
culture.
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Figure 6.
Purkinje cell differentiation in
culture. Cerebellar cells were isolated from 18-d-old WT
(A-C) and
SRC-1/
(D-F) mouse embryos. Purkinje cell bodies
and their dendritic branches at 6, 11, and 16 d in culture were
identified by immunocytochemical staining for calbindin. Scale bars, 50 µm.
|
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Finally, the time course for the generation of Purkinje cell precursors
during early embryonic development was studied. During mouse cerebellar
development, the precursors of Purkinje cells and neurons in deep
cerebellar nuclei originate in the ventricular zone during 11-13
d.p.c. These precursors start to migrate from the ventricular zone to
the location just beneath the proliferating granule cells during 14-17
d.p.c. (Hatten and Heintz, 1995). Because SRC-1 is expressed in the
cerebellar primordium (Fig. 5A) at early stages of
cerebellar development, the absence of the SRC-1 protein in
SRC-1/
mice may affect the generation of Purkinje cell precursors. To define
the developmental time course of Purkinje cell precursors in WT and
SRC-1/
mice, the birth of Purkinje cells was followed by BrdU labeling (Vogel
et al., 2000). Cerebellar precursor cells were labeled at 10.5, 12.5, or 13.5 d.p.c. by injecting a single dose of BrdU into SRC-1
heterozygous pregnant females. The incorporated BrdU in the nuclear DNA
was examined by immunostaining at P5 when calbindin-positive Purkinje
cells became reliably detectable in both WT and
SRC-1/
mice (Fig. 7A). A large number of BrdU-positive Purkinje
cells were clearly detected at P5 in WT pups that received BrdU at
10.5 d.p.c. In contrast, no BrdU-positive Purkinje cells could be
detected at P5 in
SRC-1/
littermates that received identical amounts of BrdU at 10.5 d.p.c. (Fig.
7Ba,Bb,C).
When BrdU was injected at 12.5 d.p.c., the number of BrdU-labeled
Purkinje cells at P5 in
SRC-1/-deficient
mice was threefold higher than in their WT littermates (Fig.
7Bc,Bd,C). However, when
BrdU was given at 13.5 d.p.c., only a small number of BrdU-labeled
Purkinje cells were observed at P5 in both WT and
SRC-1/
mice (Fig. 7Be,Bf,C). These
results demonstrate that the birth of Purkinje cell precursors in
SRC-1/
mice is delayed by ~2 d at early embryonic developmental stages, suggesting that the delayed Purkinje cell development at the neonatal stage is caused partially by the delayed birth of Purkinje cell precursors at earlier embryonic developmental stages.
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Figure 7.
Delayed generation of Purkinje cell precursors.
A, Time course for treatment and analysis. Precursor
cells were labeled by injecting a single dose of BrdU into multiple
pregnant SRC-1+/ females at 10.5, 12.5, and
13.5 d.p.c. Cerebellum samples were obtained at P5 for analysis of
BrdU-labeled Purkinje cells (PCs). B, The nuclei of
BrdU-labeled PCs on the midsagittal sections of cerebella were
identified by immunostaining with an antibody specific to BrdU. After
antibody reactions, the slides were counterstained either with
hematoxylin (a, b) or with methyl green
(c-f). Note that BrdU-positive PCs, indicated by
P, were observed primarily at E10.5 for WT mice
(a) and at E12.5 for
SRC-1/ mice (d).
Scale bars, 50 µm. C, Quantitative analysis
of BrdU-labeled PCs. BrdU-labeled PCs on three adjacent midsagittal
sections prepared from each cerebellum at P5 were counted. Each average
number of BrdU-labeled PCs per section for each genotype and each time
point was calculated from a total of 12 sections, which were prepared
from four mice. When BrdU was given at E10.5 (10.5 d.p.c.), the number
of BrdU-labeled PCs at P5 in WT mice was significantly higher than in
knock-out (ko) (SRC-1/) mice
(p < 0.001; t test).
