Traumatic spinal cord injury is usually accompanied by development of cystic cavities surrounded by glial scars which severely impair regeneration of severed axons (Sandvig et al., 2004; Silver and Miller, 2004; Yiu and He, 2006). Glial scars are composed of various types of cells and extracellular matrices. Of these, astrocytes play key roles in the formation of glial scars. Upon injury, they upregulate the expression of intermediate filament proteins (known as glial fibrillary acidic proteins, GFAP) to become hypertrophied (Pekny and Nilsson, 2005). These reactive astrocytes also secret various extracellular matrix (ECM) proteins and form dense physical barriers together with ECM molecules. Although glial scars may exert some beneficial effects by regulating local immune responses and promoting tissue repair (Faulkner et al., 2004; Rolls et al., 2009), mounting evidence indicates that glial scars constitute a major component of post-injury environment unfavorable to spontaneous axonal regeneration (Yiu and He, 2006).
Formation of astrocytic glial scars impedes growth of regenerating axons both physically and chemically. Hypertrophic astrocytes pose mechanical barriers to growing axons (McKeon et al., 1991). Deletion of both GFAP and vimentin, major intermediate filaments of astrocytes upregulated after CNS injury, resulted in enhancement of axonal regeneration after spinal cord injury (Ribotta et al., 2004). At the same time, they also produce growth inhibitory extracellular matrix (ECM) molecules such as chondroitin sulfate proteoglycans (CSPGs) (McKeon et al., 1999; Tang et al., 2003). CSPGs, potent inhibitory ECM molecules, consist of a core protein and many sugar side chains, which are large, sulfated
glycosaminoglycans (GAG) chains covalently attached to core proteins (Smith and Strunz, 2005; Galtrey and Fawcett, 2007). There are several different CSPG species upregulated after SCI such as neurocan, brevican, phosphacan, and NG2 proteoglycan (Jones et al., 2003;
Tang et al., 2003). Regardless of different classes of core proteins, major inhibitory functions appear to reside in GAG chains. Several studies have reported that enzymatic degradation of GAG chains by chondroitinase ABC enhances axonal regeneration and improves functional recovery after SCI (Bradbury et al., 2002; Caggiano et al., 2005; Tester and Howland, 2008;
Tom and Houle, 2008). These studies highlight importance of manipulating reactive astrocytes and ECM proteins to enhance regeneration and functional recovery after SCI.
Previous studies have shown that various inflammatory cytokines are implicated in induction of reactive astrocytosis. Especially, early induction of TGFβs after SCI is considered to be essential in reactive astrocytosis (Logan et al., 1994; Reilly et al., 1998;
Fitch et al., 1999; Gomes et al., 2005; Buss et al., 2008). Treatment of neutralizing antibodies against TGFβs led to marked attenuation of CNS scarring, suggesting their causative role (Logan et al., 1999; Moon and Fawcett, 2001). In addition to TGFβs, however, many other factors are implicated in activation of astrocytes at least in vitro. Several proinflammatory cytokines such as IL-1β, IFNγ and TNFa (Hewett et al., 1993; John et al., 2003) also strongly induce astrocytosis. Furthermore, it has been shown that nitric oxide (NO) (Brahmachari et al., 2006), oxidative stress (Rohl et al., 2008) and ciliary neurotrophic factor (CNTF) (Hudgins and Levison, 1998; Levison et al., 1998) could elicit reactive astrocytosis.
Therefore, it would be very challenging to identify a specific target of which blockade could effectively prevent reactive astrocytosis.
