J. Neurochem. (2011) 116, 374–384.
Stomatin is an important membrane raft protein which can combine skeleton protein, some ion channel, and transporter to regulate their functions. However, until now no data on its expression and function in CNS are available. In this study, we examined distribution of stomatin in CNS of rat, and investigated the effects of hypoxia exposure and glucocorticoid on stomatin expression in cerebral cortex of rat. Immunofluorescence staining revealed a broad expression of stomatin protein in many areas of adult rat brain and spinal cord, including the ventral horn of spinal cord, causal magnocellular nucleus of hypothalamus, the V layer of the cerebral cortex, solitary nucleus, 10 and 12 nuclei, and so on. Hypoxia or ischemic hypoxia significantly up-regulated stomatin expression in cerebral cortex, and the up-regulation was independent on adrenocortical steroids since it also occurred in adrenalectomized (ADX) rats. Moreover, treatment of ADX or sham-operated rats with dexamethasone, a synthetic glucocorticoid alone could significantly stimulate expression of stomatin in lung and heart, but not in cerebral cortex. However, dexamethasone could enhance the hypoxia-stimulated expression of stomatin in cerebral cortex of ADX rats. These findings suggested that stomatin might be involved in various physiological functions and cellular events of neurons in CNS under physiological conditions and play a potential protective role under hypoxic conditions.
acid-sensing ion channels
degenerin/epithelial sodium channel
hypoxia inducible factor 1α
middle cerebral artery
Stomatin, also known as band 7.2 b was first identified in human erythrocytes and the absence of the protein is associated with a form of hemolytic anemia known as hereditary stomatocytosis (Stewart et al. 1992; Gallagher and Forget 1995). Stomatin is widely expressed in many cell types, and localized in detergent-resistant membrane domains enriched in cholesterol and sphingolipids, therefore it is involved in membrane organization and cholesterol-dependent regulatory processes (Salzer and Prohaska 2001; Umlauf et al. 2004, 2006; Kadurin et al. 2009). As a lipid raft protein for membrane microdomains, it is believed to combine various membrane proteins, such as ion channel, skeleton protein, transporter, and signal transducer to regulate their biological activities although the mechanism is unknown. For example, studies indicate that stomatin co-localizes with actin microfilaments in human amniotic epithelial cell line (Snyers et al. 1997) and interacts with actin in red blood cells, which is important in maintaining the structure and in modulating the function of stomatin-containing membrane rafts (Wilkinson et al. 2008). A stomatin-specific, raft-based process is also involved in storage-associated vesiculation in red cells (Salzer et al. 2008). In addition, stomatin has also been shown to bind to the glucose transporter to decrease the rate of glucose uptake (Zhang et al. 2001).
Recent studies report that stomatin can regulate the function of degenerin/epithelial sodium channels (DEG/ENaC) that are voltage-insensitive, amiloride-blockable cation channels (Fricke et al. 2000; Kellenberger and Schild 2002). Mechanosensory protein 2, a highly homologous protein to stomatin in Caenorhabditis elegans is found to interact with the DEG/ENaC, and the interaction of these two proteins is necessary for normal mechanosensation in the worm (Goodman et al. 2002; Huber et al. 2006). Several acid-sensing ion channels (ASICs) that are H+-gated members of the DEG/ENaC family are expressed in sensory neurons in vertebrate, where they play a role in response to nociceptive, taste, and mechanical stimuli. Recent study shows that stomatin co-localizes with ASIC proteins and modulates the physiological properties of ASICs. For example, stomatin potently reduced acid-evoked currents generated by ASIC3 and accelerated the desensitization rate of ASIC2 (Price et al. 2004). Moreover, stomatin is known to express in almost all sensory neurons of the dorsal root ganglia in mice, and lacking stomatin in mice leads to a specific loss of mechanoreceptor function in vivo (Martinez-Salgado et al. 2007; Wetzel et al. 2007). These observations indicate that stomatin has important implications for mammalian mechanosensation and nociception. Since DEG/ENaCs and some of ASICs, such as ASIC1 are also expressed in CNS (Alvarez de la Rosa et al. 2003; Amin et al. 2005), where they participate in synaptic plasticity and some forms of learning, it is interesting to ask if stomatin acts in CNS by interacting with ion channels or other membrane proteins.