However, when BrdU was given at E12.5, the number of BrdU-labeled PCs
at P5 in WT mice was significantly lower than in knock-out
(SRC-1/) mice (p < 0.001; t test). There were no statistical differences
between two groups when BrdU was given at E13.5.
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| |
Discussion |
The importance of hormones and their cognate NRs for the
development and function of the CNS caused us to investigate the potential roles and molecular mechanisms that the newly identified NR
coactivators have in the brain. In this study, a number of experiments
were designed to characterize the role of SRC-1 in brain development
and function. We first identified the potential sites of SRC-1 function
in the adult mouse brain by examining the expression of the SRC-1
transcript. We found that SRC-1 is preferentially expressed in specific
brain regions. Although the expression pattern of SRC-1 in the mouse
brain is similar to that observed in the rat brain (Meijer et al.,
2000), the identification of Purkinje cell-specific expression of SRC-1
mRNA and protein in the mouse cerebellum is a novel finding. These
SRC-1-positive brain regions contain high levels of multiple NRs (Lopes
da Silva and Burbach, 1995; Morimoto et al., 1996; Shughrue et al.,
1997). The overlapping of SRC-1 and NRs in these specific brain regions highlights their potential functional partnership in mediating hormone
actions. In addition, in the brain of mouse embryos, SRC-1 is expressed
much more broadly, with its highest expression localized in the
olfactory epithelium (Misiti et al., 1999). These results suggest that
SRC-1 function in the brain may be both region specific and
developmental stage specific.
Next, we addressed whether SRC-1 is required for generation and
maintenance of mature brain structure by analyzing the brain histology
of adult
SRC-1/
mice. Although serial brain sections were examined carefully, no
structural defects were observed in the brains of both male and female
adult
SRC-1/
mice. This observation indicates that SRC-1 is not essential for the
formation and maintenance of mature mouse brain morphology.
Subsequently, we compared the brain function of
SRC-1/
mice with age-matched WT mice using a battery of well established
behavior tests.
SRC-1/
mice performed poorly in the hanging wire tests, suggesting that SRC-1/
mice may have a dysfunction of motor strength. In the rotarod tests,
although
SRC-1/
mice initially showed coordination comparable with that of WT mice,
serial trials revealed that
SRC-1/
mice made significantly slower improvement in their performance on the
rotarod, suggesting that the motor learning and the motor performance
of
SRC-1/
mice are partially impaired. In the Morris water maze,
SRC-1/
mice exhibited significantly longer escape latencies when compared with
WT mice, which was primarily attributable to a slower swim speed, as
demonstrated by further analysis. Because SRC-1 is expressed at
extremely low levels in the skeletal muscle, the sum of these results
suggests that
SRC-1/
mice have a moderate motor dysfunction.
No other abnormalities in brain function, such as dysfunction of
spatial learning, could be detected in
SRC-1/
mice, although SRC-1 was highly expressed in functional regions including hippocampus and amygdala. Because in vitro studies
have indicated that SRC coactivators have overlapping function in
coactivation of multiple transcription factors when expressed in the
same type of cells (Xu and O'Malley, 2002), and we have reported that
SRC-2 mRNA is more abundant in total RNA samples from
SRC-1/
mouse brains (Xu et al., 1998), we have hypothesized that other members
of the SRC family may compensate for the loss of SRC-1 during brain
development and function.
To test our hypothesis, spatial distributions and relative expression
levels for all three SRC coactivators in brain regions of both WT and
SRC-1/
mice were compared in parallel experiments. Our results demonstrated the following. (1) In WT mice, SRC-1 is more abundantly expressed than
other SRC family members in multiple brain regions described above.
SRC-2 is expressed at lower levels in all brain regions where SRC-1 is
expressed, except in the brainstem. SRC-3 mRNA is detectable at a very
low level only in the hippocampus by in situ hybridization.
