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Molecular mechanisms underlying the role of statins in the induction of brain plasticity and subsequent improvement of neurologic outcome after treatment of stroke have not been adequately investigated. Here, we use both in vivo and in vitro studies to investigate the potential roles of two prominent factors, vascular endothelial growth factor (VEGF) and brain-derived neurotrophic factor (BDNF), in mediating brain plasticity after treatment of stroke with atorvastatin. Treatment of stroke in adult mice with atorvastatin daily for 14 days, starting at 24 hours after MCAO, shows significant improvement in functional recovery compared with control animals. Atorvastatin increases VEGF, VEGFR2 and BDNF expression in the ischemic border. Numbers of migrating neurons, developmental neurons and synaptophysin-positive cells as well as indices of angiogenesis were significantly increased in the atorvastatin treatment group, compared with controls. In addition, atorvastatin significantly increased brain subventricular zone (SVZ) explant cell migration in vitro. Anti-BDNF antibody significantly inhibited atorvastatin-induced SVZ explant cell migration, indicating a prominent role for BDNF in progenitor cell migration. Mouse brain endothelial cell culture expression of BDNF and VEGFR2 was significantly increased in atorvastatin-treated cells compared with control cells. Inhibition of VEGFR2 significantly decreased expression of BDNF in brain endothelial cells. These data indicate that atorvastatin promotes angiogenesis, brain plasticity and enhances functional recovery after stroke. In addition, VEGF, VEGFR2 and BDNF likely contribute to these restorative processes.
We have previously demonstrated that treatment of stroke in the rat with HMG-CoA reductase inhibitors (statins), atorvastatin and simvastatin, with treatment initiated 24 hours after onset of stroke, significantly improves neurologic outcome compared with control animals (Chen et al, 2003b). These agents induce angiogenesis, neurogenesis and synaptogenesis in the ischemic brain, and this brain remodeling likely contributes to the functional recovery (Chen et al, 2003b). However, the molecular triggers for the statin-induced brain plasticity are not known. In addition, our prior observation of statin-induced angiogenesis, neurogenesis and synaptogenesis raises the question of how these aspects of brain plasticity are coupled, and if there are specific molecular factors that can integrate these remodeling events in the brain.
Vascular endothelial growth factor (VEGF) and brain-derived neurotrophic factor (BDNF) are two important neurotrophic factors that have multiple effects on sustaining and evoking elements of brain plasticity (Jin et al, 2002; Sun et al, 2003). Vascular endothelial growth factor is an angiogenic agent, which promotes neurogenesis and stem-progenitor cell migration (Jin et al, 2002; Sun et al, 2003). Likewise, BDNF regulates neuronal survival, cell migration, and synaptic function (Aguado et al, 2003; Gorski et al, 2003). Statins increase VEGF in endothelial and osteoblast cells (Chen et al, 2003a; Maeda et al, 2003). Both VEGF and BDNF increase in ischemic brain with other neurorestorative treatments of stroke, such as with bone marrow stromal cells (Chen et al, 2002; Matsuno et al, 2004). It is therefore reasonable to propose that VEGF and BDNF might be upregulated in the brain after treatment with a statin, and may orchestrate brain plasticity.
In this study, we therefore examined the effects of a widely employed hydrophilic statin, atorvastatin, on angiogenesis, neuronal plasticity and neuronal migration in a mouse model of permanent middle cerebral artery occlusion (MCAO). We show that atorvastatin increases cerebral expression of VEGF and its receptor VEGFR2 induces endothelial cell proliferation and angiogenesis, and promotes production of neurotrophic factors, BDNF and VEGF, which may act in concert to induce brain plasticity and to foster functional recovery after stroke in mice.
Adult male C57BL/6J mice (age 2 to 3 months, weight 24 to 28 g) were purchased from Charles River (Wilmington, MA, USA). Mice were anesthetized with halothane. Permanent right MCAO was induced by advancing a 6-0 surgical nylon suture (8.0 to 9.0mm determined by body weight) with an expanded (heated) tip from the external carotid artery into the lumen of the internal carotid artery to block the origin of the MCA (Mao et al, 1999). Atorvastatin was dissolved in methanol (20mg atorvastatin dissolved in 20mL saline with 10 μL methanol) and injected subcutaneously (s.c.) at 24 hours after MCAO. An atorvastatin dose of 10 mg/kg was selected based on pretreatment studies of stroke in mice (Gertz et al, 2003; Laufs et al, 2000).