Cerebral hypoxia can be caused by low oxygen in the environment, whereas the presence of areas of hypoxia in brain is a prominent feature of cerebral infarction, inflammation, and tumor in brain. Glucocorticoid (GC) plays an essential role in stress response and homeostatic regulation, especially in adaptation to hypoxic condition. In addition, GC has been widely used to treat hypoxia-induced tissue edema and injury in clinical. Previous study showed that dexamethasone (Dex), a synthetic GC and interleukin-6 up-regulated the expression of stomatin in human amniotic epithelial cells (Snyers and Content 1994). Our recent study also found that Dex up-regulated the expression of stomatin in human A549 cells, an alveolar type II cell line, and lung of rat. These findings strongly suggested that stomatin may play a role in stress response or under other pathophysiological conditions. Since to date no data on the distribution and function of stomatin in CNS are available, the purpose of this study is to determine the distribution of stomatin in rat CNS and investigate the effects of hypoxia and GC treatment on expression of stomatin.
Animals and hypoxia exposure
This study was approved by the Institutional Ethics Committee (protocol No. M2008-004/20080123), following the guidelines of the National Institutes of Health for the care and use of laboratory animals. Adult male Sprague-Dawley rats (250–300g; Shanghai SLAC laboratory animal company, China) were housed in Plexiglas cages (60 × 30 × 24cm), with food and water ad libitum. The animals were maintained in a pathogen-free barrier facility with a 12 : 12-h light–dark cycle. Rats were given ≥ 2 weeks to habituate to the environment before experimentation. Rats randomly selected were put in a normobaric hypoxia cabin (40 L; Yangyuan hyperbaric chamber company, Shanghai, China) flushed with a gas mixture of 8% O2 and 92% N2 (Vivona et al. 2001) for 8, 12, and 16 h. After hypoxia exposure termination, animals were killed by pentobarbital sodium anesthesia and decapitation. Brains were dissected and the prefrontal cortex was separated from subcortical areas of the brain for stomatin detection.
Four normal control adult rats (250–300 g), five hypoxia exposed rats and five rats (250–300 g) with middle cerebral artery (MCA) occlusion were anesthetized with sodium pentobarbitone and perfused through the aorta with 0.9% NaCl solution and 4% paraformaldehyde in 0.1 m phosphate-buffered saline (PBS, pH 7.2). The brains and spinal cords were dissected out immediately and immersed in 4% paraformaldehyde in 0.1 m PBS (pH 7.2) for 2–4 h. The tissues were then transferred to 25% sucrose in PBS and kept in the solution until they sank to the bottom. Thereafter, the tissues were rapidly frozen by immersion in isopentane at −70°C for 2 min. Coronal sections (20 μm) of the brains and spinal cord were cut with a Leica cryostat (CM1900, Leica, Nussloch, Germany) and floated in PBS. The sections were washed 3 × 5 min in PBS, and then pre-incubated in antiserum solution (10% normal bovine serum, 0.2% Triton-X-100, 0.4% sodium azide in 0.01 m PBS pH7.2) for 30 min, followed by the incubation with stomatin antibody (goat-anti-rat; Santa Cruz Biotechnology, Santa Cruz, CA, USA) diluted 1 : 50 in antiserum solution (1% normal bovine serum, 0.2% Triton-X-100, 0.4% sodium azide in 0.01 m PBS pH7.2) at 25°C overnight. Subsequently, the sections were incubated with Cy3-conjugated donkey anti-Goat IgG (Jackson Immuno-Research Laboratory, West Grove, PA, USA) diluted 1 : 400. All the incubations were separated by 3 × 10 min washes in PBS. A negative control of omitting the primary antibody was carried out. No staining was observed in those sections. The density of stomatin immunostaining signal was scored as absent (−), weak (+), moderate (++), heavy (+++) (Collo et al. 1996; Guo et al. 2008). Images were taken with the Nikon digital camera DXM1200 (Nikon, Chiyoda-ku, Japan) attached to a Nikon Eclipse E600 microscope (Nikon).
