Acute Hepatocyte Growth Factor Treatment Induces Long-Term Neuroprotection and Stroke Recovery via Mechanisms Involving Neural Precursor Cell Proliferation and Differentiation

Animals

Experimental procedures were performed according to the European Institutes of Health guidelines for the care and use of laboratory animals and approved by local authorities. Animals were housed under a regular circadian rhythm regimen with free access to food and water. In total, 119 (including sham operations) adult male C57/BL6N mice weighing 23 to 28 g were used. If not stated otherwise, each experimental group consisted of eight to nine animals. Mortality rates for animals of experimental groups including survival periods for up to 7 days were zero for each experimental condition. For animals surviving 4 weeks after induction of stroke, we observed survival rates of 88.9% (vehicle treated) and 87.5% (HGF treated). All experiments were performed in a blinded manner with regard to the kind of treatment chosen for each animal. For analysis of postischemic cell proliferation, animals were intraperitoneally injected with 5-bromo-2-deoxyuridine (BrdU; Sigma, Steinheim-Taufkirchen, Germany) on days 4 to 6 (7-day survival) or on days 8 to 18 (28-day survival), with a daily dose of 50 mg/kg body weight.

Induction of Focal Cerebral Ischemia

Induction of focal cerebral ischemia was performed as previously described (Doeppner et al, 2009). Briefly, animals were anesthetized (0.8% to 1.5% isoflurane, 30% O₂, remainder N₂O) and rectal temperature was maintained employing a feedback-controlled heating system. Regional cerebral blood flow was assessed using a laser Doppler flowmeter connected to a probe that was attached to the skull overlying the core region of the middle cerebral artery territory. Cerebral ischemia was induced by transient occlusion of the middle cerebral artery using a silicon-coated filament (180 μm diameter; Doccol, Redlands, CA, USA), which has been removed after 45 minutes to allow reperfusion of the middle cerebral artery. Laser Doppler flowmeter recordings continued for 15 minutes after thread removal to monitor appropriate reperfusion (>80% of initial regional cerebral blood flow). Sham-operated animals underwent the same experimental procedure except for insertion of silicon-coated monofilaments.

Stereotactic Intracerebral Injections

Stereotactic surgeries were performed during reperfusion under constant anesthesia as described above. The mice were placed in a stereotactic apparatus (Kopf Instruments, Düsseldorf, Germany) and the skull was exposed followed by drilling a hole at 0.4 mm anterior and 1.8 mm lateral (left ischemic hemisphere) from bregma. From these coordinates, intrastriatal injections were made 3.5 mm ventral to the bregma using a 10-μL Hamilton syringe (Switzerland). Control mice received 5 μL of 0.1 mol/L phosphate-buffered saline (PBS) per animal and day as vehicle, whereas treated animals obtained HGF at doses of 1, 5, or 10 μg on occasion of each injection, diluted in 5 μL 0.1 mol/L PBS. At the end of injection, the syringe was kept in place for additional 5 minutes before removal. For experiments with survival periods of 7 or 28 days, all animals received additional injections on days 1 to 3 using the same protocol as described above. For dose-finding studies, HGF was kindly provided by Genentech (San Francisco, CA, USA), whereas for the remaining experiments, HGF was purchased from R&D Systems (Wiesbaden-Nordenstadt, Germany). In the latter experiments, an HGF dose of 5 μg per animal and day was used.

