Abstract

Mutations in the MLC1 gene, which encodes a protein expressed in brain astrocytes, are the leading cause of MLC, a rare leukodystrophy characterized by macrocephaly, brain edema, subcortical cysts, myelin and astrocyte vacuolation. Although recent studies indicate that MLC1 protein is implicated in the regulation of cell volume changes, the exact role of MLC1 in brain physiology and in the pathogenesis of MLC disease remains to be clarified. In preliminary experiments, we observed that MLC1 was poorly expressed in highly proliferating astrocytoma cells when compared with primary astrocytes, and that modulation of MLC1 expression influenced astrocyte growth. Because volume changes are key events in cell proliferation and during brain development MLC1 expression is inversely correlated to astrocyte progenitor proliferation levels, we investigated the possible role for MLC1 in the control of astrocyte proliferation. We found that overexpression of wild type but not mutant MLC1 in human astrocytoma cells hampered cell growth by favoring epidermal growth factor receptor (EGFR) degradation and by inhibiting EGF-induced Ca+ entry, ERK1/2 and PLCγ1 activation, and calcium-activated KCa3.1 potassium channel function, all molecular pathways involved in astrocyte proliferation stimulation. Interestingly, MLC1 did not influence AKT, an EGFR-stimulated kinase involved in cell survival. Moreover, EGFR expression was higher in macrophages derived from MLC patients than from healthy individuals. Since reactive astrocytes proliferate and re-express EGFR in response to different pathological stimuli, the present findings provide new information on MLC pathogenesis and unravel an important role for MLC1 in other brain pathological conditions where astrocyte activation occurs.

Introduction

Astrocytes have an essential role in water and ion homeostasis, energy metabolism, control of neurogenesis, myelination and synaptic transmission in the central nervous system (CNS). Consistent with this multiplicity of physiological functions, astrocytes are involved in a wide spectrum of CNS pathologies where loss of astrocyte homeostatic functions and/or gain of toxic functions may occur as a consequence of specific pathogenic insults (1). In addition, recent studies have shown that mutations of genes encoding astrocyte-specific proteins such as GFAP and MLC1 are the direct cause of rare genetic leukodystrophies like Alexander's disease and megalencephalic leukoencephalopathy with subcortical cysts (MLC) (2–4). MLC disease is clinically characterized by macrocephaly, deterioration of motor functions with ataxia and spasticity, epileptic seizures and mental decline that often aggravate after minor head trauma or common infections (5–9). Brain magnetic resonance imaging revealed the presence of subcortical cysts and diffuse white matter edema, while analysis of rare brain biopsies showed myelin vacuolization (10,11) and reactive astrocytes with swollen end-feet (12). More than 60 mutations (including missense, splice site, insertions and deletions) in the MLC1 gene have been associated with MLC disease (13–15).

To date, MLC1 protein function is still unknown. Amino acid sequence examination suggests that MLC1 is a highly hydrophobic protein with eight predicted transmembrane domains, short amino- and carboxylic cytoplasmic domains (16–20), and several putative phosphorylation sites (19,20). In the human and mouse CNS, MLC1 is expressed almost exclusively in astrocytes and also in Bergmann glia of the cerebellum, but not in neurons and oligodendrocytes, the myelin forming cells (16–18,21–23), suggesting that myelin degeneration in MLC disease is the consequence of a primary astrocyte defect. In vitro and in vivo studies indicate that MLC1 might be involved in the regulation of astrocyte volume changes by favoring the activation of calcium and chloride currents and regulatory volume decrease (RVD) following astrocyte swelling in response to ionic imbalance (23–28). Besides being hallmarks of many pathological conditions, cell volume changes also accompany cell proliferation, differentiation and death (29). Previous studies revealed that inhibition of cell proliferation favors MLC1 protein expression in primary rat astrocytes (12). In mouse and human brain, MLC1 gene expression is developmentally regulated and increases substantially from late embryonic to postnatal period (21,23), when astrocyte progenitor cell proliferation declines and astrocyte maturation occurs (30,31). This temporal profile suggests that MLC1 may be implicated in the cellular processes leading to progenitor cell proliferation arrest and astrocyte differentiation/maturation. To verify this possibility, in this study we have investigated the relationship between unmutated and mutated MLC1 and cell proliferation, and the underlying molecular mechanisms in astrocytes.

Results

MLC1 expression inversely correlates with astrocyte proliferation

Preliminary experiments revealed substantial differences in endogenous MLC1 expression between astrocytes displaying markedly different proliferation rates (i.e. tumoral and primary astrocytes). Using western blotting (WB), we found that highly proliferating human U251 astrocytoma cells and rat C6 glioma cells express lower levels of MLC1 protein than the less proliferating primary human and rat astrocytes (Fig. 1A). MLC1 mRNA level was also higher in primary astrocytes than in tumoral astrocytes (Fig. 1B). We analyzed further the link between MLC1 protein expression and cell proliferation in U251 astrocytoma cell lines overexpressing recombinant MLC1 wild type (WT) or carrying a pathological missense mutation (MLC1-S280L) (25–27). Treatment of WT MLC1-expressing cells with the antiproliferative agent AraC (2 µM) for 5 days increased the amount of MLC1 protein (Fig. 1C). This finding is in line with the data obtained in primary rat astrocytes (12) and indicates the existence of an inverse relationship between MLC1 expression and astrocyte proliferation levels. To monitor the growth rate of MLC1-expressing cells, the level of Ki67, a proliferation marker expressed in all phases of the cell cycle, was investigated by immunofluorescence (IF) and real-time RT-PCR. These experiments showed that the percentage of proliferating Ki67+ cells (Fig. 2A) and the level of Ki67 mRNA (Fig. 2B) were significantly lower in astrocytoma cells expressing WT MLC1 than in control mock-infected U251 cells (Ø) and in U251 cells expressing mutated MLC1 (MLC1-S280L). Accordingly, RNA interference-mediated down-regulation of MLC1 mRNA in primary rat astrocytes resulted in an increase in the proliferation of MLC1 KO cells (Fig. 2C). Supporting further the negative control of MLC1 on astrocyte proliferation, we found that the number of MLC1+ cells (not shown) and MLC1 protein expression levels progressively decreased from the second to the seventh in vitro passage in WT MLC1-expressing U251 cells but not in U251 cells expressing mutated MLC1 (Fig. 1 SD). Analysis of the degradation kinetics of WT and mutant MLC1 protein in astrocytoma cells treated with the protein synthesis inhibitor cycloheximide (CHX) excluded the possibility that the decrease of MLC1 observed during cell passages was due to a lower stability of the WT protein compared with the mutant protein (Fig. 2 SD).

Figure 1.

MLC1 expression in primary and tumoral astrocytes. (A) WB analysis of MLC1 protein performed in human U251 astrocytoma cells and fetal astrocytes, rat C6 glioma cells and primary astrocytes reveals higher levels of MLC1 protein in primary astrocytes than tumoral astrocytes. Equal amounts of proteins were loaded on the gel, as shown by β-actin protein levels. One representative experiment out of three is shown. (B) MLC1 mRNA levels were evaluated by RT-PCR in the same cell cultures (as in A). Human and rat primary astrocytes expressed higher levels of MLC1 mRNA than tumoral astrocytes (U251 and C6). Normalization was performed using β-actin as control gene. One representative experiment out of three is shown. (C) Effect of the antimitotic agent AraC on MLC1 protein level. U251 cells expressing WT MLC1 were grown in the absence or presence of AraC (2 µM); after 5 days, protein extracts were analyzed by WB. An increase in the total amount of MLC1 protein is observed in the presence of the antimitotic agent. β-Actin is used as internal control for protein loading. Molecular weight markers (kDa) are indicated on the left side of A and C.

Figure 1.