The sections from the frontal cerebral cortex of control and hypoxia rats were stained with 0.5% cresyl violet (Sigma-Aldrich, St Louis, MO, USA), dehydrated, coverslipped, and then analyzed under a bright-field microscope (Nikon).
Determination of the water content of brain
The water content of brain tissue was detected using the dry–wet weight technique. Five adult male Sprague-Dawley rats were put in normal pressure hypoxia chamber filled with 8% O2 and 92% N2 for 12 h. At the end of the experiment, the animals were anesthetized and killed, and the brains were harvested quickly. Brain wet weight was determined, then put the brain into 56°C baker for 72 h and collected the dry tissue to weight. The water content was calculated with the following formula:
Sprague-Dawley rats were anesthetized with 15% chloral hydrate (300 mg/kg, intraperitoneally). Permanent focal cerebral ischemia was induced by electrocoagulation of the distal portion of the MCA using a modified method described by Tamura and McGill (Tamura et al. 1981; McGill et al. 2005). Briefly, a segment of right MCA between the olfactory bundle and the inferior cerebral vein was electro-coagulated. The coagulated artery was severed with microscissors to ensure complete stop of blood supply. Brain samples were harvested 24 h after MCA occlusion. Coronal sections of 2 mm thickness were immediately stained with 2% 2, 3, 5-triphenyltetrazolium chloride as previously described (Bederson et al. 1986). Moreover, total RNA in cerebral cortex around the ischemia area or contralateral control area was extracted for stomatin detection.
Adrenalectomy and Dex supplementation
Animals were anesthetized with 15% chloral hydrate (300 mg/kg, intraperitoneally), and adrenal glands were removed via the dorsal approach, as previously described (Fleshner et al. 2001). Sham animals underwent the same surgery, except the adrenal glands were left intact. Adrenalectomized (ADX) rats were given 0.9% saline ad libitum to compensate for sodium loss after the operation and allowed to recover for 5 days before Dex supplementation or/and hypoxia exposure.
The ADX rats were injected intramuscularly with 5 or 10 mg/kg body weight of Dex (Sigma-Aldrich) dissolved in a 0.9% NaCl solution. Control ADX rats were treated with 0.9% NaCl alone (Folkesson et al. 2000). At the same time of Dex supplementation, hypoxia exposure was done in the ADX rats of combined treatment group. Rats were anesthetized and killed at 12 or 24 h after treatments, and lung, heart, and cerebral cortex were isolated for RNA extraction.
Total RNA was extracted with TRIzol reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s protocol. A total of 2 μg RNA was subjected to synthesize first-strand cDNA by reverse Transcription System (Promega, Madison, WI, USA) using random primers. The resulting cDNA was employed for real-time PCR using Power SYBR Green PCR Master Mix (Toyobo, Osaka, Japan) on Mastercycler ep realplex (Eppendorf, Hamburg, Germany). The sequences of the forward and reverse primers of stomatin were as follows: 5′-TGGTGGCTGTCTCGTTC-3′ and 5′-ATTCCTGAAGGTAGTTTGC-3′, yielding a 361 bp product. Beta-actin was used as an internal control; the sequences of the forward and reverse primers were as follows: 5′-ATGGTGGGTATGGGTCAGAAG-3′ and 5′-TGGCTGGGGTGTTGAAGGTC-3′, yielding a 265 bp product. Thermal cycling conditions consisted of an initial denaturing step (95°C, 2 min) followed by 40 cycles of denaturing (94°C, 30 s), annealing (60°C, 30 s), and extending (72°C, 30 s). The specified mode of reaction was controlled with the melting curve. Amplification of stomatin cDNA was normalized to that of β-actin. Results are expressed as a fold-increase of stomatin cDNA compared with that of control.
