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First published online 8 May 2007
doi: 10.1242/jcs.03448
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Research Article |
Department of Neurobiology, Second Military Medical University, Shanghai 200433, China
Author for correspondence (e-mail: chenghe{at}online.sh.cn)
Accepted 28 March 2007
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Summary |
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 Top Summary IntroductionResultsDiscussionMaterials and MethodsReferences |
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Key words: Nogo, Olfactory ensheathing cells, Migration, Adhesion, RhoA, Spinal cord injury
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Introduction |
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TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
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; Doucette et al., 1983). Olfactory ensheathing cells (OECs) are the glial cells that derive from the olfactory placode and envelop olfactory axons in the course of migration from the olfactory epithelium to the bulb (Ramon-Cueto and Avila, 1998). The cells are different from the typical glia in terms of existing in both the peripheral nervous system and CNS and sharing the phenotypes of both astrocytes and Schwann cells (Ramon-Cueto and Avila, 1998; Gudino-Cabrera and Nieto-Sampedro, 2000). OECs have been reported to pioneer the olfactory nerve pathway and provide a conducive substrate for growing primary olfactory axons, although the specific mechanisms remain undescribed (Tennent and Chuah, 1996).
The repair of CNS damage continues to be a major challenge, in particular that of spinal cord injury. Owing to the axonal growth-promoting properties, OEC transplantation has emerged as a very promising experimental therapy to treat axonal injuries (Franklin and Barnett, 2000; Raisman, 2001; Ramon-Cueto and Santos-Benito, 2001). Transplanted OECs have been shown to migrate with regenerating axons through an unfavorable CNS environment (Li et al., 2004), and to mingle well with astrocytes in adult brain (Li et al., 1998; Lakatos et al., 2000; Richter et al., 2005). Therefore, the migrating ability of OECs in the CNS was thought to be essential for neural regeneration and re-ensheathment after spinal cord injury. However, whether transplanted OECs can migrate over long distances in the CNS is still controversial (Smale et al., 1996; Gudino-Cabrera et al., 2000; Takami et al., 2002; Collazos-Castro et al., 2005; Deng et al., 2006).
The regeneration of CNS axons following injury is drastically restricted by the presence of inhibitory molecules within myelin (Schwab and Bartholdi, 1996; Ng et al., 1996). Three inhibitors that have been identified are Nogo, myelin-associated glycoprotein (MAG) and oligodendrocyte myelin glycoprotein (OMgp) (He and Koprivica, 2004). Nogo is a member of the reticulon family and occurs in three forms, Nogo-A, Nogo-B and Nogo-C, which are generated from alternate splicing (GrandPre et al., 2000). All three isoforms of Nogo share a 66-amino-acid-residue luminal/extracellular domain (Nogo-66), which inhibits axonal extension and fibroblast spreading (Brittis and Flanagan, 2001; Fournier et al., 2001). The molecular cloning of Nogo (Chen et al., 2000; GrandPre et al., 2000; Prinjha et al., 2000) led to the identification of a neuronal surface glycosyl phosphatidylinositol (GPI)-linked receptor that binds with high affinity to Nogo-66, termed the Nogo-66 receptor (NgR) (Fournier et al., 2001). The low affinity neurotrophin receptor, p75, has been identified as a coreceptor for NgR, and transduces a signal upon interaction with myelin ligands (Wang et al., 2002; Wong et al., 2002). The downstream signaling pathway involves the activation of small GTPases of the Rho family, which in turn regulate cytoskeletal protein assembly and mediate inhibitory effects on neurite growth (Yamashita et al., 2002; Yiu and He, 2003). The mRNA for NgR has been found in abundance in OECs (Woodhall et al., 2003), and the NgR coreceptor p75 is well known as a marker protein for OECs. However, little is known about the function of the receptors in OECs.