MLC1 expression in primary and tumoral astrocytes. (A) WB analysis of MLC1 protein performed in human U251 astrocytoma cells and fetal astrocytes, rat C6 glioma cells and primary astrocytes reveals higher levels of MLC1 protein in primary astrocytes than tumoral astrocytes. Equal amounts of proteins were loaded on the gel, as shown by β-actin protein levels. One representative experiment out of three is shown. (B) MLC1 mRNA levels were evaluated by RT-PCR in the same cell cultures (as in A). Human and rat primary astrocytes expressed higher levels of MLC1 mRNA than tumoral astrocytes (U251 and C6). Normalization was performed using β-actin as control gene. One representative experiment out of three is shown. (C) Effect of the antimitotic agent AraC on MLC1 protein level. U251 cells expressing WT MLC1 were grown in the absence or presence of AraC (2 µM); after 5 days, protein extracts were analyzed by WB. An increase in the total amount of MLC1 protein is observed in the presence of the antimitotic agent. β-Actin is used as internal control for protein loading. Molecular weight markers (kDa) are indicated on the left side of A and C.

Figure 2.

WT MLC1 protein inhibits cell proliferation. (A) The proliferative activity of mock-infected U251 cells (Ø) and cell lines overexpressing MLC1 WT or carrying mutated S280L MLC1 was evaluated by immunostaining with anti-Ki67 pAb to count the nuclei of proliferating cells and DAPI to count total cell nuclei. The bar graph shows that in WT MLC1-expressing cells, the fraction of Ki67+ proliferating cells is significantly lower than that of control cells (Ø) and cells expressing mutated MLC1 (MLC1-S280L). Data are presented as percentage of Ki67+ cells in the total cell population. Means ± SEM of three experiments are shown (*P < 0.05, ***P < 0.001). (B) Real-time RT-PCR for Ki67 performed in control cells (Ø) and in WT and S280L MLC1-expressing cells confirms lower proliferative activity of WT MLC1-expressing cell line. Data are expressed as 2-ΔCt relative to GAPDH. Means ± SEM of three experiments are shown (*P < 0.05). (C) RT-PCR experiments using specific primers to detect MLC1 and Ki67 mRNA in primary rat astrocytes transfected with MLC1-specific shRNA or control shRNA. Down-regulation of MLC1 mRNA by shRNA (left panel) causes an increase in Ki67 mRNA level (right panel). Means ± SEM of three experiments are shown (*P < 0.05).

Figure 2.

WT MLC1 protein inhibits cell proliferation. (A) The proliferative activity of mock-infected U251 cells (Ø) and cell lines overexpressing MLC1 WT or carrying mutated S280L MLC1 was evaluated by immunostaining with anti-Ki67 pAb to count the nuclei of proliferating cells and DAPI to count total cell nuclei. The bar graph shows that in WT MLC1-expressing cells, the fraction of Ki67+ proliferating cells is significantly lower than that of control cells (Ø) and cells expressing mutated MLC1 (MLC1-S280L). Data are presented as percentage of Ki67+ cells in the total cell population. Means ± SEM of three experiments are shown (*P < 0.05, ***P < 0.001). (B) Real-time RT-PCR for Ki67 performed in control cells (Ø) and in WT and S280L MLC1-expressing cells confirms lower proliferative activity of WT MLC1-expressing cell line. Data are expressed as 2-ΔCt relative to GAPDH. Means ± SEM of three experiments are shown (*P < 0.05). (C) RT-PCR experiments using specific primers to detect MLC1 and Ki67 mRNA in primary rat astrocytes transfected with MLC1-specific shRNA or control shRNA. Down-regulation of MLC1 mRNA by shRNA (left panel) causes an increase in Ki67 mRNA level (right panel). Means ± SEM of three experiments are shown (*P < 0.05).

MLC1 affects EGFR activation

Most glioblastoma/astrocytoma cells, including U251, constitutively express the active phosphorylated form of the epidermal growth factor receptor (EGFR), which plays an essential role in promoting cell proliferation both in vivo and in vitro (32,33). We thus investigated whether MLC1 could influence astrocytoma cell proliferation by interfering with EGFR expression and activation.

Using WB with an Ab recognizing both the phosphorylated and non-phosphorylated forms of the EGFR, we found that the amount of phosphorylated EGFR (high-molecular-weight band) was significantly lower in WT MLC1-expressing cells than in mock-infected U251 cells (Ø) and in cells expressing MLC1 different pathological mutations (C125R, S280L, S246R) (Fig. 3A and B). No reduction in EGFR phosphorylation was observed in U251 astrocytoma cells stably expressing the potassium channel Kir2.1, excluding possible unspecific effects due to overexpression of a plasma membrane-associated protein (Fig. 3A and B). Accordingly, in WT MLC1-expressing astrocytoma cell cultures, EGFR immunoreactivity was observed in the cytoplasm and intracellular perinuclear vesicles of cells expressing high levels of MLC1 protein (≥50% of the total cell population), where it co-localized with MLC1 protein (Fig. 3C, asterisks), and was not detected in the plasma membrane where receptor phosphorylation occurs (34 and references therein) (Fig. 3C, arrowheads). On the contrary, in the cell subset expressing low levels of WT MLC1 and in astrocytoma cell cultures expressing mutated MLC1 (MLC1-S280L), EGFR was mainly localized in the plasma membrane (Fig. 3C, arrows). Altogether, these results indicate that WT, but not mutated, MLC1 inhibits EGFR translocation at the plasma membrane and ensuing phosphorylation.

Figure 3.

WT MLC1 inhibits EGFR phosphorylation. (A) WB analysis of proteins derived from mock-infected U251 astrocytoma cells (Ø) and U251 astrocytoma cells expressing MLC1 WT (WT), different MLC1 pathological missense mutations (C125R, S280L, S246R), or the potassium channel Kir2.1 reveals that EGFR is constitutively phosphorylated (pEGFR) in mock-infected U251 cells and astrocytoma cell lines expressing mutated MLC1 and Kir2.1, but not in cells expressing MLC1 WT. β-Actin is used as loading control. Molecular weight markers are indicated on the left (kDa). One representative experiment out of three performed is shown. (B) Densitometric analysis of phosphorylated EGFR protein bands after normalization with the amount of the β-actin was evaluated in the same samples shown in A. Means ± SEM of three experiments are shown (***P < 0.001). (C) Double IF staining of astrocytoma cells overexpressing WT and S280L MLC1 with anti-MLC1 pAb (red) and anti-EGFR mAb (green) shows that EGFR immunoreactivity is not detectable in the plasma membrane of WT MLC1-expressing cells (arrowheads), while it is present in the plasma membrane of cells expressing low levels of MLC1 and in cells expressing the S280L MLC1 mutation (arrows). In WT MLC1+ cells, EGFR is present in intracellular perinuclear areas where it co-localizes with MLC1 (asterisks). Scale bars: 10 μm.

Figure 3.

WT MLC1 inhibits EGFR phosphorylation. (A) WB analysis of proteins derived from mock-infected U251 astrocytoma cells (Ø) and U251 astrocytoma cells expressing MLC1 WT (WT), different MLC1 pathological missense mutations (C125R, S280L, S246R), or the potassium channel Kir2.1 reveals that EGFR is constitutively phosphorylated (pEGFR) in mock-infected U251 cells and astrocytoma cell lines expressing mutated MLC1 and Kir2.1, but not in cells expressing MLC1 WT. β-Actin is used as loading control. Molecular weight markers are indicated on the left (kDa). One representative experiment out of three performed is shown. (B) Densitometric analysis of phosphorylated EGFR protein bands after normalization with the amount of the β-actin was evaluated in the same samples shown in A. Means ± SEM of three experiments are shown (***P < 0.001). (C) Double IF staining of astrocytoma cells overexpressing WT and S280L MLC1 with anti-MLC1 pAb (red) and anti-EGFR mAb (green) shows that EGFR immunoreactivity is not detectable in the plasma membrane of WT MLC1-expressing cells (arrowheads), while it is present in the plasma membrane of cells expressing low levels of MLC1 and in cells expressing the S280L MLC1 mutation (arrows). In WT MLC1+ cells, EGFR is present in intracellular perinuclear areas where it co-localizes with MLC1 (asterisks). Scale bars: 10 μm.