Western blot analysis
Protein extracts for western blot analysis were prepared by washing the brains three times with PBS, dissecting the cerebral cortex, and homogenizing the cerebral cortex using a homogenizer with ice-cold radio immunoprecipitation assay lysis buffer (contain 1 mmo-phenylmethylsulfonyl fluoride; Shenergy Bilcolor BioScicnce & Technology Company, Shanghai, China). Protein concentrations in the extracts were measured using the bicinchoninic acid Protein Assay Kit (Bio-Rad Laboratories, Hercules, CA, USA). The same amount of protein for each sample was electrophoresed on 10% sodium dodecyl sulfate–polyacrylamide gel, and then transferred to a nitrocellulose membrane. The membrane was blocked with 5% non-fat milk in tris-buffered saline Tween-20 (25 mm Tris-HCl, pH 8.0; 144 mm NaCl; 0.1% Tween-20), and incubated with primary antibodies against stomatin (sc-48309 1 : 500; Santa Cruz), hypoxia inducible factor 1α (HIF1α) (1 : 500; Santa Cruz), or β-actin (1 : 5000; Sigma-Aldrich) overnight at 4°C followed by the secondary antibody, horseradish peroxidase-conjugated rabbit anti-goat IgG antibody (Santa Cruz), or rabbit anti-mouse IgG antibody (HuaMei Biotechnology company, Shanghai, China). Immunodetected proteins were visualized using enhanced chemiluminescence assay kit (Shanghai Shenergy Biocolor Bioscience & Technology Company) following the manufacturer’s recommended protocol.
Statistical analysis was performed using one-way anova followed by Fisher’s least significant difference test, and p < 0.05 was considered significant. Data are expressed as the means ± SD.
Distribution of stomatin expressing neurons in the CNS
Stomatin immunoreactivity was to be in a wide-ranging expression pattern throughout the brain and spinal cord. Stomatin immunoreactivity was dense in many regions, such as ventral horn of spinal cord, causal magnocellular nucleus of hypothalamus, the V layer of the cerebral cortex, solitary nucleus, and 10 and 12 nuclei. The distribution pattern and immunostaining density of stomatin protein were summarized in Table 1.
|Olfactory bulb||Anteroventral nucleus||+|
|Mitral cells||++||Laterodorsal nucleus||+|
|Granular layer||+−||Mediadorsal nucleus||+|
|Tufted cells||+−||Ventrolateral nucleus||+|
|Anterior olfactory nucleus||++||Ventromedial nucleus||+|
|Cerebral cortex||Reticular nucleus||+|
|Agranular insular cortex||++||Paraventricular nucleus||+|
|Anterior cingulate cortex||++||Central medial nucleus||−|
|Ventrolateral orbital cortex||++||Xiphoid nucleus||−|
|Piriform cortex||++||Rhomboid nucleus||+|
|Parietal cortex||++||Suprachiasmatic nucleus||+|
|Occipital cortex||++||Medial preoptic nucleus||+|
|Subcortical telencephalon||Medial preoptic area||+|
|Nucleus diagonal band||++||Supraoptic nucleus||++|
|Bed nucleus stria terminalis||+||Paraventricular nucleus||++|
|Medial septa nucleus||+||Ventromedial nucleus||+|
|Lateral septal nucleus||+||Arcuate nucleus||+|
|Islands of Calleja||−||Tuberomammillary nucleus||+|
|Olfactory tubercle||+||Causal magnocellular nucleus||+++|
|Caudate Putamen||+||Zona incerta||+|
|Globus pallidurn||+||Lateral geniculate nucleus||+|
|Substantia innommata||+||Medial geniculate nucleus||+|
|Medial globus pallidurn||+||Substantia nigra zona compacta||++|
|Lateral olfactory tract nucleus||+||Substantia nigra zona reticulate||+|
|Anterior cortical amygdaloid nucleus||+||Supramammillary nucleus||+|
|Basolateral amygdaloid nucleus||++||Ventral tegmental area||+|
|Central amygdaloid nucleus||++||Oculomotor nucleus||+|
|Medial amygdaloid nucleus||++||Edinger-Westphal nucleus||+|
|Amygdalohippocampal area||++||Central gray||+|
|Hippocampal fields CAl–CA4||++||Superior colliculus||+|
|Dentate gyrus||+||Inferior colliculus||+|
|Medial habenular nucleus||++||Trochlear nucleus||++|
|Lateral habenular nucleusa||+||Pedunculopontine tegmental nucleus||+|