By using a much more sensitive method (X-gal staining), SRC-3
expression can also be detected in the olfactory bulb in addition to
the hippocampus (Xu et al., 2000). (2) In SRC-1/
mice, SRC-2 expression is slightly elevated, at least in the hippocampus and cerebellum. SRC-3 expression is also slightly elevated
in the hippocampus. When one or more of the family members are
disrupted, the partially overlapping expression of SRCs and upregulation of other SRC members may provide compensatory mechanisms to support brain development and function. The coexpression patterns of
all SRCs in the hippocampus of WT mice may explain, at least in part,
why
SRC-1/
mice can still maintain their normal hippocampal function in the
learning and memory tests. The notion of functional redundancy among
SRC family members is further supported by the lethal phenotype of the
double SRC-1 and SRC-2 knock-out mice, although each individual knock-out mouse line shows nearly normal life span (data not shown).
Purkinje cells serve as the sole output neurons for the cerebellar
cortex and affect motor neuron activity. Motor learning tasks are
mediated by Purkinje cells in the cerebellum and are frequently
implicated in altered function of Purkinje cells (Linden, 1994; Raymond
et al., 1996). Because SRC-1 is highly and specifically expressed in
Purkinje cells of the cerebellum, the lack of SRC-1 may affect Purkinje
cell function. Therefore, the moderate dysfunction of motor learning
observed in adult
SRC-1/
mice can be attributed to the absence of SRC-1 in Purkinje cells. However, it is also possible that motor dysfunction in adulthood is a
result of developmental problems at early stages. To test this
possibility, we examined brain development at late embryonic and
neonatal stages, with our focus on the brain regions expressing high
levels of SRC-1. Indeed, a delay in Purkinje cell development was
identified at the neonatal stage in
SRC-1/
mice. Both calbindin and synaptophysin molecular markers for Purkinje
cells were undetectable in the cerebella of
SRC-1/
pups at P0 although these two molecules were detected in their WT
littermates as expected (Leclerc et al., 1989; Anderson et al., 1998).
Analysis of the time course for the origination of Purkinje cell
precursors by BrdU labeling made it clear that the delay in Purkinje
cell development at the neonatal stage is related to a deficit in the
generation of precursor cells during 10.5-12.5 d.p.c. in
SRC-1/
embryos. During this developmental stage, SRC-1 is expressed in the
cerebellar primordium in WT embryos, suggesting that removal of SRC-1
from this region may directly affect the generation of Purkinje cell
precursors. However, once the precursors of Purkinje cells in
SRC-1/
embryos were generated, they were able to differentiate into Purkinje
cells as WT precursors did when analyzed in a culture system (Maeda et
al., 1989; Nakanishi et al., 1991).
The time course analysis revealed that the generation of
SRC-1/
Purkinje cell precursors was delayed ~2 d when compared with WT
precursors. In
SRC-1/
pups at neonatal stages, the calbindin-positive Purkinje cells were
nearly undetectable at P0 and became detectable at P3. Even at P5, the
number of calbindin-stained Purkinje cells in
SRC-1/
pups was still significantly lower than that in WT pups. The number of
Purkinje cells in
SRC-1/
mice finally reached that of their WT littermates during P5-P10. These
observations indicate that the time differences between WT and
SRC-1/
pups with respect to the final morphological maturation of Purkinje cells after birth are longer than the delay in the generation of
Purkinje precursors. To uncover the underlying mechanisms, we further
compared the temporal expression patterns of SRC coactivators in
Purkinje cells. Our analysis indicates that SRC-1 transcripts exist in
cerebellar primordium in embryos and in Purkinje cells after birth, but
SRC-2 mRNA was undetectable in Purkinje cells even at P10. Similar to
adult mice, SRC-3 was undetectable in the cerebellum at these stages by
in situ hybridization. The different temporal expression
patterns of SRCs suggest that SRC-1 is the major player in the SRC
family for Purkinje cell development at embryonic and neonatal stages,
and SRC-2 may participate at a later stage. More interestingly, SRC-2
mRNA became detectable at P10 in
SRC-1/
Purkinje cells, suggesting that this earlier expression of SRC-2 in
SRC-1/
Purkinje cells might play a compensatory role to support the maturation
of Purkinje cells in
SRC-1/ mice.
