Astroglia are both a target for and a source of cytokines. The latter are soluble mediators that may serve as communication signals between astroglia and neighboring neural cells (Benvensite,1992; Merrill,1992; Eddleston and Mucke,1993). Astroglia exhibit remarkable cellular responses following a variety of insults to the central nervous system (CNS). The most striking response is the induction of astrogliosis. Astrogliosis is characterized by astrocyte proliferation, hypertrophy, and increases in the synthesis of glial fibrillary acidic protein (GFAP) (Bignami and Dahl,1995). The mechanisms by which astrogliosis is induced are poorly understood. It has been demonstrated by previous studies that the expression of GFAP is regulated by a wide variety of hormones, cytokines, and growth factors (Morrison et al.,1985; Laping et al.,1994; Gomes et al.,1999).
Interleukin-1β (IL-1β), a multifunctional cytokine, is secreted by activated astroglia and microglia in brain, where it exerts a diverse range of activities on immune function and coordination of many aspects of the acute phase response to trauma and infection (Fontana et al.,1982; Giulian et al.,1986; John et al.,2005). In a number of different inflammatory and degenerative conditions of the CNS, an increase in the expression of IL-1β has been extensively documented (Hopkins and Rothwell,1995). Previous studies have suggested that IL-1 acts as an astroglia growth factor during brain development, and it may play an important role in astrogliosis following injury to the CNS (Lachman et al.,1987; Giulian et al.,1988a,1988b). Following exposure to cytokines (IL-1β, TNF-α, and IFN-γ), astroglia undergo functional changes that involve both gene expression and protein synthesis (Benvensite,1992; Merrill,1992) and secretion of the proinflammatory cytokine IL-6 (Frei et al,1989; Benveniste et al.,1990; Aloisi et al.,1992,1995).
IL-1β induces morphological changes in human fetal astroglia, which reflect on their functional status. Previous studies have shown that IL-1β downregulates the expression of GFAP mRNA in rat astroglia, however, its effect on the protein level is not clear (Oh et al.,1993). The effect of IL-1β on the organization and expression of cytoplasmic β-actin isoform and α-sm actin isoform, which is expressed by astroglia is not known (Abd-El-Basset and Fedoroff,1991; Lecain et al.,1991). It is known that different actin isoforms may have different functions that translate into overall functional capabilities of cells (Khaitina,2001; Chaponnier and Gabbiani,2004; Reisler and Egelman,2007). The neither functional significance of the presence of α-sm actin isoform in astroglia is not yet known nor is the mechanism that regulates the expression of different actin isoforms in astroglia.
In this study, we examine the effect of mouse recombinant IL-1β on the morphology of astroglia and on the organization and expression of GFAP, cytoplasmic β-actin and α-sm actin isoforms in cultured mouse astroglia during short-term (2 days) and long-term (range, 4–8 days) treatment.
MATERIALS AND METHODS
The cerebral hemispheres of newborn Balb/c mice were isolated aseptically and the meninges were removed. The neopallia were dissected out and then forced through a sterile 75-μm Nitex mesh. The cells were suspended in a modified Eagle's minimum essential medium containing 5% horse serum. For culturing, 5 × 104 nigrosine-excluding cells were plated in 60-mm Nunc Petri dishes containing 11 × 22 mm glass coverslips in a total volume of 3-mL growth medium. Aliquots of 5 × 106 nigrosine-excluding cells were plated in 75 cm2 Nunc culture flasks. The cultures were incubated at 37°C in a humidified atmosphere of 5% CO2 in air. After 3 days of incubation, the growth medium was removed, the cell debris and nonattached cells were washed off, and fresh medium was added. The cultures were then incubated for 20 days with a medium change every 2–3 days. After 10–12 days, some culture plates and flasks were incubated for 1, 2, 4, and 8 days in medium to which 200 international units (U)/mL of IL-1β (Sigma, St. Louis, MO) had been added and in which the horse serum had been lowered to 1%. The new medium was also changed every 2–3 days. In controls, the cells were incubated with medium containing 1% horse serum (without IL-1β).