|Anterior nuclei||+++||Pontine nuclei||+|
|Raphe dorsalis||+||Dorsal motor vagus nucleus||+++|
|Mesencephalic trigeminal nucleus||++||Solitary tract nucleus||+++|
|Dorsomedial tegmental area||+||Lateral reticular nucleus||++|
|Locus ceruleus||+||Curate fasciculus||++|
|Subceruleus nucleus||+||Area postrema||+|
|Pontine reticular nucleus, ventral part||+||Purkinje cells||++|
|Pontine reticular nucleus, caudal part||+||Granular layer||−|
|Red nucleus||++||Cerebellar nucleus||++|
|Motor trigeminal nucleus||++||Spinal cord|
|Superior olivary nucleus||+||Lamina I||+|
|Principal sensory trigeminal nucleus||++||Lamina II||+|
|Spinal trigeminal nucleus||++||Lamina III||++|
|Paragigantocellular nucleus||+||Lamina IV||+|
|Abducens nucleus||+||Lamina V||++|
|Facial nucleus||+++||Lamina VI||++|
|Accessmy facial nucleus||++||Lamina VII||++|
|Ambiguous nucleus||++||Lamina VIII||+++|
|Inferior olive||++||Lamina IX||+++|
|Hypoglossal nucleus||+++||Lamina X||++|
Main olfactory bulb, accessory olfactory bulb, and primary olfactory cortex showed stomatin immunoreactivity at moderate levels. Some of olfactory nerve branches and lateral olfactory tract were heavily immunostained. Moderate stomatin immunoreactivity was detected in the bodies of mitrial cells and piriform cortex (Fig. 1a). Moderate to heavy stomatin immunoreactivity was observed in all cerebral cortexes. Typically, stomatin immunostaining was mainly localized in II, III, IV, and V layers of cerebral cortex (Fig. 2e and f). Pyramidal cells of layer V showed the denser labeling than those of other layers. Moderate to heavy immunostaining nerve tracts were observed in the subcortex regions, such as corpus callosum, septal nucleus, caudate putamen, external capsule, fonix, and fimbria of the hippocampus. Moderate immunostaining neurons were observed in the vertical and horizontal parts of diagonal band, entopeduncular nucleus, substantia innommata. Weak to moderate immunostaining neurons were demonstrated in all the regions of amydaloid complex (Fig. 1b and c). Pyramidal cells in the cornu ammonis 1–3 of hippocampal fields showed moderate immunostaining (Fig. 1d and e). Granule cells of dentate gyrus showed weak immunostaining. In the thalamus, heavy immunostaining was showed in anterodorsal thalamic nucleus, but most of other thalamic nuclei showed weak immunostaining (Fig. 1f). Heavy immunostaining in optic chiasm and causal magnocellular nucleus, moderate immunostaining in paraventricular and supraoptic nuclei was detected in the hypothalamus (Fig. 1g). In the median eminence varicose axonal-like fibers for stomatin immunoreactivity were detected. Weak immunostaining was showed in other regions of the hypothalamus. In the midbrain neurons of substantia nigra, compact part showed moderate stomatin immunostaining (Fig. 1h). The medial geniculate nucleus, ventral tegmental area, oculomotor nucleus, etc. expressed weak stomatin immunoreactivity, although scattered neurons were immunostained moderately for stomatin. Moderate immunostaining in Purkinje cells and interposed cerebellar nucleus and weak immunostaining in the axonal-like fibers was detected in white matter of cerebellum (Fig. 1i). Stomatin immunostaining distributed widely in hindbrain. Heavy immunostaining was detected in motor trigeminal nucleus, hypoglossal nucleus, dorsal motor vagus nucleus, etc. (Fig. 1j and k). Weak to moderate immunostaining was detected in the pontine reticular nucleus, red nucleus, etc. Stomatin immunostaining was observed in the spinal gray matter at cervical, thoracic, lumbar, and sacral levels. The most prominent staining was seen in the ventral horn. Heavy immunostaining was detected in motoneurons. Weak to moderate stomatin immunostaining was also detected in other regions of the gray matter (Fig. 1l).