MLC1 induces EGFR degradation

The lack of EGFR in the plasma membrane and its predominant distribution in intracellular organelles in WT MLC1-expressing U251 cells suggest that MLC1 could be involved in the control of EGFR trafficking/degradation. To verify this possibility, we treated U251 cells with epidermal growth factor (EGF), which stimulates EGFR phosphorylation at the plasma membrane and promotes subsequent endocytosis for receptor sorting to the recycling or degradation pathway (33–36), and investigated EGFR degradation in the presence or absence of the protein synthesis inhibitor CHX, as previously described (37). WB showed that in WT MLC1-expressing cells, EGFR phosphorylation was induced within 30 min treatment with EGF/CHX and markedly reduced after 1 h (Fig. 4A and B). In contrast, in S280L MLC1-expressing cells, EGFR was abundantly phosphorylated both in basal conditions and after treatment with EGF/CHX for 1 h (Fig. 4A and B). In both cell lines, EGFR was almost undetectable after 4 h EGF/CHX treatment (Fig. 4A and B). We next evaluated EGFR activation/degradation kinetics in a more physiologically relevant setting, such as primary rat astrocytes, which express constitutively both MLC1 and EGFR (22,38). Similarly to what observed in WT MLC1-expressing astrocytoma cells, EGFR was phosphorylated in primary astrocytes after 15–30 min of EGF stimulation and rapidly degraded after 1 h treatment in the presence of CHX (Fig. 4C and D).

Figure 4.

WT MLC1 induces EGFR degradation. (A) WB analysis of protein extracts from WT and S280L mutated MLC1 protein expressing U251 cells treated with EGF for 30 min, 1 and 4 h, in the absence (CTRL) or presence of the protein synthesis inhibitor CHX indicated that EGFR is phosphorylated (upper band) after 30 min of EGF activation and rapidly degraded after 1 h in WT but not in S280L MLC1-expressing cells. The excitatory amino acid transporter-2 (EAAT2) is used as loading control. Molecular weight markers are indicated on the left (kDa). (B) Densitometric analysis of EGFR protein bands revealed by WB after normalization with EAAT2 protein levels in the same samples. Data are expressed as percentage of the value measured in control untreated cells (100%). Means ± SEM of three experiments are shown (*P < 0.05). (C) WB analysis of protein extracts derived from primary rat astrocytes, untreated (CTRL) or treated with EGF + CHX for 15 min, 30 min and 1 h shows that in these cells EGFR is phosphorylated after 15 min stimulation and degraded after 1 h. β-Actin is used as loading control. Molecular weight markers are indicated on the left (kDa). (D) Densitometric analysis of EGFR protein bands in primary astrocytes revealed by WB and normalized for protein content with β-actin. Data are expressed as percentage of the value measured in control untreated cells (100%). Means ± SEM of three experiments are shown (***P < 0.001).

Figure 4.

WT MLC1 induces EGFR degradation. (A) WB analysis of protein extracts from WT and S280L mutated MLC1 protein expressing U251 cells treated with EGF for 30 min, 1 and 4 h, in the absence (CTRL) or presence of the protein synthesis inhibitor CHX indicated that EGFR is phosphorylated (upper band) after 30 min of EGF activation and rapidly degraded after 1 h in WT but not in S280L MLC1-expressing cells. The excitatory amino acid transporter-2 (EAAT2) is used as loading control. Molecular weight markers are indicated on the left (kDa). (B) Densitometric analysis of EGFR protein bands revealed by WB after normalization with EAAT2 protein levels in the same samples. Data are expressed as percentage of the value measured in control untreated cells (100%). Means ± SEM of three experiments are shown (*P < 0.05). (C) WB analysis of protein extracts derived from primary rat astrocytes, untreated (CTRL) or treated with EGF + CHX for 15 min, 30 min and 1 h shows that in these cells EGFR is phosphorylated after 15 min stimulation and degraded after 1 h. β-Actin is used as loading control. Molecular weight markers are indicated on the left (kDa). (D) Densitometric analysis of EGFR protein bands in primary astrocytes revealed by WB and normalized for protein content with β-actin. Data are expressed as percentage of the value measured in control untreated cells (100%). Means ± SEM of three experiments are shown (***P < 0.001).

Consistent with the WB data, stimulation with EGF for 30 min in the presence of CHX induced the appearance of EGFR+ intracellular vesicles in both WT and mutant MLC1-expressing astrocytoma cells, as result of EGF-induced EGFR endocytosis (Fig. 5A and B). However, after 1 h treatment with EGF/CHX, the number of EGFR+ intracellular vesicles was strongly reduced in WT compared with S280L MLC1-expressing cells (arrowheads in Fig. 5A and B, respectively; Fig. 5C). Noteworthy, in WT MLC1-expressing cells, MLC1 and EGFR co-localized in intracellular organelles at both time periods (asterisks in Fig. 5A), suggesting a common endocytic route for the two proteins. Altogether, the above results demonstrate that expression of WT, but not mutated, MLC1 favors the degradation of the activated EGF/EGFR complex.

Figure 5.

Immunolocalization of EGFR in response to EGF stimulation in astrocytoma cells expressing WT and mutant MLC1. Double IF staining with anti-MLC1 pAb (red) and anti-EGFR mAb (green) was carried out in WT (A) and S280L MLC1 (B) expressing U251 astrocytoma cells treated with EGF for 30 min and 1 h, in the absence or presence of the protein synthesis inhibitor CHX. In untreated WT MLC1+ cells (CTRL), EGFR was not detected in the plasma membrane (empty arrows), whereas it is localized in cells expressing low levels of MLC1 or expressing the S280L mutant (arrows). After 30 min exposure to EGF, EGFR was mainly localized in intracellular vesicles in both WT and mutant MLC1 (MLC1-S280L) expressing cells, as the result of EGF-induced endocytosis. After 1 h exposure to EGF, the number of EGFR+ intracellular vesicles is strongly diminished in WT MLC1+ compared with mutant expressing cells (arrowheads in A versus B). In WT MLC1+ cells, EGFR is present in intracellular perinuclear areas where it co-localizes with MLC1 (asterisks). Scale bars: 10 μm. (C) According to mean fluorescence intensity (MFI) analysis, EGFR fluorescence intensity after 1 h exposure to EGF is strongly diminished in WT MLC1+ when compared with mutant expressing cells.

Figure 5.

Immunolocalization of EGFR in response to EGF stimulation in astrocytoma cells expressing WT and mutant MLC1. Double IF staining with anti-MLC1 pAb (red) and anti-EGFR mAb (green) was carried out in WT (A) and S280L MLC1 (B) expressing U251 astrocytoma cells treated with EGF for 30 min and 1 h, in the absence or presence of the protein synthesis inhibitor CHX. In untreated WT MLC1+ cells (CTRL), EGFR was not detected in the plasma membrane (empty arrows), whereas it is localized in cells expressing low levels of MLC1 or expressing the S280L mutant (arrows). After 30 min exposure to EGF, EGFR was mainly localized in intracellular vesicles in both WT and mutant MLC1 (MLC1-S280L) expressing cells, as the result of EGF-induced endocytosis. After 1 h exposure to EGF, the number of EGFR+ intracellular vesicles is strongly diminished in WT MLC1+ compared with mutant expressing cells (arrowheads in A versus B). In WT MLC1+ cells, EGFR is present in intracellular perinuclear areas where it co-localizes with MLC1 (asterisks). Scale bars: 10 μm. (C) According to mean fluorescence intensity (MFI) analysis, EGFR fluorescence intensity after 1 h exposure to EGF is strongly diminished in WT MLC1+ when compared with mutant expressing cells.