The critical roles of ROR and TH in Purkinje cell development have
been well established. ROR heterozygous mutant mice show accelerated
dendritic atrophy and cell loss, suggesting that ROR has a role in
mature Purkinje cells (Zanjani et al., 1992). ROR homozygous mutants
exhibit severe tremors and body imbalance caused by a cell-autonomous
defect in the development of Purkinje cells (Hamilton et al., 1996;
Dussault et al., 1998). These ROR-deficient Purkinje cells show
immature morphology and synaptic arrangement and a reduction in
numbers. Hypothyroidism causes reduced dendritic arborization of
Purkinje cells and less synaptogenesis at neonatal stage similar to
that seen in ROR mutant mice (Koibuchi and Chin, 2000).
Interestingly, TH treatment alters the timing of expression of the
ROR gene in Purkinje cells, and ROR mutation blocks Purkinje cell
response to TH (Messer, 1988; Koibuchi and Chin, 1998). Disruption of
SRC-1 may affect Purkinje cell development and maturation caused by a
partial impairment of the ROR and TR functions on the basis of the
following findings: (1) the members in the SRC family serve as
transcriptional coactivators for ROR and TR (Takeshita et al., 1996,
1997; Atkins et al., 1999); (2) SRC-1 is coexpressed with ROR and TR
in the Purkinje cells; (3) only SRC-1 in the SRC family is expressed in
the cerebellar primordium and Purkinje cells during embryonic and
neonatal stages; and (4) the SRC-1-deficient mice exhibit thyroid
hormone resistance (Weiss et al., 1999). In addition to NRs, SRC-1
deficiency may also affect the function of some other transcription
factors during Purkinje cell development because SRC-1 also interacts
with and augments a number of non-NR transcription factors such as
AP-1, serum response factor, and NF- B (Xu and O'Malley, 2002).
Characterization of the specific molecular targets of SRC-1 and the
degree of the functional impairments of related transcription factors
in
SRC-1/
mice during Purkinje cell development deserve additional studies.
In summary, the expression of individual SRC members in the mouse brain
is spatially and temporally regulated. Their partial overlapping
expression patterns may determine their functional redundancy
during brain development and function. In the cerebellum, SRC-1 is
selectively expressed in Purkinje cells. Disruption of the SRC-1 gene
in mice delays the generation of Purkinje cell precursors at an early
embryonic stage and further delays the maturation of Purkinje cells
after birth. Although the morphology of
SRC-1/
Purkinje cells develops to the same extent as WT Purkinje cells by P10,
adult
SRC-1/
mice still exhibit moderate motor dysfunction, suggesting that the
abnormal development of Purkinje cells at early stage may have a
negative impact on the cerebellar function in adulthood. For the first
time, these findings demonstrate that SRC coactivators play an
important role in cerebellar Purkinje cell development and motor function.
| |
FOOTNOTES |
Received June 24, 2002; revised Oct. 7, 2002; accepted Oct. 14, 2002.
This work was supported by National Institutes of Health Grants
U54-HD-07495 (B.W.O.) and DK58242 (J.X.), by National Institute of Mental Health Grant IMH 60420 (R.L.D.), and by a Research Award (E.N.) from the Uehara Memorial Foundation. We thank Drs. M.-J. Tsai,
F. DeMayo, H. Y. Zoghbi, F. A. Pereira, and N. J. McKenna for valuable discussions.
Correspondence should be addressed to Dr. Jianming Xu, Department of
Molecular and Cellular Biology, Baylor College of Medicine. One Baylor
Plaza, Houston, TX 77030. E-mail:
jxu{at}bcm.tmc.edu.
| |
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TOP
ABSTRACT
Introduction