After MCAO, mice were randomly divided into the following groups: Group 1, saline (0.25 mL) daily for 14 days as a control group (n=12); Group 2, atorvastatin (10 mg/kg) daily for 14 days (n=11); Group 3, saline (0.25 mL) daily for 7 days as a control group (n=10); Group 4, atorvastatin (10 mg/kg) daily for 7 days (n=10). To identify newly formed DNA in ischemic brain, mice received injections of bromodeoxyuridine (BrdU, Sigma Chemical, 100 mg/kg in 0.007N NaOH physiologic saline), intraperitoneally (i.p.) starting 1 day after MCAO and daily thereafter until the mice were killed. Functional tests were performed on Groups 1 and 2. Groups 1 and 2 were killed at 14 days after MCAO for immunostaining; mice in Groups 3 and 4 were killed at 7 days after MCAO and brain tissue extracts were obtained for enzyme-linked immunosorbent (ELISA) assay. In addition, physiologic parameters (heart rate, blood pressure, blood gases) were also measured before killing in Group 3 and 4 mice treated with or without atorvastatin for 7 days after stroke (n=4/group). The left femoral artery in animals was cannulated for heart rate and arterial blood pressure (Protocol Systems, Inc.) and blood gas determination (Radiometer Copenhagen, Inc.). Arterial blood samples were analyzed for pH, arterial oxygen pressure and partial pressure of carbon dioxide by using a blood gas/pH analyzer (Radiometer Copenhagen, Inc.).
To test whether atorvastatin promotes neurologic recovery after stroke, an array of behavioral tests were performed before MCAO, and at 1, 7 and 14 days after MCAO by an investigator who was masked to the experimental groups.
Neurologic function was graded on a scale of 0 to 14 (normal score 0; maximal deficit score 14, see Table 1; Li et al, 2000). mNSS is a composite of motor, reflex and balance tests. In the severity scores of injury, one score point is awarded for the inability to perform the test or for the lack of a tested reflex; thus, the higher the score, the more severe is the injury.
For locomotor assessment, mice were tested for placement dysfunction of forelimbs with the modified foot-fault test (Hernandez and Schallert, 1988). The total number of steps (movement of each forelimb) that the mouse used to cross the grid was counted, and the total number of foot-faults for left forelimb was recorded. The percentage of foot-fault of the left paw to the total number of steps is presented.
Mice were placed between two boards with dimensions 30×20×1cm3 for each in the home cage (Zhang et al, 2002a). The nonischemic animals turn back from either left or right, randomly. The ischemic animals preferentially turn toward the impaired side. The number of turns taken on each side was recorded from 10 trials for each test.
Mice subjected to MCAO followed by treatment with saline or atorvastatin daily for 14 days (n=9) were killed at day 14 after MCAO. Brains were fixed by transcardial perfusion with saline, followed by perfusion and immersion in 4% paraformaldehyde, and the brains were embedded in paraffin. Coronal sections of tissue were processed and stained with hematoxylin and eosin (H&E) for calculation of volume of cerebral infarction (Swanson et al, 1990). The indirect lesion area, in which the intact area of the ipsilateral hemisphere was subtracted from area of the contralateral hemisphere, was calculated (Swanson et al, 1990). Lesion volume is presented as a volume percentage of the lesion compared with the contralateral hemisphere.
To measure whether atorvastatin treatment promotes trophic factor expression, angiogenesis and brain plasticity, BrdU, von Willebrand Factor (vWF), BDNF, doublecortin (DCX), tubulin isotype III (TUJ-1) and synaptophysin immunostaining were performed. Briefly, two standard paraffin blocks were obtained from the center of the lesion, corresponding to coronal coordinates for bregma −1 to 0mm and from 0 to +1mm. A series of 6-μm-thick sections were cut from the two blocks. Every 10th coronal section (6 μm thick) for a total of 5 sections from each block was used for immunohistochemical staining. For immunohistochemical analysis a mouse monoclonal antibody (mAb) against BrdU, (1:100, Boehringer Mannheim, Indianapolis, IN, USA), vWF (goat polyclonal IgG antibody, 1:200 dilution, Santa Cruz), BDNF (C-9, 1:200 dilution Santa Cruz); TUJ1, an early neuron marker, (Monoclonal Anti-B-Tubulin Isotype III, 1:400 dilution, Sigma) and doublecortin (DCX), a protein expressed in migrating neurons (C-18, goat polyclonal IgG antibody, 1:200 dilution, Santa Cruz), and synaptophysin (monoclonal antibody, clone SY 38, 1:40 dilution; Boehringer Mannheim Biochemica, Indianapolis, IN) were used. Control experiments consisted of staining brain coronal tissue sections as outlined above, but omitted the primary antibodies, as previously described (Li et al, 1998).