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), as well as an essential player in modulating axon-glial junction architecture and possibly K+-channel localization during development (Nie et al., 2003). Nogo-B, like other members of the reticulon family, is involved in modulating BACE1 activity and amyloid-
peptide generation (He et al., 2004), and might function as a pro-apoptotic protein (Tagami et al., 2000). Moreover, Nogo-B has been reported as a regulator of vascular remodeling (Acevedo et al., 2004). In the present study, we demonstrate that Nogo enhances the adhesion and inhibits the migration of OECs via RhoA activation by NgR. |
Results |
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TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
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Nogo-66 enhances the adhesion of OECs
To determine the effect of Nogo-66 on OEC adhesion, a protein-spot assay was used. We found that OECs had a significantly greater affinity for attaching to spots harbouring fusion proteins (GST–Nogo-66 or His–Nogo-66) than those containing GST or His alone (Fig. 2A). Quantification of the results showed that the numbers of OECs attaching to GST–Nogo-66 and His–Nogo-66 spots after various time points were different (Fig. 2B). The enhancement of adhesion by either GST–Nogo-66 or His–Nogo-66 was time-dependent within the first 6 hours after plating. In addition, the enhancement of OEC adhesion was dependent on the concentration of GST–Nogo-66 or His–Nogo-66 in the spot. As shown in Fig. 2C, the number of cells attached to the spotted GST–Nogo-66 or His–Nogo-66, but not to the spotted GST or His, increased with the concentration of proteins. Taken together, these results suggest that Nogo-66 has a promoting effect on the adhesion of OECs.
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NgR is a protein anchored to the membrane via a GPI linkage that can be released by PI-PLC (Fournier et al., 2001). As shown in Fig. 3C, the immunostaining of NgR was greatly reduced after treatment of OECs with PI-PLC. The results were further confirmed by western blotting. After treatment of cells with PI-PLC, NgR was released from a membrane-bound fraction to a supernatant fraction. Correspondingly, the residual amount of NgR remaining on the surface of OECs was significantly decreased. Without PI-PLC treatment, immunoreactivity for NgR was undetectable in the supernatant fraction and had no change in the membrane fraction (Fig. 3D).
NgR mediates the effect of Nogo-66 on the adhesion and migration of OECs
After confirming that endogenous NgR was expressed on OECs, we next tested if Nogo-66 was bringing about its effect on OECs through the NgR. Firstly, we investigated whether treatment of PI-PLC could attenuate the effect of Nogo-66 mediating the enhancement of cell adhesion and inhibition of cell migration. As shown in Fig. 4, the effect of Nogo-66 on the migration (Fig. 4A) and adhesion (Fig. 4B) of OECs was greatly attenuated by PI-PLC treatment in a dose-dependent manner (Fig. 4C). Secondly, pre-incubation of OECs with anti-NgR antibody which had been reported to be a function blocking antibody (Domeniconi et al., 2002), but not an irrelevant IgG, depressed the ability of Nogo-66 to inhibit the motility (Fig. 4A) and enhance the adhesion of OECs (Fig. 4B). Importantly, the characteristic of migration (Fig. 4A) and adhesion (Fig. 4B) of SCs in which endogenous NgR is undetectable was not changed by GST–Nogo-66 or His–Nogo-66. These results strongly suggested that the enhancement of OECs substratum adherence and inhibition of OECs migration mediated by Nogo-66 were dependent on the NgR expressed on the plasma membrane of these cells.
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). In addition, the Rho family GTPases participates in regulation of the actin cytoskeleton and various cell adhesion events (Van Aelst and D'Souza-Schorey, 1997; Hall, 1998). Therefore, we next tested whether RhoA was involved in the intracellular signaling of Nogo-66 on OECs. In the adhesion assay, OECs were pretreated with 10 µM Y-27632, a selective inhibitor of RhoA-associated kinase. As shown in Fig. 5A,B, pretreating OECs with Y-27632 markedly reduced the number of adherent cells on GST–Nogo-66 and His–Nogo-66. In the migration assay, inhibition of OEC motility by Nogo-66 was greatly attenuated after pre-treatment of cells with Y-27632 (Fig. 5C). To further examine whether Nogo activates RhoA in OECs, the pull-down assays with the Rho binding domain of rhotekin were performed using OECs seeded on Nogo-66 substrates. As shown in Fig. 5D, the amount of GTP-bound RhoA was significantly increased in OECs plated on GST–Nogo-66 or His–Nogo-66 substrates. All these results suggest that RhoA activation is critical for mediating the effect of Nogo-66 on OECs.