Immunohistochemistry and Terminal Transferase dUTP Nick End Labeling

Animals were intraperitoneally injected with chloralhydrate (420 mg/kg body weight) and transcardiacally perfused with 4% paraformaldehyde on day 7 or day 28 after stroke. Brains were removed, postfixed in paraformaldehyde, embedded in paraffin, and coronal sections were prepared. Sections were deparaffinized, boiled in 0.2% citrate buffer, and incubated with blocking solution. For analysis of BrdU⁺ cells and Ki-67⁺ cells, sections were exposed to blocking solution and subsequently stained with a rat monoclonal anti-BrdU antibody (1:50; 18 hours, 4°C; Serotec, Düsseldorf, Germany) or a rabbit monoclonal anti-Ki-67 antibody (1:100; 18 hours, 4°C; Abcam, Cambridge, UK), followed by incubation with a goat antirat Alexa 594 (1:500, 1 hour; Molecular Probes, Darmstadt, Germany) or a donkey anti-rabbit Alexa Fluor 568 (1:400, 1 hour; Molecular Probes) secondary antibody. For differentiation analysis of BrdU⁺ cells, sections were incubated with a mouse monoclonal anti-BrdU antibody (1:500; Roche, Basel, Switzerland), a goat polyclonal antidoublecortin antibody (1:50; Santa Cruz Biotechnology, Heidelberg, Germany), a rat polyclonal anti-glial fibrillary acidic protein antibody (1:500; Zymed, Darmstadt, Germany), or a rat biotin-conjugated antiisolectin B₄ (IB₄) antibody (1:100; Vector, Peterborough, UK) for 18 hours at 4°C. For further microglia analysis, colocalization of IB₄ with major histocompatibility complex (MHC)-II was performed using a mouse monoclonal anti-MHC-II antibody (1:200; Santa Cruz Biotechnology). For double staining against BrdU and neuronal nuclei (NeuN) or 2',3'-cyclic nucleotide 3'-phosphodiesterase, a rat monoclonal anti-BrdU antibody (see above) and a mouse monoclonal anti-NeuN antibody (1:200, 18hours, 4°C; Chemicon, Schwalbach, Germany) or a mouse monoclonal anti-2',3'-cyclic nucleotide 3'-phosphodiesterase antibody (1:400; Chemicon) were used. After washing, sections were incubated for 1 hour at room temperature with the following secondary antibodies: goat anti-mouse Cy-3 (1:400; Dianova, Hamburg, Germany) or goat anti-rat Alexa 594 (1:400; Dianova) for MHC-II and BrdU staining, goat anti-rat Alexa 488 (1:250; Molecular Probes) or donkey anti-goat Alexa 488 (1:250; Molecular Probes) for glial fibrillary acidic protein or doublecortin staining, goat anti-mouse Cy-3 (1:100; Jackson Immuno-Research, Newmarket, UK) for 2',3'-cyclic nucleotide 3'-phosphodiesterase staining, and goat anti-mouse Alexa 488 (Molecular Probes) for NeuN (1:400) staining. Terminal transferase dUTP nick end labeling (TUNEL) staining was performed incubating sections with proteinase K (7 minutes, 37°C), followed by exposure to the TdT enzyme reaction according to the manufacturer's manual (Roche). After several washing steps and exposure to TUNEL blocking solution (20 minutes, room temperature), sections were stained with a streptavidin-Alexa594-conjugated secondary antibody (2 hours, room temperature; Molecular Probes). Quantitative analyses were performed in four regions of interest within the ischemic striatum located 0.14 mm anterior, 2.5 to 3.25 mm ventral, and 1.5 to 2.25 mm lateral from bregma. For each regions of interest, three sections per animal were evaluated, for which mean values were computed.

Infarct Volumetry and Determination of Edema Formation

Infarct volume analysis was performed on days 1 and 7 after stroke. Brains were removed and cut into four slices of 2 mm each. Slices were stained with 2,3,5-triphenyltetrazolium chloride (2%), and computer-based analysis of infarct volumes was performed using the software Image J by subtracting the area of the nonlesioned ipsilateral hemisphere from that of the contralateral side. Infarct volume sizes were calculated by integration of the lesioned areas. Postischemic brain edema was measured also using Image J software as the increase of ipsilateral hemispheric volume in comparison to the contralateral hemisphere as described by Wacker et al (2009).

Analysis of Motor Coordination Deficits

For analysis of poststroke motor coordination deficits, animals were evaluated using the corner-turn and tight-rope tests. One day before induction of stroke, all animals were trained before the beginning of the actual tests on days 7, 14, 21, and 28 after ischemia. Each test was performed twice per time point, for which means were calculated. For the tight-rope test, animals were placed on a 60-cm long tight rope grasping the string with their forepaws. Healthy animals grasp the string with four legs and tail and move to reach a side pole, whereas mice suffering from ischemia-induced motor deficits cannot lift their hind legs and eventually fall off onto the cage bedding placed underneath. Maximum test time was 60 seconds and the results were scored from 0 (min) to 20 (max) according to a validated score (Gerlai et al, 2000) depending on the time animals spent on the rope and whether or not they reached the platform. Scores were used for statistical analysis.