It has been shown that in normal and tumoral cells, EGFR is partially localized in lipid rafts, the plasma membrane sub-compartments enriched in cholesterol and sphingolipids and that this localization is associated with either inactivation or degradation of EGFR induced by raft-dependent endocytosis (39,40). Since MLC1 is expressed in lipid rafts in primary rat astrocytes and U251 cells (19,20), we investigated whether EGFR degradation observed in WT MLC1-expressing U251 cells could be due to MLC1-induced receptor sequestration in caveolar lipid raft compartments. WB of raft-derived proteins (22) revealed no differences in EGFR distribution between WT and S280L MLC1-expressing cells showing that EGFR was localized in non-caveolar raft compartments in both cell lines (Fig. 3 SD). Based on these findings, it is unlikely that MLC1-mediated inactivation/degradation of EGFR occurs by means of raft-dependent endocytosis (Fig. 3 SD).

EGFR distribution is altered in macrophages from MLC patients

Blood-derived macrophages do express both MLC1 and EGFR (15,41,42). Thus, we investigated EGFR expression and intracellular distribution in macrophages obtained from three MLC patients (Pt1, Pt2 and Pt3) and three healthy controls. In control macrophages MLC1 immunoreactivity was distributed throughout the cytoplasm and in the plasma membrane (Fig. 6A). As previously described (41), MLC1 expression in macrophages from patients was lower than in control cells, particularly in Pt2 and Pt3 (Fig. 6A). Notably, it was found that in healthy macrophages, the expression of EGFR was mainly restricted to the nucleus and weakly detected in the cytoplasm (Fig. 6A). The expression of EGFR was significantly higher in both nucleus and cytoplasm of macrophages derived from Pt1 and, even more, in those from Pt2 and Pt3, showing an inverse relationship with MLC1 expression levels (Fig. 6A and B). These results indicated that MLC1 mutations can modify EGFR protein expression and distribution also in macrophages, strengthening the conclusion that MLC1 down-regulates EGFR expression and, possibly, function.

Figure 6.

Confocal imaging of MLC1 and EGFR in macrophages from healthy controls and MLC patients (Pt1, Pt2 and Pt3). (A) MLC1 is distributed in the cytoplasm and plasma membrane of control macrophages, whereas it is essentially confined around the nucleus in macrophages from Pt1, Pt2 and Pt3. EGFR distribution is detected in the nucleus and faintly in the cytoplasm of control macrophages, whereas in Pt1 and, to a higher degree in Pt2 and Pt3 macrophages, EGFR expression significantly increases in both nucleus and cytoplasm. Nuclei have been stained with Hoechst. Scale bar: 40 μm. (B) MFI of EGFR in nuclear and cytoplasmic compartments as well as in the total cell area of control (CTRL), Pt1, Pt2 and Pt3 macrophages. Data are presented as mean ± SEM (***P < 0.001, **P < 0.01).

Figure 6.

Confocal imaging of MLC1 and EGFR in macrophages from healthy controls and MLC patients (Pt1, Pt2 and Pt3). (A) MLC1 is distributed in the cytoplasm and plasma membrane of control macrophages, whereas it is essentially confined around the nucleus in macrophages from Pt1, Pt2 and Pt3. EGFR distribution is detected in the nucleus and faintly in the cytoplasm of control macrophages, whereas in Pt1 and, to a higher degree in Pt2 and Pt3 macrophages, EGFR expression significantly increases in both nucleus and cytoplasm. Nuclei have been stained with Hoechst. Scale bar: 40 μm. (B) MFI of EGFR in nuclear and cytoplasmic compartments as well as in the total cell area of control (CTRL), Pt1, Pt2 and Pt3 macrophages. Data are presented as mean ± SEM (***P < 0.001, **P < 0.01).

Downstream effects of MLC1-induced EGFR down-regulation: inhibition of Ca2+ entry and of ERK1/2 and PLCγ1 activation

In many cell types, including astrocytes, the first step of EGFR-induced cell proliferation involves intracellular calcium (Ca2+) elevation followed by activation of MAPK/ERK and PLCγ1-associated signal transduction pathways (43,44). These are also the main intracellular pathways involved in the proliferative response of astrocytes in vivo and in vitro (45,46). To investigate whether MLC1 can affect EGFR downstream pathways, we first used the FURA-2-based fluorescence video imaging technique to monitor intracellular Ca2+ alterations in U251 astrocytoma cells expressing WT or mutated MLC1 after EGFR stimulation. EGF application caused in virtually all cells a low amplitude raise of intracellular Ca2+, reaching steady state in ∼2 min application. Figure 7A shows a fluorimetric Ca2+ recording upon sequential EGF (200 ng/ml) and ATP (10 μM) applications, pointing out the slow kinetic and low amplitude of the EGF-induced signal. Noteworthy, the EGF-induced Ca2+ response showed a higher amplitude in S280L mutant than in WT MLC1-expressing cells, most likely due to the presence of a higher density of functional EGFRs (Fig. 7B and C). The distributions of the amplitudes of Ca2+ signals show that the average increase in mutant compared with WT MLC1-expressing cells is mainly due to a general increase of the response in all the cells, but also to the appearance of a small population of cells displaying a very high Ca2+ response (Fig. 7D). To characterize the nature of the Ca2+ signal, we used ion substitution experiments and pharmacological tests. The removal of Ca2+ (replaced by Mg2+) in the external solution caused a strong reduction in the EGF-induced Ca2+ increase, indicating that Ca2+ raise was mainly due to Ca2+ influx and only slightly to Ca2+ release from intracellular compartments, such as the endoplasmic reticulum (Fig. 7E). To identify the Ca2+ channels responsible for the EGF-induced Ca2+ signals, we used the store-operated channels (SOC) inhibitor SKF-96365, known to inhibit Ca2+ influx from TRP type of Ca2+ channels. When applied before and during EGF, SKF-96365 almost completed prevented the Ca2+ response. The inhibitory effect of SKF-96365 was also evident when it was applied during the EGF challenge (Fig. 7F) leading to hypothesize the involvement of TRP channels in the EGF-induced Ca2+ response. The finding that cell treatment with PD98058, the MEK/ERK kinase specific inhibitor, did not abrogate the EGF-induced Ca2+ response (data not shown) suggests that Ca2+ influx might lay upstream of ERK1/2 activation. We then analyzed the effects of MLC1-induced EGFR down-regulation on MAP kinase ERK1/2, PLCγ and AKT expression and activation by WB using Abs specific for the phosphorylated forms of these enzymes. The results obtained showed that the phosphorylated forms of both ERK1/2 and PLCγ1 were down-regulated in WT MLC1-expressing U251 cells when compared with S280L mutant expressing cells and cells infected with the empty vector (Fig. 8A and B), while no differences were observed in the expression of the phosphorylated form of AKT, another kinase activated by EGF that is mainly involved in cell survival processes (Fig. 8A and B).

Figure 7.

EGF induces intracellular Ca2+ raise. (A) Fura-2-loaded astrocytoma cells were challenged with EGF (200 ng/ml for 3 min) and then with ATP (10 μM for 30 sec). EGF induced a slow raise of Ca2+, while ATP triggered a fast Ca2+ transient of a larger amplitude. (B) The panel shows exemplificative traces (mean ± SEM) representing EGF-induced Ca2+ responses recorded from cells expressing WT MLC1 (triangles) and mutant S280L MLC1 (circles). A larger amplitude of the signal is detected in S280L cells, as shown by the average amplitude of the EGF-induced Ca2+ signals (C) and the amplitude distributions of the responses recorded in single cells (D) (207–375 cells were analyzed; ***P < 0.001). (E) When recorded in Ca2+-free condition most of the Ca2+ signal observed was abrogated (Ca2+ free: squares; CTRL: circles), depicting a major contribution of Ca2+ influx to the Ca2+ raise. (F) The TRP inhibitor SKF-96365 (10 μM) almost completely abrogated the Ca2+ response, both when applied before and during the EGF challenge.