The cells on coverslips after treatment with 200 U/mL of IL-1β for 2, 4, or 8 days, along with control cells, were fixed for 10 min with 3.7% formaldehyde in phosphate-buffered saline (PBS) at room temperature. The cells were then extracted for 2 min with −20°C methanol. The fixed cells were treated first with one of the following mouse monoclonal primary antibodies to: GFAP, α-sm actin isoform, or β-actin isoform (Sigma, St. Louis, MO) diluted 1:200 in PBS, for 45 min. The cells were then treated with the secondary antibody; fluorescein-conjugated F(ab)2 fragment of donkey anti-mouse IgG (Jackson ImmunoResearch Laboratories, Baltimore Pike) diluted 1:100 in PBS for 45 min. For double labeling of both α-sm actin and β-actin isoforms, the cells were treated first with monoclonal primary antibody to β-actin isoform for 45 min followed by rhodamine-conjugated F(ab)2 fragment of donkey anti-mouse IgG (Jackson ImmunoResearch Laboratories, Baltimore Pike) diluted 1:100 for another 45 min. The cells were then treated with fluorescein-conjugated monoclonal antibody to α-sm actin isoform (Sigma, St. Louis, MO) diluted 1:200 for 45 min. Some cells after fixation in formaldehyde were extracted with 0.1% Triton X-100 (optimal for phalloidin labeling) and then treated with monoclonal antibody to α-sm actin isoform. The cells were then treated with fluorescein-conjugated F(ab)2 fragment of donkey anti-mouse IgG for 45 min then double labeled with rhodamine phalloidin (Molecular Probe, Eugene, OR) diluted 1:50 in PBS for another 45 min.
For controls, the primary antibodies were omitted and parallel single labeling with each antibody was done. All the procedures were carried out at room temperature. After application of each antibody, the preparations were washed three times for 5 min each in PBS. The cells were mounted in 50% glycerol in PBS, pH 7.8, and examined in Olympus photomicroscope equipped with mercury vapor lamp, epifluorescence optics, and appropriate interference filters.
Fluorescent Microplate Assays
Culture flasks with monolayer of astrocytes were trypsinized using 0.25% trypsin for 15 min, centrifuged, and resuspended in culture medium containing 5% horse serum. The same number of cells (1 × 105) were plated in each well of culture plates and incubated for 2 days at 37°C in 5% CO2. The cells were incubated for another 2, 4, or 8 days in medium with 1% horse serum to which 200 U/mL of IL-1β was added along with controls without IL-1β. The medium was changed every 2 days. The cells were fixed in 3.7% formaldehyde for 10 min and then extracted for 2 min with −20°C methanol. For quantification of GFAP, α-sm actin, and β-actin, the cells were treated with monoclonal antibody to either GFAP, α-sm actin, or β-actin diluted 1:200 in PBS for 45 min. The cells were then treated with fluorescein-conjugated F(ab)2 fragment of donkey anti-mouse IgG diluted 1:100 in PBS for another 45 min. For controls, the primary antibodies were omitted. After application of each antibody, the preparations were washed three times for 5 min each in PBS using a multichannel pipette. Finally, 100 μL PBS was added to each well, and the plates were read in a microplate fluorescence reader (Skatorn) FL 600 with software KC4 version 2.5 to quantify the fluorescence in each well. The excitation and emission wavelengths were set at 485/20 and 530/25 nm, respectively, with the sensitivity specified at 100. The fluorescence reading was expressed as fluorescence units. The percentage of fluorescence intensity was calculated with normalization of controls to 100%. This experiment was repeated three times.