For the corner-turn test, animals were placed in a corner consisting of two vertical boards attached at one side with an angle of 30°. Each mouse was tested for the side chosen over 10 trials per day, that is right or left rearing in the corner. Whereas healthy animals leave the corner without side preference, mice suffering from stroke preferentially turn to the left, nonimpaired body side when leaving the corner. The laterality index was calculated according to the following formulae: (number of left turns—number of right turns)/10.

Blood—Brain Barrier Permeability

For this study, animals that were treated with vehicle, HGF (see above), or the matrix metalloprotease (MMP) inhibitor BB-1101 (0.75 mg per animal) at the beginning of reperfusion were used. Mice received intravenous injections of 2% Evans Blue (2 mL/kg) 22 hours after the stroke, as previously described (Chiba et al, 2008). Two hours later, animals were killed by transcardiac perfusion with 0.1 mol/L PBS. Brains were removed and separated into hemispheres. Left hemispheres were weighed, homogenized in 2 mL of 50% trichloroacetic acid, and centrifuged at 10,000 r.p.m. for 20 minutes. The extracted Evans Blue dye was then further diluted with ethanol, and the fluorescence signal was measured with a luminescence spectrophotometer (excitation at 620 nm, emission at 680 nm). An external standard (62.5 to 500 ng/mL) was used for the calculation of Evans Blue contents. Evans Blue contents of left hemisphere tissue was calculated from four animals as (μg) Evans Blue per (g) tissue.

Zymography of Matrix Metalloproteases

Left hemispheres from ischemic and nonischemic animals were homogenized in cold lysis buffer (basic buffer) containing 50 mmol/L Tris-HCl (pH 7.6), 150 mmol/L NaCl, 5 mmol/L CaCl₂, 0.05% BRIJ-35, 0.02% NaN3, and 1% Triton X-100. Homogenates were centrifuged at 4°C at 12,000 g for 5 minutes and supernatants were thereafter incubated with a 1:10 volume of gelatin sepharose 4B for 1 hour at 4°C with constant shaking. After centrifugation and resuspension of pellets in elution buffer (basic buffer containing 10% dimethyl sulfoxide (DMSO) and 20% volume of lysis buffer), purified samples were analyzed by zymography. Protein concentrations were determined by the bicinchoninic acid method (BCA kit, Thermo Scientific, Bonn, Germany).

Separation of MMP-2 and MMP-9 as pro-form and active form was performed using Novex Zymogram Gels (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions. Briefly, samples were incubated in nonreducing sample buffer (0.4 mol/L Tris, pH 6.8, 5% sodium dodecyl sulfate, 20% glycerol, 0.05% bromophenol blue) for 10 minutes at room temperature. Thereafter, the samples were loaded onto 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis gels containing 0.1% gelatin. After electrophoresis, the samples were incubated with 2.5% Triton X-100 twice for 20 minutes, equilibrated with developing buffer (Novex), and incubated for over 18 hours at 37°C. Gels were stained with Coomassie blue for 30 minutes and destained in washing solution (30% methanol, 10% acetic acid). White bands on a dark background indicated zones of digestion corresponding to the presence of pro-MMPs and activated MMPs on the basis of their molecular weight. As standards, 0.1 ng of human pro-MMP-9 and human pro-MMP-2 (Merck Biosciences, Darmstadt, Germany) and 0.01 ng of activated MMP-9 and activated MMP-2 (Merck Biosciences) were used. Thereafter, gels were scanned and densitometrically analyzed.