Hepatocyte growth factor (HGF) was first discovered and cloned as a mitogenic polypeptide for hepatocytes (Nakamura et al., 1989). HGF exerts its actions through tyrosine kinase receptor, c-Met (Shimamura et al., 2007). Beyond its classical effects on hepatocytes, subsequent studies have uncovered diverse functions of HGF in nervous system. It has a crucial role in developing neurons; HGF promotes neurite outgrowth from motor neuron (Ebens et al., 1996; Yamamoto et al., 1997) and improves neuron survival (Maina and Klein, 1999). It also stimulates proliferation and migration of Schwann cells (Krasnoselsky et al., 1994), olfactory interneuron precursors (Garzotto et al., 2008), and oligodendrocyte precursor cells (Yan and Rivkees, 2002; Ohya et al., 2007). One of the well established functions of HGF outside CNS is to suppress fibrosis of internal organs such as the liver and kidney (Matsumoto and Nakamura, 1992). It has been shown that liver fibrosis is actively processed by TGFβ1 (Border and Noble, 1994) and inhibition of TGFβ1 prevents the progression of liver fibrosis and even improves hepatocyte regeneration (Nakamura et al., 2000). HGF also functions to suppress fibrosis of kidney by inhibiting TGFβs production (Liu, 2004, 2006; Liu and Yang, 2006). Regulation of proteoglycans synthesis (Kobayashi et al., 2003) and extracellular matrix remodeling by HGF further contributed to attenuation of fibrosis in the internal organs (Kim et al., 1997; Liu and Yang, 2006). Interestingly, formation of fibrosis in the liver or kidney shares similarity with glial scarsring in CNS in that TGFβs are critically involved and deposition of ECM constitute an important component.
Based on this reasoning, it could be hypothesized that HGF may play a role in modulation of CNS glial scars. However, the possibility that HGFs may play an inhibitory role in the glial scars formation has not been addressed to date.
The purpose of this study was to examine whether HGFs can affect formation of glial scars and the production of various CSPGs. We found that HGF prevented both a formation of astrocyte activation and a production of chondroitin sulfate proteoglycans in activated astrocyte culture system, suggesting that HGFs may regulate scar formation. These effects appeared to be mediated by decreasing the level of TGFβ1. HGF’s effects on glial scars were also demonstrated in vivo by transplantation of HGF overexpressing mesenchymal stem cells into hemisected spinal cord tissue. Our data suggest that HGFs evidently regulate both scar formation and production of CSPGs following spinal cord injury.
Ⅱ. MATERIALS AND METHODS
A. Primary astrocyte cultures
Primary astrocyte cultures from cerebral cortex were prepared from postnatal 1 day pups as described previously (Wilson and Dixon, 1989). The upper part of the skull was separated and the meningeal tissue removed. The cerebral neocortices were isolated, placed into complete medium, which were containing 10% fetal bovine serum (FΒS), 0.01 M HEPES, 1% penicillin streptomycin in MEM/EΒSS (Minimum Essential Medium with Earle's Balanced Salts, Hyclone). The neocortices were homogenized by pipetting, were plated onto 175 cm2 tissue culture flasks (Corning). The cultures were maintained in a humidified atmosphere of 95% air 5% CO2 at 37℃ for 10~14 days after plating. Flasks were slapped to detach nonadherent cells from the bottom of flasks and then were washed with PΒS. Adherent astrocytes were removed by treatment with trypsin (0.25%, Gibco), were resuspended in complete medium, and suspended cells properly seeded in each condition.
Then microglia and oligodendrocytes were extracted. Once cells reached confluence (5-7 days), they were plated into 100 mm2 (Falcon) at a density of 1.3 X 106 cells/dish. Cells were passaged again and plated. The final percentage of GFAP-expressing cells in these cultures was found to be >95%. When the cells reached confluence, culture medium was changed to serum-free MEM containing 1 % penicillin streptomycin (P/S) and 0.01M HEPES before cytokine treatment. Astrocytes were treated in serum free medium with IL-1β (R&D systems) 10 ng/ml and IFNγ (R&D systems) 10 ng/ml, or TGFβ1 (Peprotech) 10 ng/ml with or
without HGF (Millipore) of various concentrations for 24 hours.
B. Animals and surgical procedures
Adult female Sprague Dawley rats weighing 200-250 g were used in this study. SD rats were housed in the Ajou University Animal Care and Use Committee. For surgery, animals were anaesthetized with chloral hydrate (400mg/kg, i.p.), received a dorsal laminectomy to expose the spinal cord. The dura were opened and iridectomy scissors were used to create a spinal cord right-over hemisected injury at Thoracic 8 level. Vacuum suction was used to aspirate the remaining tissues. Transplants were prepared with the concentration of 4.0 X 104 cells/ul and 5 μl of cell suspensions (total 2.0 X 105 cells) were soaked into gelfoam pledgets.
