Neuritin Mediates Nerve Growth Factor–Induced Axonal Regeneration and Is Deficient in Experimental Diabetic Neuropathy Free

All experiments were conducted in accordance with U.K. Home Office Regulations and with the Animals Act (Scientific Procedures), 1986. Male Wistar rats (250–300 g on arrival, Charles River, Kent, U.K.) were used in all studies. Diabetes was induced by a single intraperitoneal injection of STZ (Sigma, St. Louis, MO), freshly dissolved in normal saline, at a dose of 55 mg/kg, administered the morning after an overnight fast. Three days later, blood glucose was monitored (BM Acutest strips; Roche Diagnostics, Mannheim, Germany), and any STZ-administered rats with blood glucose levels <15 mmol/l were rejected from the study. Animals were then group housed with full access to food and water for 12 weeks.

Gene expression in the DRG was compared between two groups of rats: one control group and one after 12 weeks of diabetes. DRGs were dissected out from spinal segments L4 and L5, and the four ganglia were pooled from each rat. Ganglia from groups of three rats were further pooled to give enough RNA for purification, derivatization, and hybridization. Pooled ganglia were extracted and the RNA processed and hybridized to Affymetrix U34 rat microarray chips exactly as described elsewhere (28).

Rats were randomly assigned to four groups (10 rats per group), designated as follows: control untreated, untreated diabetic, diabetic given 0.1 mg/kg NGF, and diabetic given 0.5 mg/kg NGF. NGF treatment began 8 weeks after induction of diabetes and was maintained for 4 weeks until the rats were killed. Purified human recombinant NGF (GenenTech, San Francisco, CA) was given three times per week by subcutaneous injection at the back of the neck. At the end of 12 weeks, rats were killed and L4/5 DRG and sciatic nerve removed and processed for quantitative PCR or Western blotting.

To study axonal transport control, diabetic rats (as treated in study 2) were anesthetized with isofluorane (2% in oxygen), and, under sterile conditions, the left sciatic nerve was exposed at mid-thigh level. The nerve was ligated using prolene sutures (Ethicon; Johnson & Johnson, Brussels, Belgium), the wound was closed, and animals recovered under observation (29). Twenty-four hours later, animals were deeply anesthetized with sodium pentobarbitol (Sagatal, 60 mg/kg, i.p.) and perfused transcardially with 0.9% heparinized saline (0.9% NaCl, 50 units/ml heparin) followed by 4% paraformaldehyde in 0.1 mol/l phosphate buffer. Sciatic nerves were removed, postfixed for 2 h at 4°C, and then cryoprotected in 30% sucrose (in 0.1 mol/l phosphate buffer overnight at 4°C). Nerves were frozen in optimal cutting temperature embedding medium (VWR, Leicestershire, U.K.) on liquid nitrogen and stored at −80°C until processing. Alternatively, animals were humanely killed, and 5-mm segments were cut from the constricted sciatic nerve—two segments were taken proximal and two distal to the ligature, and anatomically matching segments were cut from the contralateral intact nerve. Samples were frozen on liquid nitrogen before analysis.

Adult male Wistar rats (250 g, Charles River) were killed by concussion followed by decapitation. Dissociated sensory neurons were prepared as previously described (30). Neurons were suspended in modified Bottenstein and Sato's medium (BSM) (containing 0.1 mg/ml transferrin, 20 nmol/l progesterone, 100 μmol/l putrescine, 30 nmol/l sodium selenite, and 1 μg/ml BSA, 0.01 mmol/l cytosine arabinoside and 10 pmol/l insulin in Ham's F12). Neurons were seeded onto either Lab-Tek chamber slides (Nunc; VWR) or 35-mm Petri dishes coated with 2 μg/ml laminin (L2020; Sigma). Neurons were plated for 1 h before treatment with NGF (Sigma; 0.1, 1, or 10 ng/ml in BSM) or BSM alone as a control for 24 h in 5% CO₂ at 37°C. For studies using functional inhibitors of mitogen-activated protein kinase/ERK (MEK-1/2) (U0126, 10 μmol/l), its control U0124 (10 μmol/l; Calbiochem), and PI-3K (LY294002, 10 μmol/l; Calbiochem), neurons were treated with each inhibitor 1 h following plating and then treated with NGF (10 ng/ml; Sigma) for 24 h at 37°C.