Hypoxia up-regulates the expression of stomatin in cerebral cortex of rat
To test the effect of hypoxia exposure (8% oxygen), we first examined the level of HIF1α by western blot and the water content of brain of rats, and confirmed that the protein level of HIF1α in cerebral cortex was significantly increased in a time of hypoxia exposure dependent fashion (Fig. 2a). The water content of brain also found to be significantly increased (from 77.74% to 78.23%, p < 0.05; Fig. 2b) determined by brain/body weight ratio at 12 h after hypoxia exposure, indicating that hypoxia caused brain edema.
Then, we examined stomatin mRNA and protein levels in cerebral cortex of rat after exposing to hypoxic condition for different time by real-time PCR and western blot analysis. As shown in Fig. 2(c), hypoxia exposure for 8 h significantly up-regulated stomatin mRNA, reaching to twofold of that in normoxia control at 12 h (p < 0.05). The up-regulation of stomatin mRNA was accompanied by a significant increase in stomatin protein level in the cerebral cortex in this model (Fig. 2d).
The results of immunostaining for stomatin were similar with that of western blot. As shown in Fig. 2(e–h), hypoxia exposure also up-regulated the stomatin immunoreactivity, especially in the II and III layers of the cerebral cortex (Fig. 2g) as compared with the control (Fig. 2e). The number and morphology of the cerebral cortex neurons stained by Nissl staining in the control and hypoxia groups were quite similar. No significant changes were observed (Fig. 2i, j, i′ and j′).
Ischemic hypoxia up-regulates the expression of stomatin in cerebral cortex of rat
We further examined expression of stomatin in hypoxia-ischemic model induced by electrocoagulation of the distal portion of the right MCA of the rat. As shown in Fig. 3(a), the infarct region looked pale, whereas the normal region looked red. Compared with the same area in cerebral cortex of the left site, mRNA of stomatin in the cerebral cortex around the ischemia area was decreased at 12 h, and then significantly increased at 24 and 36 h (about 2.1-fold) after MCA occlusion (p < 0.05; Fig. 3b). The increased stomatin immunoreactivity was also observed in the adjacent region of the ischemical region (Fig. 3e and f) as compared with the left site (Fig. 3c and d), whereas the stomatin immunoreactivity was almost disappeared in the ischemical regions (Fig. 3e).