Figure 7.

EGF induces intracellular Ca2+ raise. (A) Fura-2-loaded astrocytoma cells were challenged with EGF (200 ng/ml for 3 min) and then with ATP (10 μM for 30 sec). EGF induced a slow raise of Ca2+, while ATP triggered a fast Ca2+ transient of a larger amplitude. (B) The panel shows exemplificative traces (mean ± SEM) representing EGF-induced Ca2+ responses recorded from cells expressing WT MLC1 (triangles) and mutant S280L MLC1 (circles). A larger amplitude of the signal is detected in S280L cells, as shown by the average amplitude of the EGF-induced Ca2+ signals (C) and the amplitude distributions of the responses recorded in single cells (D) (207–375 cells were analyzed; ***P < 0.001). (E) When recorded in Ca2+-free condition most of the Ca2+ signal observed was abrogated (Ca2+ free: squares; CTRL: circles), depicting a major contribution of Ca2+ influx to the Ca2+ raise. (F) The TRP inhibitor SKF-96365 (10 μM) almost completely abrogated the Ca2+ response, both when applied before and during the EGF challenge.

Figure 8.

WT MLC1 inhibits phosphorylation of ERK1/2 and PLCγ1 but not of AKT. (A) WB analysis of the phosphorylation level of ERK1/2, PLCγ1 and AKT in mock-infected cells (Ø) and in cells expressing WT or S280L mutant MLC1. The phosphorylated forms of ERK1/2 and PLCγ1 were significantly less abundant in WT MLC1+ cells than in mock-infected and S280L mutant expressing U251 cells. No differences were observed in AKT phosphorylation. (B) Densitometric analysis of pERK1/2, pPLCγ1 and pAKT after normalization with the non-phosphorylated proteins ERK and β-actin and AKT, respectively. The bar graphs represent the means ± SEM of three independent experiments expressed as percentage of the values measured in U251 Ø (taken as 100%; *P < 0.05, **P < 0.01).

Figure 8.

WT MLC1 inhibits phosphorylation of ERK1/2 and PLCγ1 but not of AKT. (A) WB analysis of the phosphorylation level of ERK1/2, PLCγ1 and AKT in mock-infected cells (Ø) and in cells expressing WT or S280L mutant MLC1. The phosphorylated forms of ERK1/2 and PLCγ1 were significantly less abundant in WT MLC1+ cells than in mock-infected and S280L mutant expressing U251 cells. No differences were observed in AKT phosphorylation. (B) Densitometric analysis of pERK1/2, pPLCγ1 and pAKT after normalization with the non-phosphorylated proteins ERK and β-actin and AKT, respectively. The bar graphs represent the means ± SEM of three independent experiments expressed as percentage of the values measured in U251 Ø (taken as 100%; *P < 0.05, **P < 0.01).

Effect of MLC1 expression on KCa3.1 functionality

Intermediate conductance Ca2+-activated potassium channels (KCa3.1) are expressed in astrocytes selectively at the end-feet surrounding blood vessels, where the MLC1 protein is mainly localized in the brain, and are functionally relevant for cell proliferation and neurovascular coupling (47). In a previous study, we found that KCa3.1 channel expression in astrocytoma cells depends on ERK1/2 activity (48). Given the significant inhibition of ERK1/2 signal transduction pathway as a consequence of the MLC1-induced EGFR degradation, we verified whether the KCa3.1 channel functionality was altered in MLC1-expressing U251 cells. KCa3.1 channel activity in U251 astrocytoma cells was recorded using ionic conditions suitable for the study of K+ selective currents (see the Materials and Methods section), in the presence of 3 mm extracellular TEA to block the large-conductance Ca2+-activated K+ channels expressed in these cells (49). In these conditions, application of the KCa2/KCa3 channel opener DC-EBIO (100 μM) plus the Ca2+ agonist ionomycin (0.5 μM; EBIO/iono) activated a voltage-independent current with a reversal potential close to the K+ equilibrium potential, and blocked by the selective KCa3.1 channel inhibitor clotrimazole (CTL; 10 μM; Fig. 9). The CTL-sensitive current activated by EBIO/iono had a mean density of 4.3 ± 0.4 pA/pF at 0 mV (n = 4) and was present in all U251 cells examined. Notably, we found that U251 cells stably expressing WT MLC1 expressed a significantly lower KCa3.1 current density when compared with U251 cells infected with an empty vector (2.2 ± 0.3 pA/pF; n = 7; P < 0.001; Fig. 9), suggesting that the expression of MLC1 negatively modulates this channel. In contrast, U251 cells stably expressing mutated MLC1 (MLC1-S280L) had KCa3.1 current densities that did not differ significantly from those found in cells infected with the empty vector (3.7 ± 0.5 pA/pF; n = 7; P > 0.05), but were significantly higher than those present in WT MLC1-expressing cells (P < 0.05; Fig. 9).

Figure 9.

MLC1 down-regulates KCa3.1 current in U251 cells. (A) Time course of current at 0 mV. Inset: sample current ramps recorded from −100 to + 100 mV (from a Vh of −30 mV, duration 1 s, repeated every 5 s) in control condition (white line, CTRL), following the application of Ionomyicin 500 nM and DCEBIO 100 μM (black line, EBIO/iono) and following CTL 10 μM (white line). (B) Bar plot showing KCa3.1 current density (pA/pF) in control (Ø) and U251 cells expressing WT or mutated (MLC1-S280L) MLC1 (*P < 0.05, ***P < 0.001).

Figure 9.

MLC1 down-regulates KCa3.1 current in U251 cells. (A) Time course of current at 0 mV. Inset: sample current ramps recorded from −100 to + 100 mV (from a Vh of −30 mV, duration 1 s, repeated every 5 s) in control condition (white line, CTRL), following the application of Ionomyicin 500 nM and DCEBIO 100 μM (black line, EBIO/iono) and following CTL 10 μM (white line). (B) Bar plot showing KCa3.1 current density (pA/pF) in control (Ø) and U251 cells expressing WT or mutated (MLC1-S280L) MLC1 (*P < 0.05, ***P < 0.001).

Discussion

The main finding of this study is that MLC1 is involved in the control of cell proliferation and EGFR activation in brain astrocytes. Using different strategies to modulate MLC1 expression in primary and tumoral astrocytes, we show that MLC1 hampers astrocyte proliferation by favoring EGFR degradation and down-regulation of EGF-activated Ca2+ entry and ERK1/2 and PLCγ1-associated signaling pathways, as depicted in Fig. 10. Moreover, we also found that MLC1 causes functional inactivation of the KCa3.1 channel that is primarily responsible for astrocyte activation in response to brain injury and for cell proliferation (50–52). Altogether, these data highlight a potential role for MLC1 in the control of astrocyte development and activation in response to pathological stimuli, and provide new insights into the molecular pathogenesis of MLC disease.

Figure 10.

Schematic representation of the EGFR-activated pathways that are inhibited by MLC1 in human U251 astrocytoma cells.

Figure 10.

Schematic representation of the EGFR-activated pathways that are inhibited by MLC1 in human U251 astrocytoma cells.

