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 Previous Article | Next Article 
The Journal of Neuroscience, October 1, 2001, 21(19):7526-7533
Involvement of Nitric Oxide Released from Microglia-Macrophages
in Pathological Changes of Cathepsin D-Deficient Mice
Hiroshi
Nakanishi1,
Jian
Zhang2,
Masato
Koike3,
Tsuyoshi
Nishioku2,
Yoshiko
Okamoto4,
Eiki
Kominami5,
Kurt
von
Figura6,
Christoph
Peters7,
Kenji
Yamamoto2,
Paul
Saftig6, and
Yasuo
Uchiyama3
Laboratory of 1 Oral Aging Science and
2 Biochemical and Molecular Pharmacology, Division of Oral
Biological Sciences, Faculty of Dental Sciences, Kyushu University,
Fukuoka 812-8582, Japan, 3 Department of Cell Biology and
Neuroscience, Osaka University Graduate School of Medicine,
Suita, Osaka 565-0871, Japan, 4 Department of
Physiological Chemistry, Daiichi Pharmaceutical University, Fukuoka
815-8511, Japan, 5 Department of Biochemistry, Juntendo
University School of Medicine, Tokyo 113-0033, Japan,
6 Center for Biochemistry and Molecular Cell Biology,
Göttingen University, Heinrich-Düker Weg 12, 37073 Göttingen, Germany, and 7 Institut für
Molekulare Medizin und Zellforschung, Albert-Ludwings-Universität
Freiburg Hugstetter Strasse 55, 79106 Freiburg, Germany
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ABSTRACT |
Cathepsin D (CD) deficiency has been shown to induce
ceroid-lipofuscin storage in lysosomes of mouse CNS neuron (Koike et al., 2000 ). To understand the behavior of microglial cells
corresponding to these neuronal changes, CD-deficient (CD /) mice,
which die at approximately postnatal day (P) 25 by intestinal necrosis, were examined using morphological as well as biochemical approaches. Light and electron microscopic observations revealed that microglia showing large round cell bodies with few processes appeared in the
cerebral cortex and thalamus after P16. At P24, microglia often
encircled neurons that were occupied with autolysosomes, indicating increased phagocytic activity. These morphologically transformed microglia markedly expressed inducible nitric oxide synthase (iNOS), which was also detected in the intestine of the mice.
To assess the role of microglial nitric oxide (NO) in neuropathological changes in CD/ mice,
L-NG-nitro-arginine
methylester (L-NAME), a competitive NOS inhibitor, or S-methylisothiourea hemisulfate (SMT), an iNOS inhibitor, was administered intraperitoneally for 13 consecutive days. The total number of terminal deoxynucleotidyl transferase-mediated
biotinylated UTP nick end labeling-positive cells counted in the
thalamus was found to be significantly decreased by chronic treatment
of L-NAME or SMT, whereas neither the neuronal accumulation
of ceroid-lipofuscin nor the microglial phagocytic activity was
affected by these treatments. Moreover, the chronic treatment of
L-NAME or SMT completely suppressed hemorrhage-necrotic
changes in the small intestine of CD/ mice, resulting in normal
growth of the body weight of the mice. These results suggest that NO
production via iNOS activity in microglia and peripheral macrophages
contributes to secondary tissue damages such as neuronal apoptosis and
intestinal necrosis, respectively.
Key words:
cathepsin D-deficient mouse; microglia; nitric oxide; L-NG-nitro-arginine
methylester; apoptotic neuron; intestinal atrophy
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INTRODUCTION |
It has been shown that
cathepsin D (CD; EC 3.4.23.5)-deficient (CD/) mice, which are born
normally but die at postnatal day (P) 26 ± 1 with signs of
massive intestinal necrosis, thromboembolia, and lymphopenia (Saftig et
al., 1995), exhibit prominent neuropathological changes in CNS (Koike
et al., 2000). The mice suffering from seizures and blindness
demonstrate an accumulation of lysosomes with granular osmiophilic
deposits and fingerprint profiles in CNS neurons, which are features
typical of neuronal ceroid lipofuscinoses (NCLs) (Dawson and Cho,
2000). White Swedish Landrace sheep expressing an inactive form of the
point-mutated CD molecule have been found to show a severe loss of
neurons in the cortical area and hippocampus and accumulation of
protein-like storage materials in CNS neurons (Tyynelä et al.,
2000). Subunit c of the mitochondrial F1F0ATPase, a common storage
material of NCL except for the infantile form of NCL, is markedly
accumulated in both the CNS and peripheral cells of CD/ mice (Koike
et al., 2000). The loss of CD activity causes a novel type of
lysosomal storage disease associated with massive neurodegeneration.
The mechanism underlying neuronal damage and death, however, remains unknown.