The gelfoams were immediately implanted into right-hemisection lesions. And then rats by each transplantation groups were injected 4 sites in rostral and caudal of injured spinal cord using a hamilton syringe. Each site was given 1 ul of transplant which is prepared 4 X 104 cells/ul concentrations. So, rats totally transplanted 3.6 X 105 cells / 9ul by concentration.
Experimental groups were divided into control group with phosphate buffered saline (PΒS) soaked gelfoam, MSC only group, and HGF-MSC group with gelfoams soaked with HGF overexpressing MSCs. All animals received daily intraperitoneal cyclosporine (NORVATIS) at a dosage of 10 mg/kg beginning from one day before transplantation to sacrifice.
Prophylactic antibiotics (cefazolin) were intraperitoneally injected on the next day after each surgery, and a bladder care had been provided twice daily until the animals resumed self-voiding.
C. Immunocytochemistry
For immunocytochemistry, cells were plated on 9 mm or 12 mm coverslips coated with poly-D-lysine (Sigma). Attached cells on coverslip were fixed in 4% paraformaldehyde (PFA) for 30min at room temperature (RT) after 3 times PΒS washing each for 10 minutes. After blocking with 10% normal goat serum for 1hr, primary antibodies were applied in the same blocking solution at 4℃ overnight or for 4 hours at RT. After thorough washing with PΒS, appropriate secondary antibodies tagged with Alexa Fluor 488 or Alexa Fluor 594 (Molecular Probes, Eugene, OR) were applied for 1hr at RT. The primary antibodies used in this study were mouse anti-GFAP (Dako; 1:400), rabbit anti-c-met (Santa Cruz; 1:100), rabbit anti-phospho-c-Met (Invitrogen; 1:100), mouse anti-CS-56 (Sigma; 1:400), mouse anti-neurocan (Millipore; 1:100), mouse anti-human mitochondria (Chemicon; 1:400) and rabbit anti-Ki67 (novocasta; 1:400). The coverslips were mounted onto slides with glycerol based mounting medium (Biomeda, Foster City, CA). The images were taken using a FV 300 confocal microscope (Olympus, Tokyo, Japan). These experiments are repeated several trials by experiment number of each group as described the table 1.
D. Tissue processing and immunohistochemistry
Rats were anesthetized with an overdose of chloral hydrate and perfused with heparinized saline (0.9%) followed by 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer. The spinal cord containing lesion site was dissected and post-fixed in 4% PFA for 2 hours, followedby cryoprotection in a graded series of sucrose solutions. Cryosections for
spinal cord (20 μm thickness) were made longitudinally in a 1:10 series, mounted onto Super Frost plus slides (Fisher Scientific, Pittsburgh, PA), and stored at -20℃ until use. For immunohistochemistry, longitudinal tissue sections were treated with 10% normal goat serum (Hyclone, Logan, VT) to prevent nonspecific immunoreactivity. Primary antibodies dissolved in the same blocking solution were applied onto tissue sections at 4℃ overnight followed by appropriate secondary antibodies tagged with Alexa Fluor 488 or Alexa Fluor 594 (Molecular Probes, Eugene, OR) for 1hr at RT. The primary antibodies used for immunohistochemistry were same as those for immunocytochemistry. To quantification of fluorescence intensity, such as GFAP and CS-56, we analyzed confocal images of 3 sites that are rostral, caudal and contra lateral beyond the glial limitance from 3 cryo-section at each group.The images were taken using a FV 300 confocal microscope (Olympus, Tokyo, Japan).
Immunohistochemistry is repeated several trials by experiment number of each group as described the table 4.