siRNA was introduced into cells using the Amaxa nucleofection system (Amaxa, Gaithersburg, MD). Following dissociation, sensory neurons were suspended in 100 μl rat neuron nucleofector solution containing either a scrambled sequence of siRNA with no homology to any known proteins (50 nmol/l; Ambion) or 50 nmol/l of a combination of three neuritin siRNAs: 5′ GUGCGAUGCAGUCUUUAAGtt 3′, 5′ GGGCUUUUCAGACUGUUUGtt 3′, and 5′ GGCAGCUUAUUCGAACUCUtt 3′.

Neurons were transfected using program 0-03 of the Amaxa nucleofection system (transfection efficiency determined to be 70% using pmaxGFP positive control plasmid [Amaxa]), resuspended in BSM, treated with NGF, and incubated for 48 h at 37°C.

Total RNA was extracted from sensory neurons and measured, and quantities were normalized before being reverse transcribed to cDNA using Moloney-murine leukemia virus reverse transcriptase. Quantitative PCR analysis of neuritin and calcitonin gene-related peptide (CGRP) levels were carried out on samples in triplicates using the SYBR Green Master mix kit (Molecular probes) and the ABI Prism 7000 sequence detection system (Applied Biosystems, Foster City, CA). The primers were designed using Primer Express 1.0 software (Applied Biosystems); primer sequences are available upon request from the authors. Quantitative PCR data are expressed as cycle thresholds rather than normalizing expression to a housekeeping gene in order to prevent errors introduced by choosing an erroneous housekeeping gene whose expression may itself be altered by the experimental conditions (diabetes or neurotrophin treatment).

Samples were homogenized in ice-cold lysis buffer (Tris-HCl, pH 7.4, 1% NP-40, 2 mmol/l sodium orthovanadate, 10 mmol/l sodium fluoride, 2 mmol/l EGTA, 10 mmol/l sodium pyrophosphate, 1 mmol/l phenylmethylsulfonyl fluoride, and protease inhibitor cocktail [Sigma]). Samples (30 μg protein/lane) were separated by SDS-PAGE using 16.5% acrylamide gels. Proteins were transferred to nitrocellulose membranes by semi-dry electrophoresis. Nonspecific binding was blocked by incubation in Tris-buffered saline containing 0.05% Tween-20, 5% casein (Sigma), and 5% BSA (Fraction V, Sigma) for 1 h at room temperature and then incubated overnight at 4°C with anti-neuritin (1:500; R&D systems). Blots were washed and then incubated in horseradish peroxidase–conjugated anti-goat IgG (1:5,000; Cell Signaling) for 1 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence kit and quantified using densitometric analysis (Scion Image software, version alpha 4.0.3.2).

Sensory neurons were fixed with ice-cold 4% paraformaldehyde for 30 min and then washed three times with PBS. Fixed cells were incubated with anti-β (III) tubulin (1:1,000, Sigma), a pan-neuronal marker to label neuronal cell bodies and neurites, or anti-neuritin (1:100, R&D systems) overnight at 4°C. Cells were washed and incubated with fluorescein isothiocyanate (FITC)-conjugated donkey anti-mouse or donkey anti-goat IgG (1:200, Jackson ImmunoResearch), respectively, for 1 h at room temperature. Longitudinal sections (8 μm) of sciatic nerve or DRG were incubated for 1 h at room temperature with 5% donkey serum and 0.01% Triton X-100 in PBS and then in anti-neuritin (1:100, R&D systems) and anti-TrkA (1:1,000; Chemicon) for 24–48 h at 4°C. Sections were washed with PBS and then incubated for 1 h in tetramethylrhodamine isothiocyanate (TRITC)- or biotin-conjugated donkey anti-goat and Cy3-conjugated donkey anti-rabbit IgG (1:400; Jackson ImmunoResearch), followed by 1 h in FITC-conjugated extravidin (1:400; Vector labs). Sections were mounted in Vectorshield containing DAPI (Vector Labs). Immunofluorescence was viewed using a Leica fluorescence microscope and images acquired using a Hamamatsu camera with Wasabi software.