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Focal adhesions are at the termini of stress fibre bundles that serve in longer-term anchorage (Burridge et al., 1998), and RhoA is a central player in the formation of focal adhesions and actin stress fibre bundles (Nobes and Hall, 1995; Allen et al., 1997). To determine further whether the effect of Nogo-66 on OECs is associated with RhoA activity, OECs and Y-27632-treated OECs were double-stained for paxillin and F-actin after cells were plated on GST, GST–Nogo-66, His, or His–Nogo-66. In the OECs on GST and His, there was little staining for the focal adhesion protein paxillin (Fig. 5Eb,n). These cells revealed multiple membrane protrusions where there were thick bundles of actin filaments (Fig. 5Ec,o). In OECs plated on GST–Nogo-66 or His–Nogo-66, filamentous actin extended along the base of the cell and organized in fine bundles parallel to the cell cortex at the cell periphery (Fig. 5Eg,s). In these cells, additionally, paxillin was distributed in dense basal plaques at the peripheral focal adhesions (Fig. 5Ef,r), coinciding with the insertion points of actin filaments (Fig. 5Eh,t). Quantitative analysis revealed that the number of paxillin punctae of OECs on GST–Nogo-66 or His–Nogo-66 substrates was significantly increased (Fig. 5F). When OECs were pretreated with Y-27632 and plated on GST–Nogo-66 (Fig. 5Ei-l) or His–Nogo-66 (Fig. 5Eu-x), they exhibited a similar staining pattern and membrane protrusion to the untreated OECs plated on GST or His. The number of paxillin punctae was significantly decreased in pretreated OECs plated on Nogo-66 fusion proteins (Fig. 5F). These results demonstrated that Nogo-66, via activation of RhoA, promoted the formation of OEC focal adhesions and inhibited their membrane protrusion.
Nogo-A is involved in the augmentation of OEC adhesion and restriction of OEC migration by oligodendrocytes
As shown in Fig. 6A, cultured oligodendrocytes (OLs) exhibited positive immunostaining for Nogo-A and OL-specific markers O1 and MBP. Nogo-A was primarily localized to the cell body and major processes. To test whether endogenous Nogo-A expressed by OLs influenced OEC adhesion, OECs were labeled with Di-I (3,3'-dioctadecyloxacarbocyanine perchlorate) and applied to an adhesion assay. As shown in Fig. 6B, OECs had a significantly greater affinity for remaining attached to an OL monolayer than to laminin, indicating that OLs may express or secrete some molecule that enhances the adhesion of OECs. To test whether OLs expressing endogenous Nogo-A also inhibit the migration of OECs, a standard inverted coverslip migration assay was carried out. When inverted on GST–Nogo-66, His–Nogo-66 and OL monolayers, few OECs migrated away from the coverslip fragments, in terms of both the distances covered by OECs (Fig. 6C,D) and the number of cells emerging from the fragment (Fig. 6C,E). By contrast, OECs migrated further on laminin, GST and His controls. These results suggest that OECs migrate poorly on OL monolayers and the enhanced adhesion to OL monolayers may contribute to the decreased number of OECs migrating over OLs monolayer.
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The migration of implanted OECs is facilitated by neutralizing NgR in vivo
To investigate whether the migration of implanted OECs was affected by Nogo, an in vivo migration assay was performed as previously reported (Cao et al., 2006). OECs prelabeled with Di-I were injected into the injured spinal cord. Ten days after injection, OECs were found to migrate longitudinally and laterally from the injection sites. However, in the presence of anti-NgR antibody, OECs migrated further in the vertical direction (parallel with the long axis of spinal cord) when compared with OEC migration using normal saline (N.S.) or the irrelevant IgG, at the rostral (Fig. 8) as well as the caudal injecton site. There was no difference in migration distance between OECs injected at rostral and caudal sites (data not shown). Quantitative analysis revealed that treatment of OECs with anti-NgR increased the maximum migration distance along the vertical direction, and there was no significant difference in the maximum migration distance along the horizontal direction (Fig. 8B). These results indicated that neutralizing NgR with anti-NgR antibody facilitated OEC migration through white matter tracts in vivo.