E. Enzyme-Linked Immunosorbent Assay (ELISA)
Cultured supernatants or spinal cord lysates were subjected to ELISA analysis to determine the level of TGFβs and HGF. For cultured supernatants, primary astrocyte cultures were treated with cytokines with or without HGF (as described above) for 24 hours and supernatants were collected. Tissue samples were homogenized with a dounce tissue grinder in ice-cold homogenization buffer (50mM Tris-HCl, pH 7.6, containing 150 mM NaCl, 1mM EDTA, 0.32M sucrose) supplemented with 1 homogenized with a douncetor cocktail (Roche,
Mannheim, Germany). The homogenates were then centrifuged at 12,000 rpm for 10 min and the supernatants were used for ELISA analysis. Protein concentration of the cultured supernatants or tissue homogenates were measured using Micro BCA Protein Assay kit (Pierce, Rockford, IL), and equal amounts of samples (typically 10 to 20 µg) were loaded into 96 well plates coated with capture antibodies. Concentration of HGF was assayed using human HGF ELISA kit (immunis, EIA, Japan) and that of TGFβ1 and β2 using kits from R&D systems (MN, USA). Detailed procedures followed an instruction manual provided by corresponding manufacturers. ELISA is tried by experiment number of each group as described the table 2 and table 4.
F. Western Blot analysis
One week after surgery, 17 rats were anesthetized with overdose chloral hydrate and briefly perfused with ice-cold saline to remove blood cells. Five mm-long spinal cord blocks containing lesion site were quickly dissected and homogenized in tissue extraction buffer containing protease inhibitor cocktail (Pierce) and phosphatase inhibitor (Halt Phosphatase Inhibitor Cocktail, Themo), which safeguards against serine, threonine and protein tyrosine phosphatase activities, on ice. Protein concentration was determined using the BCA protein Assay kit (Pierce). Equal amounts (24 ug / 30ul) of proteins were analyzed by SDS-PAGE, together with size marker (Thermo, Rockford, IL). It is performed in 10% polyacrylamide gels with 8% stacking gels. Proteins were separated by constant voltage of 130 mV in RT after separated by 60 mV in stacking gels and transferred to polyvinylidene difluoride
(PVDF) membranes (Immobilon-P; Millipore, Bedford, MA). The membranes were washed in Tris-buffered saline (TΒS-, 0.9% NaCl, 10mM Tris-HCl pH 8.0) containing 0.1% Tween-20 (Sigma) (TΒS-T) and blocked 5% skim milk for 2 hours at RT. The membranes were then incubated with following primary antibodies in 5% ΒSA solution. : anti-GFAP (Dako) and anti-c-met (SantaCruz, 1:1000). Especially anti-phospho-c-met (Invitrogen, 1:1000) were used in 1% ΒSA solution. β-actin was used as a loading control for cell lysates After incubation with horseradish peroxidase conjugated secondary antibodies, immunoreactivity was visualized by chemiluminescence reagents (ECL advance western blot detection kit, GE healthcare).
For neurocan detection, tissue lysates were treated with 0.03 U chondroitinase ABC (protease-free, SEIKAGAKU), which is cut the GAG chain from core protein, for 2 hours at 37℃. SDS-PAGE was performed in 6% polyacrylamide gels with a 4% stacking gel.
Proteins were prepared by 24 ug/30ul concentration and separated by constant voltage of 60 mV in RT after separated by 40 mV in stacking gels. Separated proteins in gel were transferred to PVDF membranes (Millipore) at 4℃ for 15 hours by constant current of 150mA in cold room. The blot membrane was rinsed three times in Tris-buffered saline (TΒS, 0.9% NaCl, 10mM Tris-HCl pH7.5) containing 0.05% Tween 20 (Sigma) (TΒS-T) and blocked 5% skim milk for 2 hours at RT. The blot was incubated with anti-neurocan (Millipore) in 5% skim milk solution. Anti-neurocan antibody can detect the 1D2 (150-163 kDa, C-terminal epitope of neurocan, ), 1F6 (122-130 kDa, N-terminal epitope of neurocan) and intact neurocan (240-270kDa) (Asher et al., 2000). After incubation with horseradish peroxidase conjugated secondary antibodies dissolving in 5% skim milk, immunoreactivity
was visualized by chemiluminescence reagents (ECL advance western blot detection kit, GE healthcare). This analysis is repeated several trials by experiment number of each group as described the table 4
G. Reverse Transcription polymerase chain reaction (RT-PCR)
Total RNA was extracted from cultured cells using Trizol (Gibco) according to the manufacturer’s protocol. The amount of RNA was determined using spectoscopy at 260 nm.