Preparation and Cultivation of Subventricular Zone-Derived Neural Precursor Cells

Subventricular zone-derived NPCs were prepared from 11-to 12-week-old C57/BL6N mice according to a previously described protocol (Doeppner et al, 2010). Subventricular zones were microdissected under stereomicroscopic control and minced into small pieces, mechanically triturated, and dissociated to single-cell suspension. Cells were cultured in serum-free basic medium (DMEM-F12; PAA, Pasching, Austria), supplemented with epidermal growth factor (2 μg/mL), basic fibroblast growth factor (2 μg/mL), and penicillin-streptomycin (Invitrogen, Darmstadt, Germany). Cells were incubated under standard cell culture conditions and growth factors were added every 2 to 3 days. Cells were passaged every 7 to 10 days and used for experiments after passage 3 or 4.

Oxygen—Glucose Deprivation

For each experiment, 100,000 NPCs were used (n = 4). Before induction of oxygen—glucose deprivation (OGD), cell culture medium was substituted by a crystalloid solution (‘Thomajodin’ plus 1 mM mannitol; DeltaPharm, Dortmund, Germany), and the cells were incubated in a hypoxic chamber (1% O₂, 5% CO₂, and 94% N₂) for 45 minutes and reincubated in normal glucose containing cell culture medium for 24 hours. Control cells were incubated with 0.1 mol/L PBS 1 hour before OGD, whereas other cells were treated with recombinant human HGF at a final concentration of 20 ng/mL. The number of dead cells was counted using a LIVE/DEAD-Viability-Cytotoxicity-Assay kit (Lonza, Basel, Switzerland).

Caspase-3 Assay

Caspase-3 activity was measured in cultivated NPCs (n = 4 per condition) at 1, 2, 4, and 8 hours after OGD using the fluorophore substrate Ac-DEVD (aspartic acid-glutamic acid-valine-aspartic acid)-7-amino-4-methylcoumarin (Biomol, Hamburg, Germany). Cells were incubated either with HGF (20 ng/mL) or with the caspase-3 inhibitor z-DEVD.fmk (50 μmol/L, solved in 0.2% DMSO; Biomol) at the beginning of OGD. Controls were incubated with either 0.1 mol/L PBS or 0.2% DMSO in 0.1 mol/L PBS. At the time points given, NPCs were complemented with lysis buffer that contained 25 mmol/L HEPES (pH 7.5), 1 mM EDTA, 10 mM 1,4-dithiothreitol (Roche), 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (Sigma-Aldrich), 10% sucrose, 0.1% Triton X-100, with 10 μg/mL of pepstatin A, aprotinin, and leupeptin each plus 1 mM PMSF (all Sigma-Aldrich) as protease inhibitors. Thereafter, the cells were centrifuged and supernatants were used for caspase-3 activity measurement. For each measurement, 90 μL of samples were incubated with 10 μL of substrate (50 μmol/L final concentration) and fluorescence (355 to 460 nm) was detected at 2-minute intervals for 20 minutes. The increase in fluorescence was linear between 2 and 16 minutes. Caspase-3 activity was recalculated from the slope (fluorescence units per time) and is given as picomoles of substrate cleaved per milligram of protein per minute based on a standard curve for 7-amino-4-methylcoumarin (Biomol). Protein concentration in the supernatant was determined using the Bradford assay.

Statistics

All data are given as means ± s.d. For comparison between two groups, the Student's t-test was used, whereas for a multigroup comparison, a one-way analysis of variance followed by the Tukey's post hoc test was performed. A P value of < 0.05 was considered to be statistically significant.

Hepatocyte Growth Factor Induces Neuroprotection in a Dose-Dependent Manner

Although HGF-induced neuroprotection has been described before, these studies were mostly performed on rats and included either continuous release or overexpression of HGF. We therefore first determined whether or not a single injection of HGF is sufficient to induce neuroprotection after transient focal cerebral ischemia in mice (Figure 1A). When 1 μg of HGF was injected into the ischemic striatum during early reperfusion, no effect on infarct size was observed 24 hours after stroke. On the contrary, HGF at dosages of 5 or 10 μg, respectively, yielded significant neuroprotection after stroke. In line with this, ipsilateral edema formation was significantly reduced in animals treated with 5 or 10 μg of HGF, but not in animals treated with 1 μg of HGF (Figure 1B). Hepatocyte growth factor-induced neuroprotection was not associated with changes in regional cerebral blood flow (Figure 1C); animals from all groups exhibited similarly high reperfusion rates. As no significant difference with regard to infarct volume and edema formation between animals treated with 5 or 10 μg of HGF was observed, further in vivo experiments were performed using doses of 5 μg only.