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Discussion |
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TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
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). The ability of OECs to assist in the growth of olfactory axons in the olfactory system has led them to become compelling candidates for transplant-mediated repair of CNS lesions. Several studies have now confirmed the use of OECs in spinal cord injuries, which include the promotion of axonal regeneration following spinal cord injuries and the replacement of myelin in demyelinating diseases. The anatomical evidence of regeneration (Franklin et al., 1996; Li et al., 1997; Li et al., 1998; Imaizumi et al., 2000; Nash et al., 2002) and functional improvements (Li et al., 1998; Ramon-Cueto et al., 1998; Ramon-Cueto et al., 2000; Lu et al., 2001; Lu et al., 2002; López-Vales et al., 2006) have been noted in a variety of spinal cord repair models, including complete transection, hemisection, tract lesion, contusion and demyelination. However, there are some negative reports of OEC transplantation after spinal cord injury (Resnick et al., 2003; Barnett and Riddell, 2004), and it is still debatable whether transplanted OECs can migrate long distances in the injured CNS. Smale et al. reported that no significant cell migration was detected when OECs from fetal rat olfactory bulb were implanted into the damaged adult rat brain (Smale et al., 1996). The recent study by Takami et al. showed that grafted OECs simply disappeared from the lesion cavity with no evidence that they had migrated away into the surrounding neuropil (Takami et al., 2002). Deng and colleagues reported that both rat and human OECs showed similar migration after injection into the thoracic spinal cord. Importantly, both rat and human OECs migrated for shorter distances, in both rostral and caudal directions, in the injured cord of animals with a concomitant contralateral hemisection (Deng et al., 2006). Data from other studies also demonstrated that adult rat or human OECs migrated over shorter distances after they were transplanted into the injuried CNS (Gudino-Cabrera et al., 2000; Collazos-Castro et al., 2005). However, the underlying specific mechanisms for the poor motility of OECs in the damaged CNS remain unknown. In the present study, we first show that both Nogo-66 and Nogo-A enhance the adhesion of OECs and inhibit their migration. These studies may, at least partially, explain why OECs fail to migrate long distances in the injured CNS.
It is well documented that Nogo, MAG and OMgp are prominent components of CNS myelin inhibitory activity for adult axon-regeneration. Following injury to the spinal cord, Nogo-A mRNA is upregulated around the lesion and Nogo-A protein is strongly expressed in injured dorsal column fibres and their sprouts that entered the lesion site (Hunt et al., 2003). Meier et al. showed that Nogo-A expression is upregulated after hippocampal denervation or kainate-induced seizures (Meier et al., 2003). Studies of Bandtlow et al. (Bandtlow et al., 2004) demonstrated that Nogo-A mRNA and immunoreactivity are markedly increased in hippocampal neurons of patients with temporal lobe epilepsy. In the present study, Nogo-A was found highly expressed in OLs, which is consistent with previous reports (Moreira et al., 1999; GrandPre et al., 2000; Huber et al., 2002). Furthermore, the co-culture migration assay demonstrated that motility of OECs was significantly depressed by Nogo-A on OLs. Therefore, the elevated levels of Nogo-A, probably on OLs, in the injured CNS, may be responsible for the inhibition of the migration of transplanted OECs in the damaged CNS.
The movement of a metazoan cell involves the turnover of adhesions with the substratum on which it moves (Kaverina et al., 2002). In the present study, we found that the adhesion of OECs was enhanced both by recombinant Nogo-66 protein and by Nogo-A exogenously expressed by Cos-7 cells or endogenously by OLs. Therefore, one possibility to account for the influence of Nogo-66 on OEC migration was that Nogo-66 affected the adhesion of OECs. Adhesion sites form as a result of signaling between the extracellular matrix on the outside and the actin cytoskeleton on the inside, and they are associated with specific assemblies of actin filaments (Kaibuchi et al., 1999; Kaverina et al., 2002). Paxillin is a cytoskeletal component localized to the long-lived focal adhesions at the end of actin stress fibres that serve in longer-term anchorage (Burridge et al., 1998). Protrusion of the cell surface is an early step in several cellular processes including cell migration. The driving force for the formation and extension of membrane protrusions is the polymerization of actin (DeMali and Burridge, 2003). In the present study, we observed the influences of Nogo-66 on focal adhesions, actin stress fibers and membrane protrusions. It was apparent that Nogo-66 regulated actin reassembly, promoted the formation of focal adhesions, and inhibited membrane protrusion. Taken together, these data support the hypothesis that Nogo-66 enhances the adhesion of OECs, which inhibits their migration.