After stroke, BrdU-reactive cells are not only expressed in the endothelial cells but also in the astrocytes, and pericytes around vessels. To specifically identify BrdU-reactive cells colocalized with endothelial cells, double immunofluorescence staining of BrdU with vWF, a specific marker for endothelial cells, was performed. To visualize BDNF-reactive cell colocalization with the neuron marker MAP2, double immunofluorescence of BDNF with MAP2 was performed. FITC (Calbiochem) and cyanine-5.18 (CY5, Jackson Immunoresearch) were used for double-label immunoreactivity. Each coronal section was first treated with the primary anti-BrdU or anti-BDNF antibody with FITC, and then followed by vWF or microtubule-associated protein 2 (MAP2, monoclonal mouse anti-MAP2 antibody, Chemicon, Temecula, CA, USA) with Cy5 staining. Control experiments consisted of staining brain coronal tissue sections as outlined above, but omitted the primary antibodies, as previously described (Li et al, 1998).
For semiquantitative measurements of BDNF, TUJ1, DCX and synaptophysin densities, five slides from each block, with each slide containing 8 fields consisting of cortex and stratum from the ischemic border area as shown in (Figure 1) were digitized under a ×20 objective (Olympus BX40) using a 3-CCD color video camera (Sony DXC-970MD) interfaced with an MCID image analysis system (Imaging Research, St. Catharines, Canada). The ischemic border zone is defined as the area surrounding the lesion, which morphologically differs from the surrounding normal tissue (Nedergaard et al, 1987). The digitalized images were then contrast-enhanced to clearly differentiate positivity from background, and a thresholding procedure was established to determine the proportion of immunoreactive area within each fixed field of view (Calza et al, 2001; Chen et al, 2003b). The quantification was not analyzed stereologically. Data are presented as a percentage of area, in which the TUJ1, DCX, synaptophysin immunopositive areas in each field were divided by the total areas in the field (628×480 μm2) (Calza et al, 2001; Chen et al, 2003b). Brain-derived neurotrophic factor (BDNF) quantitative data are presented as the total of number of BDNF immunoreactive cells within each field.
Bromodeoxyuridine-immunostained sections were digitized using a ×40 objective (Olympus BX40) via the MCID computer imaging analysis system. The number of endothelial and BrdU-immunoreactive cells (nuclei on the lumen of vessel) within a total of 10 enlarged and thin-walled vessels located in the ischemic border area (as shown in Figure 1) were counted in each section (Chen et al, 2003b; Wang et al, 2004; Zhang et al, 2003). Data are presented as the percentage of number of the BrdU-immunoreactive cells within vessel/total endothelial cell number.
Enlarged and thin-walled vessels, termed ‘mother’ vessels, are formed under conditions of cerebral ischemic angiogenesis (Zhang et al, 2002b). For measurement of vascular perimeters, each vWF immunostained coronal section was digitized using a ×20 objective via the MCID computer imaging analysis system. The total perimeter of 10 enlarged and thin-walled vessels in the ischemic border area (as shown in Figure 1) was measured in each referenced coronal section, as previously described (Chen et al, 2003b; Wang et al, 2004; Zhang et al, 2003, 2002b), using MCID computer imaging analysis system (Length Trace function). The total perimeter of 10 enlarged and thin-walled vessels in the ischemic border is provided.
To test whether atorvastatin promotes the expression of VEGF and VEGFR2, ELISA analysis was performed on the ischemic brain tissue extracts (Jiang et al, 1997). Mice were subjected to permanent MCAO and treated daily with saline or atorvastatin (10 mg/kg) starting at 24 hours after stroke for 7 days. Mice were killed at day 7 after stroke. Brain extracts from control MCAO and atorvastatin-treated animals were obtained from the ischemic border (bregma −1 to 1mm, border region encompassing the ischemic core) and contralateral hemisphere. Brain tissues were dissected on ice and wet weight was rapidly measured. In 150 mg/ml of tissue was homogenized in DMEM and centrifuged for 10 mins at 10,000 g at 4°C. The brain extracts were then divided into 200 μL triplicate samples. Vascular endothelial growth factor and VEGFR2 ELISA kit (Cell Signaling Technology) were used to determine the VEGF and VEGFR2 levels.