Polyacrylamide Gel Electrophoresis and Immunoblotting
Cells grown in the flasks were washed three times in PBS, scraped off with a rubber policeman, and collected in PBS. The cells were immediately sonicated for 1 min over ice, and the protein concentration was determined according to the method of Bradford (1976). The sonicated cells were dissolved in 10% sodium dodecyl sulphate (SDS)-containing sample buffer and boiled for 3 min. The same amount of protein (20 μg/well) from each sample was analyzed electrophoretically in 10% SDS-polyacrylamide gel electrophoresis (PAGE) gel according to the method of Laemmli (1970).
The proteins in the gel were transferred to nitrocellulose sheets by a modification of the method of Towbin et al. (1979). After transfer, the nitrocellulose sheets were incubated for 1 h with 3% bovine serum albumin (BSA) in PBS. The nitrocellulose sheets were then incubated overnight at room temperature with mouse monoclonal antibodies either to β-actin isoform or to α-sm actin isoform, diluted 1:500 in PBS containing 3% BSA. The sheets were then incubated for 4 h with affinity-purified goat anti-mouse IgG conjugated to horse-radish peroxidase diluted 1:1000 in PBS containing 3% BSA. Some nitrocellulose sheets were incubated with rabbit polyclonal antibody to GFAP, diluted 1:500 in PBS containing 3% BSA, and then followed by affinity-purified goat anti-rabbit IgG conjugated to horse-radish peroxidase. The color was developed using freshly prepared 0.05% 4-chlor-1-naphthol and 0.015% H2O2 in PBS. The reaction was stopped by washing in tap water. All the chemicals and materials for electrophoresis and immunoblotting were purchased from BioRad (Missisauga, Ont, Canada). Immunoblots of five independent experiments were scanned using the Snapescan 1212 scanner and Adobe Photoshop 5.0 program. The relative levels of GFAP, α-sm actin, and β-actin expression were determined by analyzing the pixel intensity of the bands using an imaging analysis program (Image J, version 1.04b, Wayne Rasband, NIH). The percentage of the protein expression was calculated in the following manner. The average background protein levels in the lane, excluding the bands, were first subtracted from both the control and treatment bands. The percentage of the decrease or increase in intensity (I) of IL-1β treated cells (T) compared with the control (C) was calculated as follows:
Experiments were performed at least three times on different cell preparations. The data of the fluorescent intensity of the staining for GFAP, α-sm actin, and β-actin in control and IL-1β-treated astrocytes were analysis with two-tailed student's t-test. P-values < 0.05 were considered statistically significant.
When astroglia were treated with 200U of IL-1β with serum lowered to 1%, the number of astroglia with elongated processes increased compared with the control (Fig. 1).
To determine the effect of IL-1β on the organization and expression of different cytoskeletal proteins, immunostaining of astroglia treated with 200 U/mL IL-1β for 1, 2, 4, and 8 days using antibodies to GFAP, to α-sm actin, and to β-actin was preformed. The results showed the following:
The majority of the cells (about 90%) growing in the culture for 10–12 days were positive for GFAP. After treatment of astroglia for 1 and 2 days, the immunofluorescent staining for GFAP became more intense than that of the control and the majority of these cells had elongated processes (Fig. 1). Astroglia treated for 4 and 8 days became less bright and GFAP could not be detected in some cells (about 20%; not shown).
α-sm actin isoform
Astroglia treated for 1 and 2 days did not show remarkable change in the intensity or organization of α -sm actin isoform compared with the control. The number of cells expressing α-sm actin isoform decreased to 77% and 63% of control levels in 4 and 8 days treated astroglia, respectively. Double labeling of 4-day treated astroglia using antibody to α-sm actin isoform and phalloidin, which binds to F-actin, showed that all the cells had F-actin, however, some cells downregulated their α-sm actin isoform (Fig. 2).