Woodhall et al. reported that NgR mRNA is expressed in primary cultures of OECs (Woodhall et al., 2003). In our study, the expression of NgR in OECs was confirmed further by immunocytochemistry and western blot. With enzymatic cleavage of NgR and anti-NgR block, we confirmed that the effect of recombinant Nogo-66 and endogenous Nogo-A expressed by OLs on OECs was NgR-dependent. To our knowledge, this is the first evidence that functional NgR is expressed on OECs, although the role of p75, the common marker for OECs, as an essential co-receptor needs further investigation. The Rho family of small GTPases have been reported to be central players in regulating the assembly of the actin cytoskeleton, adhesion formation and membrane protrusion (Hall, 1998; Ridley, 2001; DeMali and Burridge, 2003), in which RhoA signals the formation and maturation of focal adhesions associated with actin stress fibre bundles, and Rac1 and Cdc42 stimulate the formation of protrusions in association with lamellipodia and filopodia, respectively. Formation of membrane protrusions is an early central step in the process of cell migration. When the ratio of activity between these small GTPases is no longer optimal for protrusion and polarization of the cell, the migration will stop (Cox et al., 2001). In the present study, we found that active RhoA was significantly increased in OECs plated on Nogo-66 substrates, which suggests that Nogo binding to NgR leads to activation of RhoA in OECs. One of the downstream effectors of RhoA is Rho-kinase (ROCK), which can be inhibited effectively by the compound Y-27632 (Narumiya et al., 2000). Inhibition of ROCK stimulates membrane protrusion (Cox et al., 2001; Rottner et al., 1999; Tsuji et al., 2002) and promotes cell migration (Nobes and Hall, 1999). Here, we observed that the effect of Nogo-66 on the adhesion and migration of OECs was greatly attenuated when cells were pretreated with Y-27632. In addition, we found that the Nogo-66-induced increase of focal adhesion formation, alteration of actin cytoskeletal structure, and inhibition of membrane protrusion were significantly attenuated by pretreatment of OECs with Y-27632. All these results supported the hypothesis that the activity of RhoA is involved in the regulation of adhesion and migration of OECs by Nogo-66. However, further study is necessary to clarify the detailed intracellular signaling mechanisms.
It is a continuing challenge to move toward therapeutic approaches for CNS injury. Recently, there has been great interest in the possibility that OECs have potential for use in the treatment of axonal injuries and demyelinating disease (Franklin and Barnett, 2000; Raisman, 2001; Ramon-Cueto and Santos-Benito, 2001). Upgrading the growth-promoting properties of OECs is considered a valuable strategy for promoting CNS repair; however, most of these studies involve OECs secreting additional neurotrophic factors (Cao et al., 2004; Ruitenberg et al., 2003; Ruitenberg et al., 2005). Since the inhibition of axonal outgrowth by CNS myelin is one of the major obstacles to functional recovery following CNS injury, myelin inhibitors and their receptors are recently emerging as potential therapeutic targets (Lee et al., 2003). Neutralizing Nogo-A with IN-1 antibody and NgR antagonists were proven to improve CNS axon regeneration and functional recovery after various lesions (Fouad et al., 2004; GrandPre et al., 2002). In the present study, we found that OEC migration was facilitated in vitro and in vivo when NgR was neutralized with anti-NgR antibody. It is of interest to test the idea that, combined with OEC implantation, local administration of Nogo-A antibody or NgR antagonists would improve the therapeutic properties of OECs on CNS injury.
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Materials and Methods |
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TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
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). Briefly, OECs were extracted from the olfactory nerve layer by trypsin treatment and plated on two uncoated 25 cm2 culture flasks; each was incubated for 36 hours at 37°C in 5% CO2. The non-adhesive cell suspension was collected and then seeded onto disks pre-coated with poly-L-lysine (PLL, 0.1 mg/ml), and incubated with serum-containing DMEM/F-12 supplemented with 2 µM forskolin (Sigma) and 10 ng/ml bFGF (Sigma).