Microglia are normally present as ramified cells that have small cell
bodies with numerous branching processes. Once fully activated,
ramified microglia are morphologically transformed into cells that are
characterized by large cell bodies with few processes and phagocytic
activity. As an intermediate form, microglia appear to have
large cell bodies with several thicker processes. These activated
microglia appear to be directly involved in propagation of
neuropathological events such as Alzheimer's disease because the
microglial activation leads to produce mediators of
inflammation-mediated neurodegeneration including nitric oxide (NO) and
tumor necrosis factor- (TNF-) (Meda et al., 1995; Brown et al.,
1996; Yan et al., 1996; Barger and Harmon, 1997; Tan et al., 1999;
Tanabe et al., 1999; Kim et al., 2000; Wada et al., 2000). The direct
involvement of NO in microglia-induced neuronal death has also been
investigated in in vitro (Boje and Arora, 1992; Chao et al.,
1992) and in vivo (Takeuchi et al., 1998; Matsuoka et al.,
1999) studies. However, little is known about the relationship between
microglia-induced inflammation and neurodegeneration associated with CD deficiency.
The present study attempted to characterize microglia in CNS tissues of
CD/ mice and elucidate its involvement in neurodegenerative change.
We found a prominent expression of inducible NO synthase (iNOS) in both
microglia and peripheral macrophages. The chronic treatment of
L-NG-nitro-arginine
methylester (L-NAME), a competitive NOS
inhibitor, and S-methylisothiourea hemisulfate (SMT), an iNOS
inhibitor, revealed that NO production via iNOS activities of microglia
and peripheral macrophages was significantly but differentially
associated with pathological changes in the CNS and intestine.
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MATERIALS AND METHODS |
Animals. Heterozygous (+/) mice (Saftig et al.,
1995) were transferred to the Institute of Experimental Animal
Sciences, Kyushu University Faculty of Dentistry, and kept in
conventional facilities. Selection of CD/ mice from littermates
obtained by heterozygous coupling was performed according to the method of Saftig et al. (1995) in which template genomic
DNA that was isolated from tail biopsies was examined by CD exon
4-specific PCR with primers of MCD14
(5'-AGACTAACAGGCCTGTTCCC-3') and MCD15 (5'-TCAGCTGTAGTTGCTC AC ATG-3').
Heterozygous mice that were used as control animals in the
present study show no pathological phenotypes when examined by
histological, immunocytochemical, and biochemical methods. The day of
birth was designated as P1.
Immunohistochemistry. Detailed immunohistochemical procedure
has been described previously (Nakanishi et al., 1993, 1994, 1997).
Briefly, specimens were obtained at P16, P20, and P24 from CD/ mice and control littermates that were anesthetized with sodium
pentobarbital (40 mg/kg, i.p.) and killed by intracardiac perfusion
with isotonic saline, followed by a chilled fixative consisting of 4%
paraformaldehyde in 0.2 M PBS, pH 7.4. After perfusion, the brain was removed, further fixed by immersion in the
same fixative overnight at 4°C, and then immersed in 20% sucrose, pH
7.4, for 24 hr at 4°C. Floating parasagittal sections (30 µm thick)
of the cerebral cortex and the thalamus were prepared by a cryostat and
stained by F4/80 (100 µg/ml) (Serotec, Oxford, UK), anti-GFAP
IgG (2.0 µg/ml) (Dako Japan, Kyoto, Japan) or anti-iNOS IgG (1.25 µg/ml) (Transduction Laboratories, Lexington, KY) with the
avidin-biotin-peroxidase complex method as described previously. After PBS washes, sections were reacted with 0.025%
3',3-diaminobenzidine/0.4% (NH4)2Ni(SO4)2/0.09%
H2O2/0.1
M Tris-buffered saline solution for 5-10 min.
All sections were thoroughly rinsed with PBS, mounted, and
coverslipped. As immunohistochemical control, the sections were
incubated with nonimmune rabbit IgG or mouse IgG and followed by the
same treatments described above.
For indirect fluorescent immunohistochemistry, floating parasagittal
sections (30 µm thick) of the cerebral cortex and the thalamus were
stained with F4/80 (200 µg/ml) and anti-GFAP IgG (8.2 µg/ml) for 90 min at 37°C. After a wash with PBS, F4/80 and anti-GFAP IgG-treated
sections were stained with biotinylated anti-mouse and anti-rabbit IgG,
respectively. After washes with PBS, sections were treated with 0.5%
streptavidin-Alexa 488 (Molecular Probes, Eugene, OR) for 2 hr at room
temperature. After wash with PBS, the sections were mounted in the
anti-fading medium. For double staining, floating parasagittal sections
(30 µm thick) of the cerebral cortex and the thalamus were stained
with anti-iNOS IgG (2.5 µg/ml) for 90 min at 37°C. After a wash
with PBS, the sections were treated with 0.5% anti-mouse IgG
conjugated with fluorescein isothiocyanate (Amersham Pharmacia
Biotech, Buckinghamshire, UK) for 2 hr at room temperature. To
identify microglia in the iNOS-stained sections, we treated them
further with biotinylated Griffonia Simplicifolia BS-I isolectin B4 (10 µg/ml) (GSA-I-B4; Sigma, St. Louis, MO) overnight at 4°C. After a
wash with PBS, the sections were stained with 0.5% streptavidin-Cy3
(1:100) (Amersham Pharmacia Biotech) for 2 hr at room temperature. To
identify astrocyte in the iNOS-stained sections, we treated them
further with anti-GFAP IgG (2.0 µg/ml) (Dako Japan) overnight at
4°C. After a wash with PBS, the sections were stained with 0.5%
streptavidin-Cy3 (Amersham Pharmacia Biotech) for 2 hr at room
temperature. The sections were mounted in the anti-fading medium
Vectashield (Vector Laboratories, Burlingame, CA).