To measure whether atorvastatin induces neural cell migration, SVZ explant cultures were prepared from adult male rats subjected to MCAO for 7 days (Dutton and Bartlett, 2000; Lois and Alvarez-Buylla, 1993). The methods for extracting and employing SVZ explants from rats have been widely used and are well developed (Katakowski et al, 2003). The SVZ was dissected from the ipsilateral SVZ of the rat brain. Tissue was minced with scalpels into pieces of ~0.1mm in each dimension. Explants were cultured within Matrigel (BD Biosciences) in wells with 500 μL of Neuralbasal-A Medium containing 2% B27 supplement (Invitrogen). The cultured SVZ explants were treated in the absence or presence of: (a) atorvastatin (1 μmol/L); (b) BDNF (100 ng/mL, Sigma); (c) anti-BDNF neutralizing antibody (100 ng/mL, Promega); (d) atorvastatin (1 μmol/L) and anti-BDNF neutralizing antibody (100 ng/mL) for 7 days. Cell migration from the SVZ explant was measured using a phase contrast microscope and photographed at ×4 magnification with a digital camera. The average linear distance of cell migration from the edge of the SVZ explant was captured and measured at day 7 using the MCID software. This average distance was assessed in each explant culture.
To measure whether atorvastatin promotes endothelial cell expression of VEGFR2 and BDNF, mouse brain derived microvessel endothelial cells (MBECs) (1.5×104 cells) were cultured in an eight-well chamber in the absence (control) or presence of: (a) atorvastatin (1 μmol/L); (b) neutralized anti-VEGFR2 antibody (DC101, 30 μg/ml); and (c) atorvastatin (1 μmol/L) with DC101 (30 μg/ml) for 24 hours. VEGFR2 (1:100 dilution, Santa Cruz, USA) and BDNF immunohistochemistry staining was performed on slides containing cultured endothelial cells. DAPI was used as a nuclear counterstain. All assays were performed in triplicate. VEGFR2- and BDNF-positive cells were counted in five randomly selected microscopic fields under ×20 objects. The percentage of VEGFR2 or BDNF immunoreactive cells within the total number of DAPI positive cells was presented.
Data were evaluated for normality. Data transformation or nonparametric analysis approach would be considered if data were not normal. As a result, ranked data were used for behavior test scores. Analysis of variance and covariance (ANCOVA) was used to study treatment and time effect on functional recovery. Analysis began testing for treatment by time interaction at the 0.05 significance level, followed by testing for a treatment effect if no interaction was detected, or testing the treatment effect at each time if an interaction was detected. A significant interaction (P<0.05) indicates that the effects of the treatment of atorvastatin on behavior test results depend on time. We also investigated the atorvastatin effects on lesion volume, vWF, BDNF, TUJ1, DCX, BrdU and synaptophysin expression at 14 days after MCAO, compared with the controls using two-sample t-test at the significance level of 0.05. Mean±s.d. by treatment groups are presented as data illustration.
Functional tests showed balanced neurologic deficits before treatment between control and atorvastatin-treated groups. Mice treated with atorvastatin showed significant (P<0.05, n=9/group) improvement of functional recovery on mNSS, foot-fault and corner tests at days 7 and 14 compared with saline-treated mice (Figure 2). The mortality rate of atorvastatin-treated mice was two of 11 mice, and three of 12 MCAO control mice. No significant differences of ischemic lesion volumes were detected between atorvastatin-treated (16.7±8.7%) and saline-treated (19.0±10.7%) mice. These results suggest that atorvastatin promotes functional recovery after stroke in mice, while it has no effect on reducing the lesion volume. In addition, physiologic parameters (heart rate, blood pressure and blood gases) do not show significant difference after ischemia in the atorvastatin-treated group (HR: 562 ±43/min; BP: 68.8±3.0mm Hg; pH: 7.31±0.04; PaO2: 143.0±6.4mm Hg; PaCO2: 41.3±1.9mm Hg) compared with MCAO control animals (HR: 568± 56/min; BP: 71.3±3.6mm Hg; pH: 7.35±0.06; PaO2: 139.5±10.6mm Hg; PaCO2: 43.5±3.1mm Hg).