MLC1 affects EGFR and associated signaling pathways

The EGFR is a membrane receptor with intrinsic tyrosine kinase activity that is activated upon binding of EGF or other growth factors of the EGF family. EGFR activation stimulates multiple signaling cascades that regulate several physiological processes including cell proliferation, differentiation and death (31–34). This study shows that in U251 astrocytoma cells, where EGFR is constitutively activated, overexpression of WT, but not of mutated MLC1 or an unrelated membrane protein (Kir2.1), down-regulates EGFR by inducing its degradation. The co-localization of EGFR and MLC1 in intracellular organelles after EGF-induced endocytosis suggests a possible effect of MLC1 on EGFR intracellular trafficking/sorting after receptor internalization. Membrane receptor endocytosis is a strictly regulated mechanism to turn off receptor activation, and alterations of receptor trafficking toward the recycling or degradation pathways can affect the duration, magnitude and specificity of EGFR signaling, thus influencing basilar cell biology processes (53). Although the molecular mechanisms through which MLC1 modifies EGFR endosomal sorting remain to be elucidated, the effect of MLC1 on organelle pH could play a role (27). Organelle acidification (from 6.0 in early endosomes to 5.0–5.5 in lysosomes) controls EGFR receptor–ligand dissociation after endocytosis and its trafficking toward the recycling or degradation pathways (54,55). We recently demonstrated that MLC1 decrease early endosome acidification by interacting with the V-ATPase (27) and this may in turn influence intracellular EGF/EGFR complex dissociation occurring at intraorganelle acidic pH, thus favoring receptor degradation. Alternatively, a less acidic intraorganelle pH could indirectly influence the activity of kinases involved in EGFR trafficking in the endosomal compartment (56). Interestingly, in breast cancer cells, the tumor suppressor protein BIF-1, which increases intraorganelle pH, induces EGFR degradation and cell proliferation arrest (54). It has been proposed that intracellular sorting of the EGF/EGFR complex is differently influenced by EGFR distribution in caveolar lipid raft compartments (40,57). However, we can exclude this possibility since we did not find differences in EGFR raft distribution between WT and mutant MLC1-expressing cells. As expected, we also found that WT, but not mutated, MLC1 was also able to inhibit many of the EGFR-dependent intracellular signal transduction pathways, such as intracellular Ca2+ influx, and ERK1/2 kinase and PLCγ1 phosphorylation. Ca2+ is one of the key regulators of DNA synthesis and cell proliferation, and EGF stimulation increases cytoplasmic free calcium concentration by inducing both an extracellular Ca2+ influx mediated by TRP Ca2+ channels (44) and a Ca2+ release from the endoplasmic reticulum (43). In U251 cells, we found that EGF induces a slow raise of cytoplasmic Ca2+, an effect mainly due to Ca2+ influx through TRP Ca2+ channels, as demonstrated by the inhibitory effect of SKF-96365. Noteworthy, the amplitude of the Ca2+ response was significantly higher in astrocytoma cells expressing mutated MLC1 than in WT MLC1-expressing cells.

The Ras/Raf/MEK/ERK pathway is involved in the control of cell growth, survival and invasion, and ERK1/2 activation occurs in developing astrocytes (46) and in adult astrocytes in response to injury and inflammatory stimuli (58–60). In addition in different cell types, EGFR and ERK activation plays an important role in the control of volume changes and RVD activation (29,61–63). RVD is one of the physiological processes in which MLC1 has been implicated (23–25). In astrocytes, EGFR/ERK activation of osmosensitive Cl fluxes occurs upon hypotonicity-induced cell swelling (64,65), while PLCγ1 regulates astrocyte swelling in response to oxidative stress (66). MLC1 can thus influence volume changes and RVD in astrocytes by down-modulating EGFR/ERK/PLCγ1 pathways in response to physiological and pathological stimuli.

The KCa3.1 channel is a voltage-independent, K+-selective channel activated by the binding of Ca2+ ions to calmodulin molecules constitutively associated with the channel protein (67,68). Relatively small increases of intracellular Ca2+ (few hundreds of nanomolars) induce channel opening, resulting in K+ efflux and maintenance of a driving force for subsequent Ca2+ influx. These channels play a key role in regulating Ca2+ signaling in a variety of cells (49,68,69). In astrocytes, KCa3.1 channels are moderately expressed in normal conditions and contribute to serum-induced proliferation (70). Interestingly, immunohistochemical, molecular and electrophysiological studies show that KCa3.1 channels are abundantly expressed in astrocyte processes and perivascular end-feet, where they participate to neurovascular coupling by inducing vasodilatation (47). As MLC1 is specifically expressed in astrocytes contacting CNS blood vessels, the present findings suggest that MLC1 could regulate blood–brain barrier permeability by modulating KCa3.1 functionality with the result that alteration in this astrocytic function by MLC1 mutations could contribute to the development of brain edema in MLC patients.

EGFR down-regulation: relevance for astrocyte activation and MLC pathogenesis

In the developing CNS, the EGFR pathway is implicated in astrocyte migration, differentiation, response to pathological insult and support of neuronal function (71,72). In mice, EGFR is abundantly expressed in nestin+ astrocyte progenitors that actively proliferate in the subventricular zone, and decreases during astrocyte differentiation in the late embryonic to postnatal period (72). Conversely, MLC1 is not expressed in nestin+ progenitor cells, while it increases from embryonic to postnatal life reaching a peak around Postnatal Days 5–10 to stabilize in the adult life (23). These data, along with our observation that MLC1 expression in both tumoral astrocytes and primary rat astrocytes is inversely related to proliferation, indicate that MLC1 most likely plays a specific role in the astrocyte proliferation/maturation switch during the late phase of brain development. Along this line, it can be hypothesized that astrocyte defective maturation during brain development of MLC patients may alter myelinogenesis, leading to myelin vacuolation, as observed in vanishing white matter disease (VWM/CASH), another vacuolating leukodystrophy (73). Since U251 astrocytoma cells exhibit some morpho-functional features of low differentiated fetal astrocytes (74), they can be considered a good model to study the developmental role of MLC1. Moreover, the functional characterization of astrocytoma cells overexpressing MLC1 indicated that MLC1 favors the activation of a swelling-induced chloride current and RVD (manuscript in preparation), as observed in primary astrocytes and patient-derived lymphoblasts (24), confirming the usefulness of this experimental system to study MLC1 function. In our experiments, we observed that MLC1 expression in U251 cells induces EGFR degradation with a kinetics similar to that found in rat primary astrocytes, suggesting that MLC1 protein may behave as tumor suppressor gene. Interestingly, Hepacam/Glialcam, the cell adhesion protein which binds MLC1 and whose mutations are present in a small percentage of MLC patients (75), behaves like a tumor suppressor gene by inducing cell proliferation arrest (76,77). However, MLC patients affected by mutations in MLC1 or Hepacam/Glialcam do not show increase in brain tumor incidence, suggesting that other factors are involved in the complex events leading to tumorigenesis in vivo. The evidence that mutation-induced decrease of MLC1 protein in macrophages from MLC patients leads to an increase of EGFR expression reinforces the hypothesis of the involvement of this pathway in MLC pathogenesis. In the adult life, EGFR re-expression is a key signaling pathway in the activation of astrocytes after brain injury (78,79). Astrocyte activation and proliferation in response to injury have been demonstrated in several diseases ranging from neurodegenerative and inflammatory conditions to acute invasive brain injury, such as trauma and stroke (78,79). During these processes, astrocytes undergo a functional de-differentiation, manifested as re-expression of EGFR, nestin and vimentin, and reactivation of downstream signaling pathways, like Ras, Raf and ERK, in response to EGF (38,71,72,80) and to other inflammatory mediators, like COX-2, PGE2 and nitric oxide (38,81–83). Conversely, blocking EGFR attenuates reactive astrogliosis (71,84). In this context, our results suggest a role for MLC1 in down-regulating the astrocyte response to injury. Aggravation of the clinical conditions in MLC patients after mild trauma or common infections could be the result of the inability of astrocytes expressing a mutated, and hence dysfunctional, MLC1 protein, to down-modulate their own reactivity in response to different brain insults. Consistent with this scenario, astrogliosis has been observed in brain biopsies of MLC patients (10,11). Our study has also revealed a new effect of MLC1 on the functionality of KCa3.1 channels in astrocytoma cells. In astrocytes, KCa3.1 channels are strongly up-regulated in several pathological conditions leading to astrocyte activation such as reactive astrogliosis induced in vitro by TGF-β or in vivo by spinal cord injury (50,70). Notably, the KCa3.1 inhibitor TRAM-34 prevented TGF-β-induced astrogliosis, improved tissue protection, reduced proinflammatory chemokine/cytokine expression, inhibited Na+ influx and reduced brain edema (70,85,86). Our results show that WT, but not mutated, MLC1 was able to down-modulate the functionality of KCa3.1 channels in U251 astrocytoma cells, most likely as a consequence of the reduced ERK1/2 activity, since its inhibition by PD98058 leads to a strong reduction of KCa3.1 currents (48). All these data suggest that KCa3.1 channels play a role in the altered ion and fluid homeostasis in the damaged CNS and that MLC1 could regulate astrocyte activation by indirectly inhibiting these channels.