Electron microscopy. CD-/ mice at P18 and P25 were
deeply anesthetized with sodium pentobarbital (25 mg/kg, i.p.) and
fixed by cardiac perfusion with 2% paraformaldehyde/2% glutaraldehyde buffered with 0.1 M phosphate buffer, pH 7.2. After perfusion, the brain was removed and further immersed in the same
fixative at 4°C overnight. After being washed thoroughly with the
same buffer, containing 7.5% sucrose, samples were post-fixed with 1%
OsO4 with the same buffer, containing 7.5%
sucrose, at 4°C for 2 hr; the brain was block-stained with 2%
aqueous solution of uranyl acetate for 1 hr. Then, the brain was
dehydrated with a graded series of ethanol and embedded in Epon 812. Serial ultrathin sections were cut with the ultramicrotome, stained
with lead citrate and uranyl acetate, and observed with a Hitachi
H-7100 electron microscope as described previously (Koike et al.,
2000).
Assay of NO2
accumulation. Specimens were obtained at P16, P20, and P24
from CD/ mice and control littermates that were anesthetized with
sodium pentobarbital (40 mg/kg, i.p.), and the brains and the small
intestines were rapidly removed. The whole brain and the ileum were
homogenized in 50 mM Tris-HCl, pH 7.4, containing
0.5 mM EGTA, 0.5 mM EDTA.
After centrifugation at 50,000 × g for 30 min, the
supernatant was used as the cytosolic fraction. The supernatant was
transferred to 96-well dishes with Griess reagent (Griess
Reagent Kit; Dojindo, Kumamoto, Japan) and incubated at room
temperature for 15 min. The amount of
NO2 in the cytosol was
measured spectrophotometrically by using an ELISA plate reader (model
550; Bio-Rad, Richmond, CA) with the absorbance at 540 nm.
SDS-gel electrophoresis and immunoblotting. Detailed
immunoblotting procedure has been described previously (Nakanishi et al., 1994; Amano et al., 1995). The whole brain and the ileum obtained from CD/ mice and control littermates at P16, P20, and P24 were anesthetized with sodium pentobarbital (40 mg/kg, i.p.)
and killed by intracardiac perfusion with isotonic saline. The soluble
fractions obtained from the whole brain and the ileum homogenates were
electrophoresed in SDS-polyacrylamide gels. For immunoblotting,
proteins on SDS gels were transferred electrophoretically at 100 V for
12-15 hr from the gels to nitrocellulose membranes and then incubated
at 4°C overnight under gentle agitation with the following primary
antibodies: anti-iNOS IgG (0.1µg/ml) and anti-nNOS IgG (0.1 µg/ml).
After washes, the membranes were incubated with 0.5% horseradish
peroxidase (HRP)-labeled horse anti-mouse IgG (Amersham Pharmacia
Biotech). Subsequently, membrane bound HRP-labeled antibodies were
detected by the enhanced chemiluminescence detection system (ECL kit;
Amersham Pharmacia Biotech) on x-ray film (X-Omat; Eastman Kodak,
Rochester, NY) 30-60 sec after exposure. As a control, the primary
antibody was replaced by nonimmune mouse IgG. The protein bands on
x-ray film were scanned and densitometrically analyzed by a
densitometer (Personal Scanning Imager PD110; Molecular Dynamics).
Procedure for L-NAME and SMT
administration. CD/ mice were divided into three subgroups and
intraperitoneally treated with saline, L-NAME (10 mg/kg; Affinity BioReagents, Golden, CO), a competitive NOS inhibitor,
and SMT (10 mg/kg; Research Biochemicals, Natick, MA), an iNOS
inhibitor, twice a day for 13 consecutive days. The injections began on
P12. The injection of saline to CD/ mice was stopped at P23 because
five of eight CD/ mice were dead by P23, and the condition
of the remaining animals was very serious at this stage. The wild-type
littermates were also treated with saline for 13 consecutive
days. After 12 hr of the final administration, mice were
killed by intracardiac perfusion with isotonic saline, followed by 4%
paraformaldehyde under pentobarbital anesthesia. After perfusion, the
brain and the small intestine were removed, further fixed by immersion
in the same fixative overnight at 4°C, and then immersed in 20%
sucrose for 24 hr at 4°C. Floating parasagittal sections (30 µm
thick) of the thalamus and coronal sections (30 µm thick) of the
small intestine were prepared by a cryostat. Hematoxylin-eosin staining
was performed according to standard procedures.
Terminal deoxynucleotidyl transferase-mediated biotinylated
UTP nick end labeling. Thalamic sections prepared from
CD/ mice at P23, CD/ mice at P24 treated with
L-NAME, and age-matched wild-type control were
treated with 2% H2O2 in
PBS for 20 min at room temperature. After washes in PBS, sections were
stained for terminal deoxynucleotidyl transferase-mediated biotinylated UTP nick end labeling (TUNEL) using ApopTag kit (Intergen, Purchase, NY), following the protocol provided by the manufacturer.