Five ug of RNA was reverse transcribed to cDNA using a standard RT protocol. One ul (0.4 ug) of cDNA was added to PCR-reaction premix (GenDEPOT) with 10 pM corresponding primer pairs. The following primers were used for polymerase chain reaction : GAPDH, 5’-GTG TAG TTC ACG CCC ACG TC-3’ (forward), 5’-5’-GTG ATG GCA TGG ACT 5’-GTG GT-3’
(reverse) ; neurocan, 5’-GCC ACA CTC TAC ACT CGT CCC-3’ (f), 5’TCT CCC CAG CAT AGC CCT GAT-3’(r) ; phosphacan, 5’-GCA AGT CCT GCC GTC CTT GCA 3’-(f), 5’-GGA ATA GGG ATT AGT AAC AGC-3’ (r) ; c-Met, 5’-TGT CTC TGA AAT CCA CCC GA 3’ (f), 5’ GTG TAG TTC ACG CCC ACG TC-3’ (r). Their specificity was verified using the basic local alignment search tool (BLAST) on the GenBank database. PCR amplification was performed with 35 cycles of 95 ℃ for 30 s, 55 ℃ ~ 60 ℃ (property Tm) for 30 s, 72 ℃ for 90 s. The PCR products were separated on a 1% agarose gel and stained by Ethidium Bromide (EtBr). The amount of each product was quantified by a gel document system (Bio-Rad). GAPDH expression was used as an internal reference to verify equal concentrations of cDNA in each sample. We have executed experimental number of each group as described
the table 1. And we quantified mRNA expression level of neurocan and phosphacan by band thickness using the image J program based on band thickness of non-TGFβ1 treatment.
H. Statistical Analysis
Statistical comparison of mean values was performed using one-way ANOVA followed by Tukey’s post hoc tests. All values are expressed as mean ± SD. Quantitation graphs were generated by GraphPad Prism version 4.00 (GraphPad Software, San Diego, CA, USA)
Ⅲ. RESULTS
A. HGF prevents cytokine-induced astrocytic activation in vitro.
To examine whether HGF affects formation of glial scars, we used primary cultured astrocytes isolated from 1 day pub rat’s neocortex to mimic glial scars (Fig. 1A). When more than 95% cultured astrocytes express glial fibrillary acidic protein (GFAP), we induced astrocyte activation by treatment of TGFβ1 at a concentration of 10 ng/ml (Baghdassarian et al., 1993; Reilly et al., 1998) (Fig. 1B, C). Immunocytochemistry revealed that TGFβ1-treated astrocytes upregulated GFAP positive intermediate filaments and showed hypertrophic morphology. HGF was added to the medium at three different concentrations: 5 ng/ml, 50 ng/ml, and 250 ng/ml. Fifty ng/ml and 250 ng/ml treatment of HGF obviously prevented morphological change, although 5 ng/ml was not effective (Fig. 1E-G). To quantify changes in astrocytic morphologies, we measured mean GFAP positive areas (Fig.
1G). We found that treatment of TGFβ1 clearly increased the mean size of astrocytes and co-treatment of HGF at 50 ng/ml and 250 ng/ml significantly reduced the extent of hypertrophic changes.
It has been previously reported that HGF stimulated neuronal cell survival and oligodendrocyte precursor cell proliferation (Yan and Rivkees, 2002; Ohya et al., 2007). To examine whether HGF affects proliferation of astrocytes, cells were stained with anti Ki-67, which is a cellular marker of proliferation and detects nuclei-during interphase. Ki-67 immunoreactivity was not different between different treatment groups (Fig. 2A-C).
Quantification of the number of Ki-67 positive cells showed that proliferation between TGFβ1 treatment and HGF treatment with TGFβ1 was not significantly different (Fig. 2E).
As shown in Fig 2D, DAPI counts in regular area made no difference between TGFβ1 and HGF treatment, including control.
We next examined expression of HGF receptor c-Met in primary astrocyte. RT-PCR showed that c-Met mRNA is present in primary astrocytes and its expression is greatly increased after TGFβ1 treatment (Fig. 3A). HGF treatment did not change expression of c-Met. By immunostaining with antibodies that detect the beta-chain of c-Met (Fig. 3B-D), we confirmed the expression of c-Met in HGF treated astrocytes.