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Figure 1

Hepatocyte growth factor (HGF) reduces postischemic injury and edema formation in a dose-dependent manner. Hepatocyte growth factor was injected at different dosages into the left ischemic striatum at the beginning of reperfusion followed by infarct analysis 24 hours after stroke (A). Ipsilateral edema formation at 24 hours after stroke is given as relative increase of ipsilateral hemisphere volume as compared with the contralateral nonischemic hemisphere (B). Note that HGF-induced neuroprotection was not associated with regional cerebral blood flow changes (C). Data (n = 6 animals per group) are presented as means ± s.d. *Significantly different from vehicle.

Hepatocyte Growth Factor-Induced Neuroprotection Persists in the Postacute Stroke Phase, and Is Associated with Ameliorated Motor Coordination Deficits

As acute reduction of infarct size and edema formation (Figure 1) does not imply sustained neuroprotection or beneficial functional outcome, ischemic injury was further analyzed on both day 7 and day 28 after the stroke (Figure 2). Treatment with HGF resulted in persistent neuroprotection as assessed by means of both TUNEL (day 7) and NeuN (day 28) staining (Figure 2A). We observed 407.6 ± 54.8 TUNEL⁺ and 701.8 ± 62.2 NeuN⁺ cells per mm² within the periinfarct area of HGF-treated animals compared with 649.0 ± 80.3 TUNEL⁺ and 533.3 ± 50.9 NeuN⁺ cells per mm² in control animals. In line with the sustained neuroprotection, animals treated with HGF showed reduced postischemic functional deficits as assessed by both the corner-turn and the tight-rope tests (Figures 2B and 2C). Although animals from both vehicle and HGF groups gradually improved over time, animals treated with HGF always performed better than vehicle-treated animals, thereby almost approximating test performance of healthy sham-operated animals.

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Figure 2

Sustained neuroprotection and amelioration of functional neurologic deficits induced by hepatocyte growth factor (HGF). Treatment with HGF significantly reduced the number of terminal transferase dUTP nick end labeling (TUNEL⁺) cells on day 7 after stroke (A) and significantly increased the density of NeuN⁺ neurons on day 28 (A). Long-term neuroprotection by HGF was accompanied by enhanced motor coordination performance in both the corner-turn (B) and the tight-rope (C) tests. Data are presented as means ± s.d. *Significantly different from vehicle. NeuN, neuronal nuclei.

Hepatocyte Growth Factor Stabilizes Blood—Brain Barrier Integrity via Inactivation of Matrix Metalloproteases

As development of acute infarct injury involves enhanced blood—brain barrier (BBB) permeability, we analyzed HGF-mediated effects on Evans Blue extravasation 24 hours after induction of stroke. Whereas control animals showed distinct Evans Blue extravasation, animals treated with HGF displayed significantly reduced extravasation of the dye (Figure 3A). Similarly, treatment with BB-1101 (0.75 mg per animal), a broad inhibitor of MMPs that are critically involved in BBB breakdown, also resulted in reduced Evans Blue extravasation. Whereas HGF treatment lead to subacute neuroprotection as suggested by decreased infarct volumes on day 7, animals treated with BB-1101 did not benefit from this therapy. As such, infarct volume was not influenced by BB-1101 (Figure 3B). Matrix metalloproteases gelatin zymography (Figures 3C-3E) revealed activation of MMP-2 in vehicle-treated control animals 4 hours after the stroke, which was not found in sham-operated animals. Both HGF and BB-1101 treatments reduced MMP-2 activation without changing levels of inactive pro-MMP-2. Activation of MMP-2 was, however, only transient, decreasing below levels of detection within 24 hours after stroke. On the other hand, active MMP-9 was only detected as late as 24 hours after stroke. Matrix metalloprotease-9 but not pro-MMP-9 activity was again reduced by HGF and BB-1101.