Treatment with atorvastatin significantly increased BrdU-reactive endothelial cell numbers (B and C) and vascular perimeter (E and F) in the ischemic border, as compared with the saline-treated animals (A, D). Bromodeoxyuridine with vWF double immunostaining shows that BrdU-positive endothelial cells (G and I) localize in the vessel (H and I). These data indicate that atorvastatin enhances angiogenesis in the ischemic brain. Treatment of stroke with atorvastatin significantly increased BDNF (K, arrow) immunoreactivity cell density in the ischemic border, as compared with the control mice (J, arrow). Double immunostaining showed that BDNF immunoreactive cells were colocalized with MAP2 immunoreactive cells in the cortex (M to O). Brain-derived neurotrophic factor was also detected in the endothelial cells (P, arrow). These findings suggest that atorvastatin promotes endothelial cell proliferation and angiogenesis. In addition, atorvastatin promotes BDNF expression. Brain-derived neurotrophic factor is expressed in neurons and endothelial cells.
Vascular endothelial growth factor and VEGFR2 were measured by ELISA method in the ischemic brain extract. Vascular endothelial growth factor and VEGFR2 were significantly (P<0.05) increased in the ischemic border in the atorvastatin-treated group (VEGF: 30.6±9.3 pg/g; VEGFR2: 40.2±12.6 pg/g) compared with the control group (VEGF: 19.2± 8.6 pg/g; VEGFR2: 14.9±8.0 pg/g). Vascular endothelial growth factor and VEGFR2 showed no significant difference in the contralateral hemisphere treated with atorvastatin (VEGF: 10.8± 2.2 pg/g; VEGFR2: 5.6±4.0 pg/g) compared with the contralateral hemisphere of MCAO control mice (VEGF: 12.4±4.3 pg/g; VEGFR2: 4.9±3.1 pg/g) (P>0.05). These data suggest that atorvastatin enhances VEGF and VEGFR2 expression after stroke in the ischemic brain.
To test whether atorvastatin promotes neuronal migration and plasticity, DCX, TUJ1 and synaptophysin immunostaining were performed in the brain coronal sections. Figure 4 shows that atorvastatin treatment after stroke significantly increased DCX (B), TUJ1 (F) and synaptophysin (I) expression in the ischemic border, as compared with saline-treated mice (A, E and H). Increased DCX-positive cells were observed around vessels (C). These findings suggest that atorvastatin promotes cell migration and differentiation into neurons, and migrating neuronal cells are linked with vessels.
Figure 5 shows that atorvastatin (C) and BDNF (B) promote SVZ explant cell migration, compared with control (A). Anti-BDNF antibody (D) significantly inhibited cells migration compared with control (A). Coculture anti-BDNF with atorvastatin significantly inhibited atorvastatin-induced SVZ neural cell migration, compared with atorvastatin (C) alone group. These data suggest that BDNF amplifies neural cell migration.
Figure 6 shows that atorvastatin increases VEGFR2 (B) and BDNF (E) expression in cultured endothelial cells, as compared with control (A and D). Inhibition of VEGFR2 using a neutralizing anti-VEGFR2 antibody (DC101) significantly decreased atorvastatin-induced BDNF expression in cultured endothelial cells (F). These data indicate that atorvastatin promotes VEGFR2 and BDNF expression.
In the current study, we have demonstrated that a widely and clinically used HMG-CoA reductase inhibitor (statin), atorvastatin, when administered to mice starting 1 day after MCAO, evokes significant improvement in functional neurologic recovery. This observation is consistent with our previous studies using this compound in rats (Chen et al, 2003b). In addition, we have also shown that atorvastatin promotes: (1) neurogenesis and neuronal plasticity as well as increases the expression of VEGF, VEGFR2 and BDNF in the ischemic border after stroke in mice; (2) endothelial cell proliferation and VEGFR2 and BDNF expression; (3) neural cell migration; (4) BDNF expression, which mediates neural cell migration in SVZ explants. Collectively, these in vivo and in vitro data strongly support a role for atorvastatin in promoting brain plasticity and recovery from stroke.