MLC disease is also characterized by the development of subcortical cysts. Interestingly, dysregulation of EGFR has been demonstrated in the autosomal dominant polycystic kidney disease (ADPKD), which is caused by mutations in either PKD1 or PKD2 gene and is characterized by the formation and progressive growth of cystic lesions that ultimately destroy the renal parenchyma (87). In ADPKD, cyst growth is the result of sustained proliferation of incomplete or de-differentiated epithelial cells and accumulation of fluid within the cysts. Inhibition of EGFR tyrosine kinase activity, either genetically or pharmacologically, significantly reduces renal cyst formation and improves renal function in rodent models of PKD (87). It can be hypothesized that a similar pathological process leads to cyst formation in the brain of MLC patients.

Conclusions

This study discloses for the first time an important role for MLC1 in the control of astrocyte growth and in the regulation of pathways that trigger quiescent astrocytes into reactive ones in response to brain injury. It also shows that MLC1 pathological mutations cause loss of this function, opening new perspectives for the comprehension of MLC disease pathogenesis. Since astrogliosis is a complex pathological process that occurs in a variety of CNS pathological conditions, elucidation of MLC1 protein function in the regulation of astrocyte protective versus detrimental responses could reveal new potential therapeutic targets for several neuroinflammatory and neurodegenerative diseases.

Materials and Methods

Cell cultures and treatments

The human U251 MG astrocytoma cell line and the rat C6 glioma cell line were obtained from the American Type Culture Collection (Rockville, MD) and grown in Dulbecco's modified Eagle's medium-high glucose (DMEM, Euroclone Ltd, UK) supplemented with 10% FBS (Gibco BRL, Paisley, UK) and 1% penicillin/streptomycin (Sigma Ltd, Irvine, UK), at 37°C in a 5% CO2/95% air atmosphere. Establishment, characterization and culture conditions of U251 astrocytoma cell lines overexpressing His-tagged WT or mutant (S280L, C125R, S246R) MLC1 have been reported previously (25–27). To evaluate MLC1 protein expression in U251- and U251-derived cell lines, mock-infected (Ø), WT and S280L MLC1-expressing cells were sub-cultured and collected at different passages. For cell treatments, astrocytoma cell lines were plated in 100 mm diameter dishes in serum-free medium and treated with CHX (100 μg/ml, Sigma) alone or in combination with EGF (200 ng/ml, Sigma) for different time lengths [3, 6 or 12 h (overnight, ON)] or with cytosine arabinoside (AraC 2 μM, Sigma) for 5 days. After stimulation, the cells were washed in PBS, collected by scraping and centrifuged at 2.700g at 4°C for 20 min. Cell pellets were solubilized as described below. Primary astrocyte cultures were generated from 1- to 2-day-old newborn rats, as previously described (22). Cultures enriched in human fetal astrocytes were established, grown at confluence and stored in liquid nitrogen at the second in vitro passage (22). After thawing, human astrocytes were grown for 2–3 days in DMEM-high glucose (Euroclone) supplemented with 10% FBS (Gibco BRL), and 1% penicillin/streptomycin (Sigma) at 37°C in a humidified 5% CO2/95% air atmosphere.

MLC1 RNA interference

Primary astrocytes were plated at 2 × 105 in 3 ml into a poly-lysinated six-well plates and grown overnight in a 5% CO2 incubator to achieve 50% confluence. Then, the cells were transfected with a HuSHTM shRNA Plasmid Panels-29mer targeting MLC1 (Origene, Rockville, MD, USA) or with control shRNA plasmid using Lipofectamine reagent (Life Technologies, Grand Island, NY, USA), following the manufacturer's instructions. At 48 h post-transfection, cells were harvested for RNA analysis as described below.

Total RNA extraction and RT-PCR

Total RNA derived from rat and human astrocytes, U251, C6 glioma cells and rat astrocytes transfected with shRNA was purified using SV Total RNA Isolation System (Promega, Madison, WI, USA). To carry out the RT reaction, AMV Reverse Transcriptase (Promega) was used employing oligo (dT)15 primers, according to the manufacturer's instructions. PCR reactions for human or rat MLC1 and human or rat β-actin (to normalize MLC1 mRNA levels in the different cell cultures) were performed as previously described (22). PCRs for rat Ki67 were performed using the following primers: forward 5′ ATT TCA GTT CCG CCA ATC C 3′; reverse 5′ GGC TTC CGT CTT CAT ACC TAA A 3′. PCRs were performed for 35 cycles in the following conditions: 60 s/94°C, 60 s/60°C and 180 s/72°C.

Immunofluorescence staining and confocal microscopy analysis

Cells were grown to subconfluency on polylysine-coated coverslips, and fixed and stained as described previously (25–27). The following primary antibodies (Abs) were used: affinity purified anti-MLC1 polyclonal Ab (pAb) (1:50, Atlas AB, AlbaNova University Center, Stockholm, Sweden), anti-Xpress monoclonal Ab (mAb) (1:100, Invitrogen, Carlsbad, CA, USA, to detect the X-press epitope present at the NH2 terminal of the recombinant MLC1) and anti-EGFR mAb (1:40, Invitrogen). As secondary Abs, biotin-SP-AffiniPure goat anti-rabbit IgG H + L (4.3 μg/ml; Jackson Immunoresearch Laboratories, West Grove, PA, USA) followed by incubation with 2 μg/ml streptavidin-TRITC (Jackson, UK), or Alexa Fluor 488 goat anti-mouse IgG (1:300, Invitrogen) was used. Coverslips were washed, sealed in Vectashield medium (Vector Lab, Burlingame, CA, USA) and analyzed with a laser scanning confocal microscope (LSM 5 Pascal, Carl Zeiss, Jena, Germany). NIH ImageJ software (http://rsb.info.nih.gov/ij/) was used for IF quantification, by means of threshold fluorescence intensity analysis within a region of interest corresponding to a single-cell profile.

Confocal imaging of macrophages from MLC patients and healthy controls

Monocytes were prepared from peripheral blood mononuclear cells of healthy subjects and MLC patients {Patient 1 (Pt1), 2 (Pt2) and 3 (Pt3)} after obtaining written informed consent, and cultured in vitro for 7 days to induce their differentiation into macrophages, as previously reported (41). Pt1 carried a homozygous mutation at the splice-donor GT sequence of exon 2, resulting in a TT transition (c.177 +1 g > t) (14). In Pt2 and Pt3, a homozygous splice site mutation in intron 2 upstream of exon 3 was reported (c.178–10 t > a) (15). Cells were fixed in ice-cold 4% paraformaldehyde (Sigma-Aldrich), washed with PBS and permeabilized with 0.025% Triton X-100 (5 min). Then, samples were blocked in 5% BSA/PBS and incubated for 1 h with anti-MLC1 pAb (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA, sc-86740, dilution 1:200), or anti-EGFR pAb (1:200, Invitrogen) followed by Alexa 488-conjugated goat anti-rabbit secondary antibody for 1 h at RT. Wheat germ agglutinin conjugated to Texas Red (titer of 1:200 in 1% BSA/PBS, Life Technologies) was used as a plasma membrane marker. Nuclei were counterstained with 1 ng/ml Hoechst 33 342 (Life Technologies). Confocal microscopy imaging was performed by means of Olympus Fluoview FV1000 confocal microscope equipped with FV10-ASW version 2.0 software, and sequential confocal images were acquired using a PLAPON 63× oil immersion objective (1.42 NA) with a 1024 × 1024 format and scan speed 40 μs/pixel. The average intensity of EGF receptor fluorescence was calculated using FV10-ASW software, from cytometric measurements relative to total cell area as well as nuclear and cytoplasmic compartments, in nine digital images randomly selected and analyzed for each cell sample. Hundred to 120 cells were examined for each sample analyzed.