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Figure 3

Hepatocyte growth factor (HGF) reduces blood—brain barrier (BBB) permeability and inhibits matrix metalloproteases (MMP). Blood—brain barrier permeability (A) was assessed 24 hours after stroke by means of Evans Blue extravasation (n = 4 animals per group). Infarct volumes (B) were analyzed on day 7 after stroke using 2,3,5-triphenyltetrazolium chloride (TTC) stainings. (C) Gelatin zymography of brain homogenates was performed at 4 or 24 hours after stroke (n = 4 animals per group). Animals were treated at the beginning of reperfusion. Control animals underwent sham surgery without further treatment. Human pro-MMPs (0.1 ng) and activated MMPs (0.01 ng) served as standards. (D, E) Densitometric analyses of activated MMP-2 (D) and activated MMP-9 (E) from gelatin zymography at 4 or 24 hours after stroke, respectively. Data are presented as means ± s.d. *Significantly different from vehicle.

Hepatocyte Growth Factor Enhances Postischemic Cell Proliferation

To further elucidate the mechanisms underlying the sustained neuroprotection and recovery induced by HGF, cell proliferation was analyzed using the thymidine analog BrdU on day 7 and day 28 after stroke (Figure 4A). Although cell proliferation declined over time within the ischemic striatum of animals treated with either HGF or vehicle, the number of BrdU⁺ cells in animals treated with HGF was always higher than in vehicle-treated controls. As BrdU labeling does not exclusively reflect current cell proliferation, analysis of Ki-67⁺ cells was additionally performed at the time points given (Figure 4B). Similarly, HGF treatment resulted in enhanced numbers of Ki-67⁺ cells for up to 4 weeks after induction of stroke, demonstrating that HGF enhances postischemic endogenous cell proliferation.

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Figure 4

Hepatocyte growth factor (HGF) promotes postischemic cell proliferation. Postischemic treatment with HGF resulted in increased cell proliferation as assessed by 5-bromo-2-deoxyuridine (BrdU) (A) and Ki-67 (B) staining. Cell proliferation was increased at both time points analyzed. Note a constant decline of proliferating cells over time in vehicle- and HGF-treated animals. Data are presented as means ± s.d. *Significantly different from vehicle.

Endogenous Poststroke Neurogenesis Is Enhanced After Hepatocyte Growth Factor Treatment

Differentiation analysis of BrdU⁺ cells revealed high relative amounts of BrdU⁺ cells expressing the microglial marker IB₄ on day 7 after stroke, suggesting that a majority of proliferating cells rather reflect microglia than newborn neural cells (Figure 5A). There was, however, no significant difference between animals treated with either HGF or vehicle. With regard to the state of activation of IB₄⁺ microglial cells, double staining against IB₄, and MHC-II on day 7 revealed no difference between vehicle and HGF group. We observed 93.1% ± 12.5% in the vehicle and 89.6% ± 7.5% of colocalizations in the HGF group, representing a high state of activation of microglial cells. Regarding neural differentiation of BrdU⁺ cells, animals treated with HGF showed significantly higher numbers of BrdU⁺ cells expressing the immature neural marker nestin than controls, albeit cell numbers declined over time in both experimental groups (Figure 5B). On the contrary, the number of BrdU⁺ cells expressing glial fibrillary acidic protein gradually increased over time in animals treated with both HGF or vehicle (Figure 5C). There was, however, no significant difference between these two groups. Likewise, oligodendroglial differentiation of BrdU⁺ cells was not affected by HGF (Figure 5D). Interestingly, long-term neuronal differentiation was enhanced in animals treated with HGF compared with vehicle-treated mice (Figures 5E–5H). Hepatocyte growth factor treatment resulted in increased numbers of BrdU⁺ cells expressing either doublecortin or NeuN at day 28 after stroke when compared with control animals. Whereas no mature neuronal differentiation of newborn proliferating cells was observed on day 7 (data not shown), the number of BrdU⁺ cells expressing NeuN was still low on day 28 in both experimental groups.