Statistical analysis. Data are expressed as means ± SD. The statistical analyses of the data in Figures 3A, 6,
and 9A were performed by Student's t test. The
statistical analysis of the data in Figure 7 was performed by two-way
ANOVA, followed by Scheffe's post hoc test for multiple
comparison when F ratios reached significance.
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RESULTS |
Morphological transformation of microglia in the CNS of
CD/ mice
To examine the morphological transformation of microglia during
the course of neuropathological changes in CNS tissues of CD/ mice,
brain sections from the mice were immunostained with F4/80, a marker
for microglia-macrophages. At P16 of CD/ mice, a proliferation of
F4/80-immunoreactive microglia was first observed especially in the
cerebral cortex (Fig.
1A) and the thalamus
(Fig. 1B). Processes of microglia were usually
attached to the cell bodies of large neurons in these brain regions.
These cells were considered to be activated microglia because they had
shortened and thicker processes. After P20, most microglial cells
exhibiting expanded and round cell bodies with few thick processes
appeared in the entire brain of CD/ mice. F4/80 immunoreactivity
formed an almost continuous rim in both the cortex (Fig.
1C,E) and the thalamus (Fig.
1D,F), suggesting that
microglia phagocytosed cortical and thalamic neurons. The number of
these phagocytic microglia that were characterized by their large round
cell bodies with few thick processes was dramatically increased in an
age-dependent manner; the mean incidence in the thalamus (mean ± SD cells per section; n = 5 or 6) was 31 ± 13 at
P16, 67 ± 10 at P20, and 121 ± 13 at P24. By contrast,
ramified microglia were only detected in these brain regions of control
littermates even at P24 (Fig. 1G,H).
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Figure 1.
Accumulation of activated microglia stained with
F4/80 in the cerebral cortex and the thalamus from CD/ mice.
A, C, E, G, the cortex;
B, D, F, H, thalamus.
A, B, CD/ mouse at P16;
C, D, CD/ mouse at P24;
E, F, enlargement of boxed
areas in C and D, respectively;
G, H, control littermate mouse at P24.
Scale bars: A, 50 µm; E, 20 µm.
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Phagocytosis of ceroid-lipofuscin-laden neurons by microglia
We have shown previously that neuronal cell bodies with
autofluorescent bodies increased in number in various brain regions of
CD/ mice (Koike et al., 2000). In the brain sections prepared from
CD/ mice at P24, large autofluorescent bodies were evident in the
cell bodies of neurons especially in the cerebral cortex and the
thalamus of CD/ mice. To further evaluate the possible phagocytosis
of these ceroid-lipofuscin-laden neurons by microglia, we
immunostained these sections by glial cell markers, F4/80 and anti-GFAP
antibody. Under fluorescent microscope, F4/80-labeled microglia
(green) were observed to uptake neurons laden with
large autofluorescent bodies (orange) in the cortex (Fig.
2A) and the thalamus
(Fig. 2B). On the other hand, GFAP-labeled astrocytes showed no phagocytic activity in either the cortex (Fig.
2B) or the thalamus (Fig. 2C). At the
electron microscopic level, microglia often enclosed a large area of
neuronal cell bodies that were characterized by the presence of a
number of autophagosome-autolysosome-like structures containing part
of the cytoplasm or heterogenously dense materials as reported
previously (Koike et al., 2000) (Fig. 2E).
Furthermore, microglia that completely surrounded pale neurons by their
cytoplasmic processes were also often encountered, suggesting progressive microglial phagocytosis of neurons (Fig.
2F). These observations clearly suggest that
phagocytosis of ceroid-lipofuscin-laden neurons is responsible for
morphological transformation of microglia in CNS of CD/ mice.
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Figure 2.
Phagocytosis of neurons laden with
ceroid-lipofuscin by microglia in the cortex and the thalamus of
CD/ mice. A-D, Immunostaining using
F4/80 and anti-GFAP antibody in the cortex (A, C) and
the thalamus (B, D) of CD/ mouse at P24.
A, B, Immunostaining for F4/80
(green) and autofluorescence of
ceroid-lipofuscin (orange). C,
D, Immunostaining for GFAP (green)
and autofluorescence of ceroid-lipofuscin (orange).
It was noted that microglia labeled by F4/80 extensively
engulfed neurons that were laden with large autofluorescent bodies,
whereas astrocytes labeled by anti-GFAP antibody showed no
phagocytic activity. E, F, Electron
micrographs of microglia attached and phagocytosed neurons laden with
autophagosome-autolysosome-like bodies in the cortex
(E) and the thalamus (F) of
CD/ mouse at P24. m, Microglia; n,
neuron. Scale bars: A, 20 µm;
E, 4.0 µm; F, 3.5 µm.