Astroglia treated for 1, 2, 4, and 8 days showed an increase in the intensity of β-actin isoform staining (not shown). When 8-day treated astroglia were double labeled for β-actin and α-sm actin, the cells showed intense staining for β-actin isoform and downregulation of their α-sm actin isoform (Fig. 3).
Fluorescent Microplate Assay
Fluorescence intensity of the GFAP staining significantly increased to 122% ± 2% of the control level in 2 days treated astroglia (P < 0.001, Student's t-test). It then progressively significantly decreased to 88% ± 2% and 71% ± 4% of control level in 4 and 8 days treated astroglia, respectively (P < 0.001, Student's t-test; Fig. 4A).
α-sm actin isoform
No significant changes were observed in the fluorescence intensity of α-sm actin isoform of 2 days treated astroglia (P = 0.83, Student's t-test). Fluorescent intensity significantly decreased progressively to 78% ± 2% and 64% ± 4% of the control level in 4 and 8 days treated cells, respectively (Fig. 4B; P < 0.001, Student's t-test).
After treatment of astroglia for 2 days, fluorescence intensity of β-actin isoform significantly increased to 126% ± 4% of the control level (P < 0.001, Student's t-test), then started to significantly change to 124% ± 4% (P < 0.001, student's t-test), and 114% ± 4% (P < 0.05, Student's t-test) of the control level in 4 and 8 days treated astroglia, respectively (Fig. 4C).
To determine the effect of IL-1β on the expression of different actin isoforms and GFAP in astroglia, proteins from 2 and 8 days IL-1β-treated astroglia and control astroglia were separated by SDS-PAGE, transferred to nitrocellulose sheets and treated with either antibody to α-sm actin isoform, to β-actin isoform, or to GFAP (Fig. 5). Quantification of the western blots of five independent experiments showed that the amount of GFAP in 2-day treated astroglia increased to about 125% of control, whereas, the amount of GFAP in 8-day treated astroglia decreased to 35% of control. With short-term treatment of astroglia (2 days), the amount of α-sm actin isoform did not show significant change, however, with long-term treatment (8 days), the α-sm actin isoform was decreased to about 34% of control. The amount of β-actin isoform in 2-day treated astroglia increased to 40% of control, whereas in 8-day treated astroglia it was 10%.
This study examines the effects of cytokine IL-1β on morphological changes and expression of cytoskeletal proteins in mouse astroglia. IL-1β has transient effects on astrogliosis as assayed by GFAP expression. This is manifested by an increase in the expression of GFAP for a short period followed by downregulation of GFAP expression after a long period of treatment. In addition, IL-1β has a differential effect on the expression of actin isoforms in astroglia, with upregulation of the expression of the β-actin isoform and downregulation of the expression of the α-sm actin isoform.
Our study shows that IL-1β induces morphological changes in astroglia grown in reduced serum medium. The flat polyhedral cells acquired more elongated processes. These data are in agreement with the previous study on the effect of IL-1β on the morphology of human fetal astroglia (Liu et al.,1994). The morphological changes in the shape of astroglia in tissue culture may result from interactions with many factors including activators and inhibitors of cAMP-dependent kinase, protein kinase C, and tyrosine kinase-dependent pathways (Shafit-Zagardo et al.,1988; Shirakawa et al.,1988; Abe and Saito,1999; Pascale et al.,2004).