Effects of Dex on expression of stomatin mRNA in lung, heart, and cerebral cortex of rat
It is well known that hypoxia exposure results in high levels of circulating GCs in vivo. In this study, we found that Dex up-regulated expression of stomatin in A549 cells in a time- and dose-dependent fashion. To test the effect of adrenocortical steroids, especially GC on the expression of stomatin in vivo, we examined the mRNA levels of stomatin in cerebral cortex, lung, and heart of rat underwent sham surgery or adrenalectomy with or without supplementation of Dex, a synthetic GC which has a high affinity for glucocorticoid receptor (GR). The results showed that as compared with that in sham group, adrenalectomy resulted in significant decrease of stomatin mRNA in cerebral cortex, lung, and heart. Treatment of ADX rats with Dex (5 mg/kg) for 12 h significantly increased the levels of stomatin mRNA of lung and heart (Fig. 4a and b). However, same dose of Dex could not increase the mRNA of stomatin in cerebral cortex, even extend time of treatment to 24 h (Fig. 4c). Furthermore, supplementation of normal and ADX rats with higher concentration of Dex (10 mg/kg for 12 h) still did not significantly change the expression level of stomatin(Fig. 4d). These results indicate that adrenocortical steroids can up-regulate expression of stomatin in vivo in a tissue-specific manner.
Effects of hypoxia and Dex alone or combined treatment on expression of stomatin in sham or ADX rat
To test the influence of adrenocortical steroids, especially GC on expression of stomatin in hypoxic brain, we examined the mRNA and protein levels of stomatin in hypoxic cerebral cortex of rat underwent sham surgery or adrenalectomy with or without supplementation of Dex. As shown in Fig. 5(a), hypoxia significantly increased mRNA level of stomatin both in sham rats (twofold of that in sham control group, p < 0.05) and in ADX rats (1.8-fold of that in the ADX control group, p < 0.05). Although treatment of ADX rats with Dex (5 mg/kg) for 12 h did not significantly alter the expression of stomatin, higher mRNA level of stomatin in ADX rats treated with both hypoxia and Dex were observed (2.6-fold of that in the ADX group, p < 0.01). Similar results were also observed in protein level of stomatin determined by western blot (Fig. 5b). These results indicated that hypoxia up-regulated the expression of stomatin was independent of hypothalamic–pituitary–adrenal axis (HPA) activation in vivo. Dex and hypoxia could co-induce the expression of stomatin in cerebral cortex of rat.
This is the first extensive study undertaken to investigate the distribution of stomatin protein in rat CNS using immunofluorescence staining method. Our data provide clear evidence that stomatin expression is not limited to sensory neurons. In contrast, it is widely distributed in the CNS. Strong immunoreactivity for stomatin protein was showed in some regions relating to motor system, such as the motor neurons of spinal cord, facial nucleus, motor trigeminal nucleus, hypoglossal nucleus, and dorsal motor vagus nucleus. Lots of varicose fibers immunostained by stomatin antibody in the median eminence and moderate to heavy immunoreactivity for stomatin protein were found in the supraoptic, paraventricular, causal magnocellular nucleus relating to the neuroendocrine system. It was interesting to find that stomatin was co-localized with DEG/ENaC and ASIC in the CNS. For example, all three DEG/ENaC subunits were present in the supraoptic nucleus, magnocellular paraventricular nucleus, hippocampus, most cerebral cortical areas, thalamus, solitary nucleus, and amygdala (Amin et al. 2005), whereas some ASICs, such as ASIC1 was found to distribute widely in the CNS of adult rat including the cerebral cortex, hippocampus, striatum, thalamus, midbrain, pons, spinal cord, and cerebellum (Alvarez de la Rosa et al. 2003). Although it is not known whether these channels are interacted with stomatin or not, the wide distribution of stomatin in CNS at least suggests that stomatin may be involved in various physiological functions and fundamental cellular events of neurons in CNS, especially may play an important role in the motor system and neuroendocrine system.