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Accumulation of NO2 and expression
of iNOS in microglia
Microglia have been suggested to be directly involved in
propagation of neuropathological events via production of NO (Boje and
Arora, 1992; Chao et al., 1992; Meda et al., 1995; Brown et al., 1996;
Yan et al., 1996; Barger and Harmon, 1997; Takeuchi et al., 1998;
Matsuoka et al., 1999; Tan et al., 1999; Kim et al., 2000; Wada et al.,
2000). For this reason, we measured the accumulation of
NO2, a major metabolite
of NO, in the cytosolic fraction of the brain of CD/ mice. At P20,
concentrations of NO2
measured in the cytosolic fractions of the whole brain in CD/ mice
were significantly higher than those from wild-type littermate mice and
further increased at P24. At P24, the level of
NO2 that was measured
from CD/ mice was approximately threefold higher than that from the
wild-type littermate mice (Fig.
3A). Additionally, the
expression level of iNOS in the cytosolic fraction of the brain of
CD/ mice was examined by Western blotting. A single band
with a molecular mass of ~130 kDa indicating iNOS immunoreactivity
was observed in the soluble fraction of the brain from CD/ mice at
P20 (Fig. 3B). At P24, the level of iNOS was increased by
approximately twofold. In the soluble fraction of the wild-type
littermate mice, no band corresponding to iNOS was detectable.
Moreover, a single 160 kDa band representing nNOS was observed in the
soluble fraction of both CD/ mice and control littermates at P20.
The level of nNOS showed no age-related change (Fig.
3B).
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Figure 3.
Expression of iNOS in the brain of CD/ mice.
A, Accumulation of NO2
in the cytosolic fraction of the whole brain of CD/ mice (/)
and control littermates (+/+) at P20 and P24. *p < 0.05, **p < 0.01, when compared with control
littermates (Student's t test). B,
Alteration of iNOS and nNOS proteins in the soluble fraction of the
whole brain of CD/ mice (/) and control littermates (+/+) at
P20 and P24. Arrowheads indicate the iNOS protein with
molecular mass of 130 kDa and nNOS protein with molecular mass of 160 kDa.
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Immunohistochemical localization of iNOS
To clarify the localization of iNOS in the brain of CD/ mice,
we conducted immunohistochemical staining using anti-iNOS antibody.
Immunoreactivity for iNOS was observed especially in the cerebral
cortex and the thalamus from CD/ mice after P16. Positive staining
for iNOS in these regions at P16 was localized in activated
microglia-like cells characterized by their shortened and thicker
processes. At P24, phagocytic microglia-like cells that accumulated in
both the cerebral cortex (Fig.
4C) and the thalamus (Fig.
4D) were intensely stained by anti-iNOS antibody. The
number of iNOS-positive microglia in the thalamus of CD/ mice
(mean ± SD cells per section; n = 4 or 5) was
158 ± 26 at P16, 184 ± 16 at P20, and 216 ± 22 at
P24. On the other hand, no immunoreactivity for iNOS was detected
either in the cerebral cortex (Fig. 4A) or the
thalamus (Fig. 4B) of the wild-type littermate mice.
To identify the iNOS-positive cells, iNOS-stained sections of the
thalamus were treated further with glial cell markers. At P24, the
iNOS-positive cells (Fig. 4E) corresponded well with GSA-I-B4-positive microglia (Fig. 4G), whereas iNOS (Fig.
4F) was not expressed in astrocytes identified by
anti-GFAP antibody (Fig. 4H). These observations
clearly indicate that iNOS is exclusively expressed in activated
microglia.
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Figure 4.
Immunostaining for iNOS in the brain.
A-D, Immunostaining using anti-iNOS
antibody in the cerebral cortex (A, C) and the thalamus
(B, D) of the CD/ mice and control littermates at
P24. A, B, control; C,
D, CD/. E-H, Double
immunostaining for iNOS (E) and GSA-I-B4
(G) in the same thalamus section of CD/ mouse
at P24. Double immunostaining for iNOS (F) and
GFAP (H) in the same thalamus section of
CD/ mouse at P24. Scale bars: A, 50 µm;
E, 20 µm.
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Effects of L-NAME and SMT on neurodegeneration in CNS
of CD/ mice
Although NO is known to work as a biologic messenger molecule
regulating immune function and blood vessel dilation and serving as a
neurotransmitter in both CNS and peripheral nervous system, a sustained
production and high levels of NO have cytotoxic properties (Lowenstein
and Snyder, 1992; Schmidt and Walter, 1994). We next analyzed for
neuronal death in the described brain regions of CD/ mice because
iNOS expressed in microglia is thought to be the isoform that produces
the large quantities of NO (Nathan and Xie, 1994). In the CNS of the
wild-type littermate mice, TUNEL-positive cells were found to be
densely distributed in the dentate gyrus of the hippocampus, the
olfactory bulb, and the subependyma in which apoptosis plays the
opposite role to neurogenesis in the regulation of cell numbers.