Schwann cells (SCs) were obtained from sciatic nerves of 2-day-old Sprague-Dawley rat pups and purified using a modification of the protocol previously described (Lakatos et al., 2000; Brockes et al., 1979). Cells were cultured on poly-L-lysine (100 µg/ml)-coated dishes and maintained in DMEM/F-12 containing 15% FBS and supplemented with Forskolin (2 µM, Sigma) and bFGF (10 ng/ml, Sigma). The cultures were treated with cytosine arabinoside (10–5 M, Sigma) to reduce contamination by fibroblasts.
Oligodendrocytes (OLs) were prepared from rat cerebral cortex as described (McCarthy and de Vellis, 1980; Fok-Seang et al., 1995). In brief, newborn rat cortical cells were dissociated and cultured in DMEM/F-12 containing 10% FBS. The culture medium was changed at 24 hours and twice weekly thereafter. After 10-12 days, the cultures became confluent and loosely attached macrophages were removed by shaking the flasks on a rotary shaker at 260 rpm for 1 hour at 37°C. The supernatant was discarded and the cultures were then shaken in fresh culture medium for 18 hours at 260 rpm to remove oligodendrocyte precursor cells from the cell monolayer. The detached cells were plated onto a non-coated culture dish for 7 minutes to remove adherent cells such as microglia and astrocytes. The oligodendrocyte precursor cells were then transferred to a culture dish pre-coated with PLL and left to adhere overnight. From the next day on, the medium was exchanged for serum-free chemical defined DMEM/F-12 supplemented with 5 µg/ml insulin, 50 µg/ml transferrin, 0.66 mg/ml BSA, 20 ng/ml progesterone, 100 µmol/ml putrescine, 40 ng/ml sodium selenite and 30 nmol/ml T3, and the oligodendrocyte precursor cells were cultured for another 1-10 days to differentiate into OLs. OLs were identified by indirect immunofluorescence labeling using monoclonal anti-oligodendrocyte marker O1 and anti-myelin basic protein (MBP).
Unless indicated otherwise, cells were pretreated with or without PI-PLC (0-0.1 U/ml), anti-NgR (1:100), IgG (1:100) or Y-27632 (15 µM) for 2 hours before contact with Nogo-66. For PI-PLC treatment, cells were washed once with 0.1 M PBS (PH 7.4) and then incubated with phosphatidylinositol-specific phospholipase C (PI-PLC, Sigma; 0.001, 0.01, 0.1 U/ml) in DMEM/F-12 at 37°C for the times indicated. The cells were washed three times and then processed for the migration and adhesion assay or for western blot analysis. To confirm cell viability, cells were stained with 0.4% Trypan Blue. Trypan-Blue-incorporating cells were <1% in each experiment.
Fluorescent staining
For immunocytochemical analysis, immunocytochemistry of cells cultured on coverslips were performed as previously described (Yan et al., 2003). Briefly, coverslips were fixed with 4% paraformaldehyde in PBS for 20 minutes, then permeabilized with or without 0.3% Triton X-100 in 0.1 M PBS for 15 minutes and incubated overnight at 4°C with polyclonal antibodies to NgR (Santa Cruz), S-100 (Boster) or Nogo-A (Santa Cruz), or monoclonal antibodies to paxillin (BD Transduction Laboratories), O1 (R&D Systems) or MBP (myelin basic protein, Chemicon international) diluted in PBS containing 10% normal goat serum. After washing three times with 0.1 M PBS (pH 7.4), cells were incubated with fluorescence-conjugated secondary antibody (Sigma) for 90 minutes at room temperature. For visualization of F-actin, cells were incubated with 0.1 µg/ml TRITC-phalloidin (Sigma) overnight at 4°C. Cells were then examined by fluorescence microscopy.