Next, we examined the effect of hypoxia or hypoxia-ischemia induced by electrocoagulation of the distal portion of the right MCA on expression of stomatin in cerebral cortex of rat, and found a significant up-regulation of stomatin expression in both kinds of animal models. It is well known that hypoxia exposure results in the activation of HPA and increased levels of circulating adrenocortical steroids, especially GCs in vivo. To rule out the possible effect of adrenocortical steroids on expression of stomatin in hypoxic brain, we examined the mRNA and protein levels of stomatin in hypoxic cerebral cortex of rat underwent sham surgery or adrenalectomy. We found that hypoxia also increased the mRNA and protein levels of stomain even in ADX rat, indicating that the effect of hypoxia on stomatin expression is independent of HPA activation. However, it has not been known the significance of increased expression of stomatin in brain under hypoxic condition. It has been reported that stomatin co-localizes with ASIC proteins and modulates the physiological properties of ASIC in sensory neurons in vertebrate (Price et al. 2004; Martinez-Salgado et al. 2007; Wetzel et al. 2007). Moreover, we also know that ASIC is activated by low extracellular pH caused by hypoxia, ischemia, or inflammation. Therefore, we speculate that the increased stomatin may interact with some members of ASIC or other membrane components, such as actin or other ion channels in CNS to keep molecular and functional homeostasis under hypoxic condition in vivo.
Although the effect of hypoxia on expression of stomatin is independent of HPA activation, our study shows that adrenocortical steroids are indispensable for maintaining physiological level of stomatin in vivo since adrenalectomy resulted in significant decrease of stomatin expression in cerebral cortex, lung, and heart as compared with sham group. It is well known that endogenous GC in rat is mainly corticosterone which binds mineralocorticoid receptors with an affinity 10-fold higher than co-localized GR in brain (De Kloet et al. 1998), therefore, we used Dex, a synthetic GC that had a high affinity for the GR to examine the effect of GC on the expression of stomatin in cerebral cortex of rat. We found that supplementation of ADX rat with Dex obviously up-regulated the expression of stomatin in lung and heart, but not in cerebral cortex, even at a high dose under normoxic condition. Our group previously examined the effect of GC on binding capacity of GR determined by the specific binding of [3H] Dex, and found similar results that maintaining the concentration of hydrocortisone at stress level in plasma (20–40 μg/dL) by subcutaneous injection of polyvinyl alcohol containing hydrocortisone into rats for 3 days led to significant decrease of the binding capacity of GR in cytosol of liver and spleen cells of rat, but not in cytosol of brain hemispheres (Yang et al. 1989; Xu and Tan 1990). Why Dex displays different effects in peripheral tissues and brain remains unclear. One possible reason is that, according to the literatures, Dex preferentially targets the pituitary in the blockade of stress-induced HPA activation, and administration of moderate amounts of Dex partially depletes the corticosterone in brain (De Kloet et al. 1998). Another explanation of this phenomenon may be owing to a poor penetration of Dex or cortisol into rat brain caused by the multidrug-resistance 1a -encoded p-glycoproteins in the blood–brain barrier (Bourgeois et al. 1993; Schinkel et al. 1995; De Kloet 1997; Meijer et al. 1998; Karssen et al. 2001). Although Dex alone could not up-regulate the expression of stomatin in cerebral cortex under normoxic condition, Dex was found to enhance the expression of stomatin under hypoxic condition, suggesting that there is cross-talk between hypoxia-dependent signal pathways and GC/GR-mediated signal pathways. Therefore, it will be interesting to investigate the interaction between these important stress-responsive pathways, focusing on the regulation of stomatin expression in future.
We summarize the findings as follows: (i) stomatin was widely expressed in CNS, and regional distribution was uneven, with predominant expression in motor neurons; (ii) hypoxia significantly up-regulated the expression of stomatin in cerebral cortex, which was independent on adrenocortical steroids; (iii) Dex did not induce expression of stomatin alone, but enhanced stomatin expression induced by hypoxia in cerebral cortex of rat. These findings add to the understandings of distribution and the regulation of stomatin in CNS, and suggest a potential protective role for increased stomatin in brain under hypoxic condition in vivo.
We would like to thank Prof. Xue-jun Sun (Department of Diving Medicine, Faculty of Naval Medicine, Second Militory Medical University) for provided us hypoxia chamber for rats and Dr. Liangnian Song (Herbert Irving Comprehensive Cancer Center Columbia University, USA) for critical reading of the article. This work was supported by grant from The National Basic Research Program ‘973’ No. 2006CB504100.