Besides these three brain regions, TUNEL-positive cells were found in
CD/ mice particularly in the thalamus, where phagocytic
iNOS-positive microglia accumulated most densely. At P16, only a few
TUNEL-positive cells (ranging from two to five cells per section of the
thalamus) were observed in CD/ mice. At P24, TUNEL-positive cells
were dramatically increased (24 ± 7 cells per section of the
thalamus; n = 9) and mainly distributed in the thalamus
(Figs. 5B, 6). The majority of
TUNEL-positive cells in the thalamus were
considered to be neurons because of their relatively large cell size
and location. In control littermates, TUNEL-positive cells were hardly
detectable in the thalamus, even at P24 (Fig. 5A).
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Figure 5.
TUNEL-stained sections in the thalamus and the
dentate gyrus. A, The thalamus of control littermates at
P24; B, thalamus of CD/ mice at P24;
C, thalamus of CD/ mice treated with
L-NAME at P24; D, dentate gyrus of CD/
mice treated with L-NAME at P24. Scale bar, 100 µm.
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Figure 6.
The number of TUNEL-positive cells in the thalamus
(TH) and the dentate gyrus
(DG) of the hippocampus from control littermates
(+/+), CD/ mice (/), CD/ mice treated with
L-NAME or SMT. The mean total number of TUNEL-positive
neurons in the thalamus of CD/ mice was significantly decreased by
chronic treatment of L-NAME or SMT without affecting that
in the subgranular zone of the dentate gyrus. The TUNEL-positive cells
were counted in semisequential sagittal sections of the thalamus and
subgranular zone of the dentate gyrus. Each column and
error bar represents the mean and SD, respectively.
**p < 0.01, ***p < 0.001, when
compared with the age-matched control littermates (Student's
t test).
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To assess the role of microglial NO in neuropathological changes in
CD/ mice, the competitive NOS inhibitor L-NAME (10 mg/kg, n = 6) or the iNOS inhibitor SMT (10 mg/kg,
n = 4) was administered intraperitoneally to CD/ mice at
P12 and twice a day thereafter for 13 consecutive days. The
chronic treatment of L-NAME or SMT partially but
significantly reduced the total number of TUNEL-positive cells in the
thalamus of CD/ mice (Figs. 5C, 6) without affecting the
total number of TUNEL-positive neurons in the dentate gyrus of the
hippocampus (Figs. 5D, 6). By contrast, chronic treatment of
L-NAME or SMT affected neither the number of
neurons laden with ceroid-lipofuscin nor the number of microglia that
phagocytosed these damaged storage neurons (control group,
24 ± 7 cells per square millimeter;
L-NAME-treated group, 20 ± 3 cells per
square millimeter). It was also noted that nuclei of damaged storage neurons that were phagocytosed by microglia were not stained with TUNEL
(data not shown).
Amelioration of the decrease in body weight and intestinal atrophy
in CE/ mice by chronic treatment of L-NAME or SMT
The decline of the body weight after P14 is one of the typical
pathological features of CD deficiency. In CD/ mice injected with
saline, body weight started to decline at approximately P14 as
described previously (Saftig et al., 1995) (Fig.
7). The mean weight of the CD/ mice
at P22 was 3.7 ± 0.9 gm (n = 8), which is only
50% of that of wild-type littermates (7.5 ± 0.2 gm;
n = 5). However, CD/ mice that were treated with
L-NAME or SMT did not show a significant decrease
in their body weight (Fig. 7). Although CD/ mice treated with
L-NAME or SMT showed a tendency for decline of
their body weight after P18, the difference of body weight between
these treated CD/ mice and their wild-type littermates did not
reach the statistical significance throughout the experimental
period. Furthermore, five of eight CD/ mice treated with
saline died by P23, whereas all CD/ mice treated with
L-NAME (n = 5) or SMT
(n = 5) were alive at this stage. In some experiments,
CD/ mice continued to be treated with L-NAME to determine whether L-NAME could significantly
prolong the survival of CD/ mice. The mean life span of
L-NAME-treated CD/ mice was 28.0 ± 0.8 d (n = 5), which is significantly longer than
that of nontreated CD/ mice (25.0 ± 0.6 d;
n = 8; p < 0.001; Student's t test). Thus L-NAME significantly
prolonged the survival of CD/ mice but could not abrogate the death
of CD/ mice.
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Figure 7.
Effects of L-NAME and SMT on body
weight of CD/ mice. The mean body weight of CD/ mice (/)
that were treated with saline (n = 8),
L-NAME (n = 5), and SMT
(n = 5), and control littermates (+/+)
(n = 5) was determined from P12 to P24.
L-NAME was systematically administered twice a day from P12
to P24. Each symbol and error bar represents the mean
and SD, respectively. ***p < 0.001, when compared
with the age-matched CD/ mice treated with saline (two-way ANOVA,
followed by Scheffe's post hoc test for multiple
comparison when F ratios reached significance).
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The small intestine of CD/ mice at P24 was found to exhibit
hemorrhage-necrotic appearance and atrophic changes of the ileal mucosa
as described previously (Saftig et al., 1995) (Fig.
8B,E), but no pathological changes were observed in these tissues of the
wild-type littermates (Fig. 8A,D).
Both the hemorrhage-necrotic appearance and the atrophic changes of the
ileal mucosa were markedly ameliorated by chronic treatment of
L-NAME (Fig. 8C,F)
or SMT (data not shown).