Our finding shows that IL-1β upregulates the expression of GFAP in mouse astroglia for 2 days, followed by downregulation of GFAP after 4 days of treatment. However, previous studies have shown that IL-1β does not significantly affect the level of GFAP in rat astroglia, with previous groups showing slight increases after 2 days of stimulation, followed by a slight decrease (Oh et al.,1993). Significant changes in the GFAP level were also not observed after treatment of human fetal astroglia for 1–2 days with IL-1β, however by days 5 and 7, the level of GFAP decreased (Liu et al.,1994). In situ studies show an increase in the number and intensity of GFAP staining astroglia around the site of IL-1β injection in rat brain (Giulian et al.,1988b). In addition, direct intracerbrovascular infusion of IL-6 as well as IL-1β-induced IL-6 release into CSF are strongly associated with GFAP expression in the cerebral cortex and hippocampus. Furthermore, blockage of IL-1β-induced IL-6 release by IL-10 and propranolol leads to a decrease of GFAP expression (Woiciechowsky et al.,2004). The expression of GFAP is regulated by a wide variety of hormones and cytokines and is altered in various pathological conditions (Chiu and Goldman,1985; Morrison et al.,1985; Shirakawa et al.,1988; Pawlinski et al.,2000). In brain injury, GFAP expression may be upregulated rapidly and high levels of GFAP are characteristic of astrocytic scar formation in various pathological conditions (Bignami and Dahl,1976;1995; Balasingam et al.,1994). Activated microglia by damaged neurons release cytokines and other products that can influence the development of astrogliosis (Zhang et al.,2010). The proliferation of astroglia (Giulian and Lachman,1985) and the increase in the level of GFAP for a short period in response to IL-1β stimulation may be important in limiting the site of inflammation and supports the previous suggestion that IL-1β is important in astrogliosis (Giulian et al.,1988b). However, IL-1β is not enough to upregulate the expression of GFAP for a long time. This suggests that other factors in addition to IL-1β are responsible for the astrogliosis. These data emphasis the role that reactive astrocytes play in wound healing and restoring CNS function at the site of injury and indicating the importance of preserving astrocyte populations (Faulkner et al.,2004; John et al.,2005).
Astrocytes are critical players in the regulation of cerebral blood vessels diameter and they appear capable of eliciting changes in the vessel diameters in both directions (Gorden et al.,2007) depending on the different state of brain metabolic activities (Gorden et al., 2008). Astrocytes foot processes can signal the vascular smooth muscles by different pathways (Koehler et al.,2006,2009) and adjust vascular tone to a range where dilator responses are optimal (Blanco et al.2008). In addition, astrocytes contain contactile proteins, actin and actin-binding proteins (Abd-El-Basset et al.,1991; Abd-El-Basset and Fedoroff,1994), which are upregulated in reactive astrocytes (Abd-El-Basset and Fedoroff,1997). Modulation of the expression of actin isoforms by IL-1β may contribute to changes in the regulation of the blood vessels diameters and blood flow inside the CNS and emphasize the role of astrocytes in dynamic signaling within the neurovascular units. Important questions remain on the mechanism of this signaling and how this signaling is integrated with other pathways in health and disease.
Our finding reveals that IL-1β has a differential effect on the expression of α-sm-actin and β-actin isoforms. With short-term treatment, it upregulates the expression of β-actin and has no remarkable effect on the expression α-sm actin. Long-term treatment with IL-1β decreases the amount of α-sm actin expressed by astroglia and the number of cells expressing α-sm actin. The amount of β-actin increases above the control level, however, the amount of the increases is less than that of the increases of 2 days treatment. A previous study has also shown a differential effect of dBcAMP on the expression of actin isoforms in astroglia (Abd-El-Basset,2000). Previous and present studies suggest that the expression of actin isoforms in astroglia has different regulatory mechanisms, which implicate differences in functional activities in astroglia. In addition, IL-1β upreguates the expression of actin and β-actin isoform in microglia (Abd-El-Basset et al.,2004).
In conclusion, IL-1β may orient astrocyte function in a proinflammatory direction and at the same time may upregulate some of the cytoskeletal proteins, which in turn facilitate the ability of astroglia to limit the noxious effect of an inflammatory reaction and to promote cell survival and tissue repair processes. These finding may have important diagnostic and therapeutic applications to limit noxious effects of neuronal inflammation.
This work was supported by Kuwait University Research Adminstration grant # MA037. The authors would like to thank Dr. Ananth Lakshmi KVV and Mrs. Jeena Prashanth for technical assistance.