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Figure 5

Hepatocyte growth factor (HGF) promotes neural differentiation. Fate of differentiation of 5-bromo-2-deoxyuridine (BrdU⁺) proliferating cells was analyzed using the microglial marker isolectin B₄ (IB₄) (A), the neural marker nestin (B), the astroglial marker glial fibrillary acidic protein (GFAP) (C), the oligodendroglial marker 2',3'-cyclic nucleotide 3'-phosphodiesterase (CNPase) (D), the immature neuronal marker doublecortin (Dcx) (E, F), and the marker for mature neuronal cells NeuN (G, H). Arrows indicate colocalization of BrdU⁺ cells (red) with the marker in question (green), that is Dcx (F) or NeuN (H). Scale bars: 40 μm (F) and 20 μm (H). Data are presented as means ± s.d. *Significantly different from vehicle.

Hepatocyte Growth Factor Stimulates Neural Precursor Cell Proliferation In Vitro and Protects Neural Precursor Cells Against Hypoxic Injury

As enhanced cell proliferation (Figure 4) can be either due to stimulation of cell division itself or due to the protection of proliferating cells, we analyzed NPC proliferation under physiological conditions in vitro. When NPCs were cultivated under standard cell culture conditions, neurosphere formation was observed in NPCs treated with either vehicle or HGF (Figures 6A and 6B). However, incubation of NPCs with HGF resulted in significantly higher numbers of neurospheres as compared with control NPCs (Figure 6C). Likewise, treatment of NPCs with HGF resulted in significantly enhanced numbers of NPCs incorporating BrdU (Figure 6D), suggesting that HGF stimulates proliferation of SVZ-derived NPCs in vitro. Apart from HGF-mediated stimulation of NPC proliferation, HGF also preserved cultivated NPC from injury when exposed to 45 minutes of OGD followed by subsequent reexposure to oxygen and glucose under standard cell culture conditions (Figure 6E). Whereas control cells developed prominent and distinct cell injury, NPCs incubated with HGF either 4 hours before OGD or at the beginning of OGD showed significantly enhanced survival (Figure 6E). Furthermore, HGF-induced neuroprotection of NPCs in vitro was associated with attenuated caspase-3 activation. As such, significantly reduced caspase-3 activity was observed for up to 8 hours after OGD in cells that were treated with HGF. As a matter of fact, caspase-3 activities in HGF-treated cells were as low as caspase-3 activities from NPCs treated with z-DEVD.fmk, a well-known inhibitor of caspase-3. These data suggest that HGF promoted the proliferation and survival of NPCs via a mechanism involving caspase-3.

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Figure 6

Hepatocyte growth factor (HGF) increases proliferation of neural precursor cells (NPCs) and protects against hypoxic injury in a caspase-3-dependent way in vitro. Neural precursor cells cultured in serum-free basic medium were treated with vehicle (= 0.1 mol/L phosphate-buffered saline (PBS)) (A) or HGF (B) every second day. (C) Neurosphere formation was quantitatively analyzed on day 7 at passage 3 to 4. For assessment of NPC proliferation (D), cells were cultured as described above, but were additionally incubated with 5-bromo-2-deoxyuridine (BrdU) 4 hours before the end of the experiment. (E) NPCs were exposed to oxygen—glucose deprivation (OGD) for 45minutes followed by 24 hours recultivation under standard cell culture conditions. In some experiments (‘HGF before OGD’), vehicle or HGF was added 4 hours before induction of OGD, whereas in other experiments, vehicle or HGF was administered at the beginning of OGD only (‘HGF after OGD’). (F) Caspase-3 activity was determined at 1, 2, 4, and 8 hours after the end of OGD in NPC lysates by means of caspase-3-mediated cleavage of nonfluorescent z-DEVD-7-amino-4-methylcoumarin (AMC) into fluorescent AMC (‘AMC release’). AMC release is given as pmol/min referred to total protein contents. Cells were incubated with HGF (20 ng/mL) or the caspase-3 inhibitor z-DEVD.fmk (50 μmol/L) before induction of OGD. Phosphate-buffered saline and 0.2% DMSO served as control vehicles. Data are presented as means ± s.d. out of four experiments (A—F). *Significantly different from vehicle. Scale bar: 100 μm. DEVD, aspartic acid-glutamic acid-valine-aspartic acid; DMSO, dimethyl sulfoxide.