Quantitative analysis of cell proliferation by immunofluorescence and real-time RT-PCR

Cell proliferation was evaluated in U251 cells, both mock-infected (Ø) and overexpressing WT or mutated MLC1. Quantitative examination of the proportion of proliferating cells was performed by indirect IF with a pAb specific for the proliferation marker Ki67 (1:150, Spring Biosciences, Canada). To quantify total cells, we used 4,6-diamidino-2-phenylindole (DAPI) nuclear staining. Sixty to 80 cells were counted for each sample analyzed. For real-time RT-PCR, total RNA was isolated from cell pellets using RNeasy Micro kit (QIAGEN, Valencia, CA, USA), including on-column DNAse digestion, according to the manufacturer's instructions. RNA integrity and quality were checked on ethidium bromide containing 1% agarose gels in Tris-borate/EDTA buffer. For each sample, 1 μg of purified RNA, measured by Nanodrop (Thermo Fisher Scientific, Waltham, MA, USA), was reverse transcribed using high-capacity reverse transcription kit (Life Technologies) and then analyzed in triplicate by PCR, using the Applied Biosystems 7500 Real-Time PCR System, Gene Expression Master Mix and inventoried FAM-labeled gene expression assays (all from Life Technologies). The relative amounts of Ki67 (assay code: Hs01032443_m1) transcript were calculated using the comparative Ct (threshold cycle number at a cross-point between amplification plot and threshold) method and normalized to the internal GAPDH control (assay code: Hs99999905_m1).

Protein extract preparation and western blotting

U251 astrocytoma cell lines, C6 glioma cells and human fetal astrocytes were lysed and analyzed by WB as previously described (20,25). For WB of phosphorylated proteins, 0.1 mm sodium orthovanadate (Na3VO4) and phosphatase inhibitor cocktail (Roche Diagnostics, Mannheim, Germany) were added to the lysis buffer. Protein samples were subjected to SDS-PAGE using gradient (4–12%) pre-casted gels (Life Technologies), transferred to a nitrocellulose membrane, and immunoblotted ON at 4°C with the following Abs: anti-MLC1 pAb (1:1500, in-house generated), anti-actin mAb (1:2000, Santa Cruz Biotechnology), anti-caveolin-1 pAb (1:1000, Santa Cruz Biotechnology), anti-EAAT2 (1:2000, Santa Cruz Biotechnology), anti-EGFR mAb (1:1000, Abcam), anti-pAkt (Ser473) pAb, anti-Akt pAb, anti-pERK1/2 (Thr202/Tyr204) pAb, anti-ERK1/2 pAb and anti-pPLCγ1 (Tyr782) pAb (1:1000, Cell Signaling Technology, Danvers, MA) in PBS + 3% BSA. Membranes were then incubated with horseradish peroxidase-conjugated anti-mouse or anti-rabbit Ab (1:10 000; Thermo Scientific, Missouri, MO, USA) for 1 h at RT. Immunoreactive bands were visualized using an enhanced chemiluminescence reagent (Pierce, Thermo Fisher Scientific, Rockford, IL, USA), according to the manufacturer's instructions and exposed on X-ray films. Densitometric analyses of WB experiments were performed using NIH ImageJ software or Bio-Rad ChemiDoc XRS system.

Detergent-resistant microdomain (DMR/lipid rafts) preparation by sucrose gradients

Detergent-resistant microdomains (DRMs) from U251 astrocytoma cell lines overexpressing WT and mutated MLC1 were prepared as previously described (19,20). Samples were precipitated with acetone (1:4 v/v) and proteins analyzed by SDS-PAGE and WB.

Intracellular Ca2+ measurement by FURA-2

Intracellular Ca2+ measurement by FURA-2 was performed as previously described (24). Briefly, Fura-2-AM loading was achieved by exposing cell cultures to a solution containing 2.5 µM Fura-2-AM for 50 min at RT, then the culture was washed and after 10 min it was used for recording. The solution used for loading and recording had the following composition (mm): 140 NaCl, 5 KCl, 2.5 CaCl2, 1 MgCl2, 10 d-glucose and 10 HEPES/NaOH (RT, pH 7.4, 290 mOsm/l). Ca2+-free solutions were made by replacing Ca2+ with an equal amount of Mg2+ and by adding EGTA (0.5 mm). For the inhibition of the store-operated calcium channels, 1-(2-(4-methoxyphenyl)-2-(3-(4-methoxyphenyl)propoxy)ethyl-1H-imidazole hydrochloride (SKF96365) was used (10 µM). A perfusion system allowing the local change of the solution bathing the cells under study was used (Rapid Solution Changer 100; Bio-Logic, Claix, France). An inverted microscope (Axiovert 135, Zeiss, Germany) equipped with an objective 20X (0.75 NA) was utilized for fluorescence video imaging. Fura-2-loaded cells were exposed to the excitation wavelengths 340 and 380 nm by means of a monochromator (Polychrome II, Photonics, Germany), and the emission light at 510 nm was collected by a digital camera (PCO, Sensicam, Germany) and recorded on the hard disk of a PC computer (Dell, USA). Recording and analysis of the data were made possible by the use of the Imaging Workbench software package (INDEC Systems, CA, USA). For further data processing and presentation, the Origin 7.5 software package (Microcal Software, NJ, USA) was utilized. Ca2+ concentrations were expressed as the ratio between the emission at 340 and 380 nm. Traces represent the mean ± SEM of the signals recorded in single cells in representative experiments.

Electrophysiology

For electrophysiological measurements, U251 astrocytoma cells were used 3 days after plating in Petri dishes at 1.5 × 104 cells/ml. Macroscopic currents were recorded using the perforated patch method, as described in Fioretti (48). The external bathing solution consisted of the following (in mm): NaCl 106.5, KCl 5, CaCl2 2, MgCl2 5, MOPS 5, glucose 20 and Na-gluconate 30 at pH 7.25, and the intracellular solution was: K2SO4 57.5, KCl 55, MgCl2 5 and MOPS 10 at pH 7.2. Electrical access to the cytoplasm was achieved by adding amphotericin B (200 μM) to the pipette solution. Access resistances in the range of 10–20 MΩ were achieved within 10 min following seal formation. External solution was conditioned with octanol (1 mm) to block gap junctions and TEA (2 mm) to inhibit large-conductance calcium-activated K+ channels expressed in these cells (data not shown). Stock solutions of DCEBIO, CTL (SIGMA) and ionomycin (TOCRIS) were made by dissolving these drugs in DMSO at concentrations of 100, 10 and 0.5 mm, respectively. Experiments were carried out at RT (18–22°C).

Statistical analysis

Statistical differences were calculated using Student's t-test to compare two groups, and data are presented as mean ± standard error of the mean (SEM). A P-value of <0.05 was set as significant.

Supplementary Material

Supplementary Material is available at HMG online.

Funding

This work was supported by the Telethon Italy (Grant Nos GEP14134 and GGP1118/3 to E.A.) and the ELA Foundation (ELA 2012-0021F2 to A.L.).

Acknowledgements

We thank Dr Francesca Aloisi for critical reading of the manuscript and stimulating comments. We also thank Gianfranco Macchia for the technical assistance in the preparation of protein rafts from U251 cells.

Conflict of Interest statement. None declared.

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Author notes

These authors equally contributed to this work.