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Figure 8.
Effects of L-NAME on pathological
changes in the small intestine of CD/ mice.
A-C, The small intestine from control
littermate at P24 (A), CD/ mouse at P24
(B), and CD/ mouse at P24
(C) treated with L-NAME. The
treatment of L-NAME markedly ameliorated
hemorrhage-necrotic appearance of the small intestine from CD/
mice. D-F, Longitudinal sections through ileum from
control littermate at P24 (D), CD/ mouse at
P24 (E), and CD/ mouse at P24
(F) treated with L-NAME. The
treatment of L-NAME markedly ameliorated atrophic changes
of the ileal mucosa of CD/ mice. Scale bars: A, 5 mm; D, 100 µm.
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Accumulation of NO2 and expression
of iNOS in peripheral macrophages
We also measured the accumulation of
NO2 in the cytosolic
fraction of the small intestine of CD/ mice. At P16, the cytosolic fractions of the small intestine from the wild-type littermate mice
contained NO2 at a very
low concentration and its level was not significantly changed even at
P24. (Fig. 9A). On the other
hand, the mean concentration of
NO2 measured in the
cytosolic fractions of the small intestine from CD/ mice was
increased after P16; the
NO2 level was
approximately eightfold higher in CD/ mice at P24, compared with
P16. By Western blotting, a single band with a molecular mass of ~130
kDa representing iNOS was observed in the soluble fraction of the small
intestine of CD/ mice at P20, but not at P16 (Fig. 9B).
In corresponding samples from littermate mice at P16 and P24, iNOS was
not detectable. Indirect immunofluorescent staining with anti-iNOS
antibody revealed that iNOS was expressed in macrophage-like cells in
the small intestine of CD/ mice at P24 (Fig. 9C). No
staining was observed in the small intestine of control
littermates.
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Figure 9.
Expression of iNOS in the small intestine of
CD/ mice. A, Accumulation of
NO2 in the cytosolic fraction of the
small intestine of CD/ mice (/) and control littermates (+/+)
at P16 and P24. The number of experiments is shown in parentheses.
**p < 0.01, when compared with control littermates
(Student's t test). B,
Expression of iNOS proteins in the soluble fraction of the small
intestine of CD/ mice (/) and control littermates (+/+) at P16
and P20. Arrowhead indicates the iNOS protein with
molecular mass of 130 kDa. C, Immunostaining for iNOS in
the small intestine of CD/ mouse (/) and control littermate
(+/+) at P24. Scale bar, 50 µm.
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DISCUSSION |
In the CNS of CD/ mice after P20, we observed a marked
accumulation of morphologically transformed microglia exhibiting expanded and round cell bodies with few thick processes. They phagocytosed damaged storage neurons laden with ceroid-lipofuscin and
expressed iNOS. Furthermore, TUNEL-positive cells appeared especially
in the thalamus at the terminal stage of CD/ mice. NO and the super
oxide anion, which is generated in mitochondria, react rapidly to form
a peroxinitrite anion (Beckman et al., 1990). This, in turn, generates
highly toxic hydroxyl radicals and hydrogen peroxide. Although NO is
synthesized from L-arginine by NOS, iNOS is thought to be
the isoform that produces the large quantities of NO that can result in
tissue damage or death (Nathan and Xie, 1994). In the present study,
iNOS was found to be markedly expressed in activated microglia of
CD/ mice after P20. To directly address the possible involvement of
NO in tissue damage and cell death in the CNS of CD/ mice, we
examined effects of L-NAME, a potent competitive NOS
inhibitor, and SMT, an iNOS inhibitor. The chronic treatment of
L-NAME or SMT partially but significantly decreased the
number of TUNEL-positive cells in the thalamus at the terminal stage of
CD/ mice without affecting the number of TUNEL-positive cells in
the dentate gyrus of the hippocampus, in which apoptosis plays the
opposite role to neurogenesis in the regulation of cell numbers.
However, the accumulation of ceroid-lipofuscin in neurons and the
phagocytosis of these damaged storage neurons by microglia were not
changed by these treatments.
New Zealand sheep that express an inactive form of the point-mutated CD
molecule exhibit some neuropathological changes similar to CD/ mice
such as neuronal storage of autofluorescent granules and neuronal death
(Tyynelä et al., 2000). Thus, it appears that both neuronal
storage and neuronal death are directly attributable to the
defective activity of CD. The activation of microglia and subsequent
production of NO through iNOS are considered to be responsible for
neuronal death of CD/ mice, because microglial phagocytosis of
storage neurons and iNOS expression were found to precede an appearance
of TUNEL-positive cells in the thalamus. In both in
vitro and in vivo studies, NO has been
demonstrated to be a major causal factor for microglia-mediated
neuronal death (Boje and Arora, 1992; Chao et al., 1992; Meda et al.,
1995; Brown et al., 1996; Yan et al., 1996; Barger and Harmon, 1997;
Takeuchi et al., 1998; Matsuoka et al., 1999; Kim et al., 2000).