Boyden chamber migration assay
To measure the ability of OECs to migrate, OEC migration studies were performed using a 24-well Boyden chamber (Costar) containing polycarbonate membranes (8 µm pore size), with slight modification of a previously described protocol (Klemke et al., 1997; Yan and Rivkees, 2002; Yamauchi et al., 2003). In brief, the undersides of polyethylene terephthalate filter membranes were coated with laminin or fusion proteins (100 ng/ml diluted in laminin) overnight at 4°C. Cells were detached by trypsin/EDTA and then seeded onto the upper chamber at a density of 4 x105 cells in 250 µl of culture medium containing 1% serum per well. The upper chambers were inserted into the tissue-culture wells and 750 µl culture medium containing 1% serum was added to the lower chambers. After incubation for 8 hours at 37°C, non-migratory cells on the upper membrane surface were removed with a cotton swab, and migratory cells migrating through the membrane pores and invading to the underside surface of the membrane were fixed with 4% paraformaldehyde and stained with Coomassie Brilliant Blue. For quantitative assessment, the number of stained, migrating cells was then counted under a microscope at five fields per filter in three independent experiments.
Inverted coverslip migration assay
The inverted migration assay was performed according to a previous protocol (Fok-Seang et al., 1995). To prelabel SCs or OECs, cells were incubated with 25 µg/ml Di-I (Molecular Probes, a sulfonated carbocyanine fluorescent tracer: 3,3'-dioctadecyloxacarbocyanine perchlorate) for 5 min at 37°C. Coverslips (8 mm) were broken to produce an approximately 1x2 mm fragment. Di-I-labeled or nonlabeled cells were plated onto the coverslip fragments precoated with PLL. After the cells had been allowed to attach for 16-18 hours, the coverslips were washed with culture medium three times to remove any loose cells and then inverted so that cells faced downward onto tissue culture surfaces coated with laminin, GST, GST–Nogo-66, His or His–Nogo-66, or onto cell monolayers or extracellular matrix. These cultures were then incubated for a further 3 days to allow cell migration from the edge of the inverted fragment and fixed with 4% paraformaldehyde. The maximum migration distance from the edge of each of the coverslip fragments was measured and the number of cells present within rows (50 µm x 1 mm area) progressing outward from the edge of the coverslip fragment was counted. Experiments were carried out in the presence of 12 µg/ml aphidicolin (Sigma), a mitotic inhibitor, in order to be certain that movement away from the coverslip fragment was due to migration alone and did not contribute to proliferation.
Preparation of culture conditioned medium and matrix
Oligodendrocyte-conditioned serum-free medium was obtained by growing confluent OLs for 2 days in condition-defined medium that consisted of DMEM/F-12 supplemented with 1% N2 (vol/vol), 10.1 ng/ml T3, 400 ng/ml T4, 0.035% bovine serum albumin (BSA) and 20 µM leupeptin. Oligodendrocyte-conditioned serum-free medium prepared in this way was filtered (0.22 µm filter) and used without further dilution. Extracellular matrix from OLs was obtained from oligodendrocyte monolayers by incubating these cells in distilled water for 2 hours. Cellular debris was removed by several washes with culture medium prior to assay.
Cells adhesion assay
The adherent ability of OECs when contacted with Nogo-66 was detected by two different approaches. OECs or SCs were plated on recombinant GST or His and the respective GST or His fusion proteins (10-100 µg/ml solution in 1 µl-sized spots) that were dried down onto dishes coated with poly-L-lysine (PLL). Cells were incubated at 37°C before fixing with 4% paraformaldehyde after various time points (Dorries et al., 1996). The number of cells attached to GST or His fusion protein spots were then counted under a microscope. To analyze the adhesion of OECs to substrates or cell monolayers, a total of 20,000 Di-I-labeled OECs or SCs were placed onto 24-well plates coated with laminin GST, GST–Nogo-66, His or His–Nogo-66, or a complete monolayer of OLs or Cos-7 cells transfected with Nogo-A. The cultures were incubated for 30 minutes at 37°C on a rotary shaker at 25 rpm. After several washes with culture medium, cultures were fixed in 4% paraformaldehyde for 20 minutes at room temperature and the number of adhering cells was determined by counting the number of stained cells in a counting grid under a 20x objective and repeating the counts in adjacent grids to cover one complete diameter of each well.