Fig. 1. Astrocytic activation in vitro glial scars model A: Illustration of primary astrocyte cultures and experimental scheme. B: GFAP stained astrocytes in serum free medium. B: 10 ng/ml treatment of TGFβ1 induced astrocytic hypertrophy. C: 5ng/ml treatment of HGF. D-F:5 ng/ml, 50 ng/ml, and 250 ng/ml HGF was added to culture medium with TGFβ1 respectively. Cells were treated with the above conditions for 24 hours and were fixed for immunocytochemistry with GFAP. G: Quantification of GFAP positive areas. *p<0.05,
**p<0.01 and ***p<0.001 by one-way ANOVA followed by Tukey’s post hoc test for the comparison of mean GFAP areas between different groups.
Fig. 2. Absence of HGF effect on astrocyte proliferation A-C: Ki-67 immunocytochemistry and DAPI staining of primary astrocytes without TGFβ1 (A), with TGFβ1 (B), and TGFβ1 with HGF 50 ng/ml (C). D: DAPI counting results by groups. E:
Quantification of Ki-67 positive cells counts among the all strocytes in the one taken picture.
Fig. 3. Expression of HGF receptor, c-Met, in primary astrocytes A: RT-PCR analysis showed mRNA expression of c-Met in primary astrocytes with or without TGFβ1 and HGF contreatment condition. B-D: Images of immunocytochemistry with c-Met and DAPI in primary astrocytes treated with TGFβ1 and HGF 50 ng/ml. B: Immunoreactivity of DAPI. C. Immunoreactivity of c-Met. D: merge image of DAPI and c-Met immunoreactivities.
B. HGF decreases expression of chondroitin sulfate proteoglycans (CSPGs)
To examine whether HGF affects production of CSPGs that is a major component of glial scars, we measured mRNA expression of neurocan and phosphacan, which are two species of CSPGs upregulated following cytokine stimulation (McKeon et al., 1999; Asher et al., 2000) and spinal cord injury (Jones and Tuszynski, 2002). TGFβ1 induced astrocytic activation resulted in dramatic increases in neurocan and phosphacan mRNAs.
Quantification data showed that neurocan mRNA expression was elevated 8 times higher than control level. HGF treatment at a concentration of 50 ng/ml almost completely blocked the increase of neurocan mRNA expression (Fig. 4A, B). Phosphacan mRNA level was also increased 2 times with TGFβ1, and HGF co-treatment evidently reduce the expression level (Fig. 4C, D). These experiments confirmed that HGF obviously suppresses mRNA production of CSPGs.
Fig. 4. Modulation of chondroitin sulfate proteoglycans (CSPGs) mRNA expression by HGF A: Quantification of mRNA expression of neurocan. B: neurocan RNA analysis by RT-PCR. C: Quantification of mRNA expression of phosphacan D: phosphacan RNA analysis by RT-PCR. *p<0.05, **p<0.01 , and ***p<0.001 by one-way ANOVA followed by Tukey’s post hoc test for the comparison with hypertrophic effects in primary astrocytes.
Table 1. Experimental groups and treatment number in vitro
C. Effects of HGF on TGFβ secretion
Following spinal cord injury, TGFβ1 and TGFβ2 were secreted from reactive astrocytes around the lesion site. TGFβ1 induces inflammatory responses and is involved in initial formation of the glial scars, while TGFβ2 maintains glial scars at later time points (Buss et al., 2008). Inhibition of TGFβ1 and TGFβ2 functions by neutralizing antibodies reduced the extent of scar formation (Logan et al., 1999; Moon and Fawcett, 2001). Therefore, TGFβs
Following spinal cord injury, TGFβ1 and TGFβ2 were secreted from reactive astrocytes around the lesion site. TGFβ1 induces inflammatory responses and is involved in initial formation of the glial scars, while TGFβ2 maintains glial scars at later time points (Buss et al., 2008). Inhibition of TGFβ1 and TGFβ2 functions by neutralizing antibodies reduced the extent of scar formation (Logan et al., 1999; Moon and Fawcett, 2001). Therefore, TGFβs