Takeuchi et al. (1998) reported that iNOS was induced in activated
microglia surrounding necrotic lesions induced by an injection of
ethanol. They also clearly indicated that iNOS can be induced in
microglia to produce NO sufficient to cause neuronal death on acute
brain injury without bacteria or molecules from their cell walls (lipopolysaccharides).
One possible mechanism underlying expression of iNOS high enough to
produce NO causing neuronal death is binding and/or phagocytosis of
damaged storage neurons by microglia. After engulfment of
apoptotic cells, peripheral macrophages have been reported to increase
the secretion of anti-inflammatory cytokines such as transforming growth factor- (TGF-) and decrease secretion of the
proinflammatory cytokines such as TNF-, interleukin
(IL)-1 , IL-8, and IL-12 (Voll et al., 1997; Fadok et al., 1998;
Freire-de-Lima et al., 2000). TGF- further depresses the
expression of iNOS and the production of NO by shifting the arginine
metabolic pathway to the ornithine synthetic one. On the
other hand, phagocytosis of opsonized apoptotic cells via the
complement receptor type 3 (CR3) has no significant effect on the
secretion of pro- or anti-inflammatory cytokines (Fadok et al., 1998).
However, CR3 contributes to the induction of iNOS and NO production in
peripheral macrophages (Matsumoto et al., 1998). Furthermore,
gangliosides that are particularly rich in neuronal cell membrane have
been reported recently to activate microglia to produce proinflammatory
mediators, including NO and TNF- (Pyo et al., 1999). Our
observations here showed that degenerating storage neurons in the
thalamus exhibited morphological features distinct from apoptosis at
both the light and electron microscopic levels. In our preliminary
experiments, TNF- was found to be increased in the thalamus of
CD/ mice after P20 (~45 pg/mg protein at P24). More recently,
Wada et al. (2000) have demonstrated that the inflammatory response
initiated by activated microglia play a pivotal role in neuronal
apoptosis of Sandhoff disease, an inherited glycolipid neuronal storage disease. Therefore, it is considered likely that microglia is activated
to produce NO through iNOS by binding and/or phagocytosis of damaged
storage neurons to initiate an intensive inflammatory response in the
CNS leading to secondary neuronal damage evidenced by an appearance of
TUNEL-positive cells. Although presumptive neuronal apoptosis has been
investigated by TUNEL staining, it is now recognized that this assay
can no longer define cell death as apoptosis (Grasl-Kraupp et al.,
1995; Wang et al., 1999). We therefore referred to an appearance of
TUNEL-positive neurons as neuronal damage rather than neuronal
apoptosis. In summary, CD deficiency causes lysosomal
storage in neurons, which stimulate and become internalized by
microglia. Microglial NO induces neuronal damage in adjacent neurons
and an inflammatory response in the CNS. The neuronal damage induced by
NO can effectively be prevented by treatment with
L-NAME or SMT.
In the course of our experiments, we have unexpectedly found that the
chronic treatment of L-NAME or SMT almost completely suppressed the decrease in the body weight of CD/ mice after P14.
The closer histochemical analysis here revealed that this treatment
markedly ameliorated a severe hemorrhage-necrotic appearance of the
small intestine and atrophic changes of the ileal mucosa of CD/
mice. Furthermore, the accumulation of
NO2 and expression of
iNOS were also found in the small intestine of CD/ mice. These
observations are consistent with the notion that NO has potential
cytotoxic effects on the intestine (Boughton-Smith et al., 1993;
Laszlo et al., 1994). Furthermore, L-NAME
significantly prolonged the survival of CD/ mice but could not
abrogate the death of CD/ mice. We speculate that most of
L-NAME-treated CD/ mice died from seizure because they
still showed signs of seizure without severe loss of body weight and
intestinal necrosis. On the basis of these observations, NO production
through iNOS expressed in peripheral macrophages may lead to a
breakdown of mucosa in the small intestine of CD/ mice. Although
the mucosal damage is one of the most typical pathological features of
CD/ mice, no such pathological change in the small intestine was
observed in White Swedish Landrace sheep (Tyynelä et al., 2000).
Thus, the expression of iNOS and resultant mucosal damage is not
directly attributable to the defective activity of CD. Further studies will be needed to clarify the factor responsible for the expression of
iNOS in the intestine of CD/ mice.
| |
FOOTNOTES |
Received Feb. 9, 2001; revised July 9, 2001; accepted July 9, 2001.
This work was supported by Grants-in-Aid on Priority Area from the
Ministry of Education, Science, Sports and Culture, Japan.
Correspondence should be addressed to either Dr. Hiroshi Nakanishi,
Laboratory of Oral Aging Science, Division of Oral Biological Sciences,
Faculty of Dental Sciences, Kyushu University, Fukuoka 812-8582, Japan,
E-mail: nakandeg{at}mbox.nc.kyushu-u.ac.jp., or Dr. Yasuo Uchiyama,
Department of Cell Biology and Neuroscience, Osaka University Graduate
School of Medicine, Suita, Osaka 565-0871, Japan, E-mail:
uchiyama{at}anatl.med.osaka-u.ac.jp.
| |
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ABSTRACT
INTRODUCTION