Atorvastatin increases VEGF, VEGFR2 expression as well as promotes endothelial cell proliferation in the ischemic border. Vascular endothelial growth factor is an angiogenic factor. Vascular endothelial growth factor exerts biologic functions via two closely related receptor tyrosine kinases VEGFR1 (flt-1) and VEGFR2 (flk-1). Most of the VEGF properties, such as mitogenicity, chemotaxis and induction of morphologic change, are mediated by its interaction with VEGFR2 (Waltenberger et al, 1994). Vascular endothelial growth factor/VEGFR2 has been shown to be essential for endothelial cell proliferation and differentiation (Breier et al, 1992). Binding of VEGF to VEGFR2 leads to the receptor phosphorylation and subsequent activation of PI3K/Akt and other downstream signaling proteins (Gliki et al, 2002; Nakashio et al, 2002). Statin-induced capillary-like tube formation is inhibited by a neutralizing antibody against VEGFR2 and inhibition of PI3K (Chen et al, 2003b). We propose that administration of atorvastatin promotes VEGF and VEGFR2 expression in the ischemic border, which may facilitate induction of endothelial cell proliferation and angiogenesis.
The 10 mg/kg dose of atorvastatin used in this study is consistent with the neuroprotective dose of atorvastatin used in pretreatment of mice after stroke (Gertz et al, 2003; Laufs et al, 2000). We and others have shown that in vitro and in vivo, atorvastatin has a U-shaped dose–response curve on inducing angiogenesis (Chen et al, 2003b; Urbich et al, 2002; Weis et al, 2002). However, a 2.5 mg/kg dose of atorvastatin was shown to decrease angiogenesis in brain tumor and in a model of inflammation (Weis et al, 2002). This apparent discrepancy may be attributed to our use of the stroke model versus tumor model.
In addition to its role in inducing angiogenesis, VEGF also stimulates neurogenesis and axonal outgrowth, and improves the survival of mouse superior cervical, dorsal root ganglion neurons, and mesencephalic neurons (Hess et al, 2002; Sondell et al, 1999). Vascular endothelial growth factor is mitogenic for astrocytes and promotes growth/survival of neurons (Silverman et al, 1999). Our data have shown that atorvastatin promotes angiogenesis, neuronal plasticity as well as increases VEGF/VEGFR2 expression. We propose that the atorvastatin-induced increase of VEGF/VEGFR2 may not only cause angiogenesis but also provide a supportive microenvironment, which can enhance the neuronal and synaptic plasticity.
Neurogenesis and synaptic reorganization are important for functional improvement after stroke (Gomez-Fernandez, 2000; Hallett, 2001; Shingo et al, 2001). Our data show that atorvastatin treatment after stroke induces the expression of BDNF and synaptic proteins, and neuronal migration in the ischemic border, and some neuronal migrating cells are localized to blood vessels. These data are consistent with other reported studies showing that increasing synaptic activity elicits compensatory angiogenesis (Black et al, 1989). An index of axonal sprouting, GAP43, appears to be colocalized with angiogenesis after MCAO (Kawamata et al, 1997). Neurogenesis occurs in close proximity to blood vessels, where VEGF expression is high and angiogenesis is ongoing (Palmer et al, 2000). The newly activated and expanded vasculature substantially increases the production and release of BDNF, whose induction is both spatially and temporally associated with recruitment of new neurons (Leventhal et al, 1999). Cerebral endothelial cells are a potential source for bioactive BDNF (Bayas et al, 2002). Vascular endothelial cells may contribute to circulating BDNF (Nakahashi et al, 2000). Atorvastatin and BDNF induce neuronal migration in the SVZ explant culture. Inhibition of BDNF decreases neuronal migration in the SVZ explant. These data suggest that neuronal plasticity is closely linked with angiogenesis and BDNF may facilitate atorvastatin-induced neuronal plasticity after stroke.
In summary, treatment of stroke at 24 hours after MCAO in mice with a widely used atorvastatin reduces neurologic deficits associated with stroke. Statin-mediated functional benefit might be derived from the upregulation of trophic factors such as VEGF/VEGFR2, BDNF and induction of angiogenesis, neurogenesis, and synaptic plasticity.
We wish to thank Cynthia Roberts for technical assistance support. This work was supported by NINDS grants PO1 NS23393, PO1 NS42345 and RO1 NS47682.