Western blot analysis
To detect the expression of NgR, OECs, SCs (as negative control) and PC12 cells (as positive control) were harvested after being rinsed briefly with ice-cold PBS and lysed in SDS gel sample buffer. To confirm the ability of PI-PLC to release NgR from the membrane, the supernatant and membrane fractions of OECs were collected after cells were treated with 0.1 U/ml PI-PLC. The samples were denatured by boiling for 10 minutes with SDS gel sample buffer, then centrifuged for 10 minutes at 12,000 g at 4°C. Proteins in the supernatants were separated by 12% SDS-polyacrylamide gel and then transferred onto nitrocellulose membranes. Membranes were then blocked with 10% low-fat milk in 1x TBST and incubated with specific primary antibody against NgR (Chemicon). To control for differences in protein loading, membranes were also incubated with anti-GAPDH antibody (Sigma). After incubating with horseradish peroxidase (HRP)-conjugated secondary antibodies (Sigma; 1:10,000), immunoreactive bands were visualized by chemiluminescence reagents (ECL, Amersham).
RhoA activity assay
Active RhoA was determined with the GST-Rhotekin-binding domain as described previously (Ren et al., 1999). OECs were plated onto the dishes coated with Nogo-66 fusion proteins (GST–Nogo-66 or His–Nogo-66) or control proteins (GST or His) for 6 hours. Cells were washed with ice-cold PBS and lysed in RIPA buffer (50 mM Tris, pH 7.2, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 500 mM NaCl, 10 mM MgCl2, 10 µg/ml each of leupeptin and aprotinin, 1 mM phenylmethylsulfonyl fluoride). Cell lysates were clarified by centrifugation at 12,000 rpm at 4°C for 5 minutes, and equal volumes of lysates were incubated with GST-Rhotekin-binding domain (20 µg) bound to beads at 4°C for 60 minutes. The beads were then washed four times in buffer B (Tris buffer containing 1% Triton X-100, 150 mM NaCl, 10 mM MgCl2, 10 µg/ml each of leupeptin and aprotinin, and 0.1 mM phenylmethylsulfonyl fluoride) at 4°C. Bound RhoA proteins were detected by western blotting using a monoclonal antibody against RhoA (Santa Cruz, 1:500). The amount of GTP-bound RhoA was normalized to the total amount of RhoA in cell lysates as previously described (Ren et al., 1999), and statistical analysis was performed for the results of three independent experiments.
In vivo migration assay
To investigate whether Nogo affected the migration of implanted OECs, an in vivo migration assay was performed as previously described (Cao et al., 2006). Animal care and use followed recommended NIH guidelines. Female adult Sprague-Dawley rats (200-250 g) were anaesthetized with 2% pentobarbital sodium. Laminectomy was performed to expose the dorsal surface of the T7-9 segment, followed by a hemisection at T8 using scalpels. A 1 mm segment was removed on the left side of the T8 spinal cord. Animals were divided into three experimental groups: (1) Normal saline (N.S.) group (n=6), (2) anti-NgR group (n=8) and (3) IgG group (n=7). Di-I-labeled OECs were deposited into two injection sites at the rostral as well as the caudal stump, + mm from the lesion cavity, using a sterile glass needle. A volume of 0.5 µl containing 5x105 cells in DMEM was grafted into each site. Rats in group 1 were injected with untreated OECs and received N.S. (10 µl/day) through a pipe embedded in the subarachnoid space. In group 2, rats were injected with OECs pretreated with anti-NgR antibody (Santa Cruz, 1:100) for 30 minutes and received the antibody (10 µl/day) instead of N.S. In group 3, rats were injected with OECs pretreated with an irrelevant goat IgG (1:100) for 30 minutes and received the IgG (10 µl/day).
Ten days after injury, the experimental animals were perfused and the fixed spinal cords (T7-T9) were post-fixed for 8 hours in 4% PFA. A series of consecutive sagittal sections (8 µm) were cut and collected in PBS. To quantify the motility of the implanted OECs in the spinal cord, we selected five representative midsagittal sections in each animal and measured the size of the region invaded by Di-I-labeled cells from the graft site under a fluorescence microscope (IX70, Olympus). To describe the motorial characteristic of grafted OECs in detail, the area covered by grafted OECs (area) and the maximum migration distance along the vertical (length) or horizontal (width) direction were included in the parameters measured.
Statistical analysis
All data present represent the results of at least three independent experiments, using cells prepared at different times. Statistical analysis was performed using unpaired Student's t-test. All data are presented as mean ± s.d.
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