For each experimental condition, images of 30 neurons were acquired from randomly selected fields of view. The mean number of neurite-bearing cells (defined as those with neurites longer than 1.5 times cell body diameter) was calculated from these images. The length of the longest neurite from each cell was calculated using SigmaScan software (SPSS), as was a measure of total neurite outgrowth. A series of concentric circles was overlaid onto an image of each neuron, and the number of times neurites crossed each circle was calculated. This gave us a measure of both total neurite outgrowth (total number of crosspoints) and neurite branching structure (crosspoints related to distance from the cell body). All of these measurements were repeated in at least three independent cultures.

Data are expressed in all graphs as the mean ± SEM. Statistical analysis was conducted using Prizm software using ANOVA followed by Dunnett's post hoc test.

We first established that neuritin was expressed in adult control rat DRG using conventional PCR and Western blotting techniques (data not shown). Using immunocytochemical analysis, we examined the distribution of neuritin immunoreactivity in adult rat lumbar (L4/5) DRG. Neuritin immunoreactivity was clearly localized within the cytoplasm of a population of sensory neurons in the DRG (Fig. 1A). Small-diameter sensory neurons expressed higher cytoplasmic levels of neuritin immunoreactivity (Fig. 1A, arrows) than large-diameter neurons (Fig. 1A, asterisks). Neuritin immunoreactivity colocalized with TrkA, the high-affinity receptor for NGF (Fig. 1B). Immunofluorescence was abolished by omission of the primary antibody, indicating the neuritin immunoreactivity to be primary-antibody specific (data not shown). Neuritin immunoreactivity was also maintained in cultures of dissociated adult rat sensory neurons treated with control media (in the absence of neurotrophic support, Fig. 1C) or with NGF (Fig. 1D). Neuritin immunoreactivity was detected in neuronal cell bodies and in puncta throughout the neurites (Fig. 1D and E).

We used immunohistochemistry to characterize axonal transport of neuritin in the sciatic nerve of control adult rats. Ligation of the sciatic nerve caused an increase in neuritin levels proximal and distal to the ligature site (Fig. 2A). Immunohistochemical analysis showed that neuritin immunoreactivity increased in axons proximal and distal to the ligature site (Fig. 2A and B, arrows) while contralateral, nonligated sciatic nerve showed no neuritin (Fig. 2C). Accumulations were also quantified by Western blotting (see below). Neuritin is therefore transported bidirectionally along axons in the sciatic nerve.

Since neuritin was expressed in predominantly small-diameter sensory neurons, we investigated the effect of NGF on neuritin expression. Quantitative real-time PCR was used to assess neuritin mRNA levels in sensory neurons treated for 24 h with NGF (0.1, 1, and 10 ng/ml). NGF caused a dose-dependent increase in neuritin mRNA (Fig. 3A) and protein (Fig. 3B). Quantification of neuritin protein levels using densitometric analysis showed that these NGF-mediated increases in neuritin were significant compared with control (Fig. 3C).

The observation that neuritin is expressed in adult rat sensory neurons, particularly small-diameter neurons, and that its expression is increased in response to NGF in vitro, indicating a potential role for neuritin in mediating downstream actions of NGF.

To investigate the intracellular signaling pathways responsible for the NGF-mediated upregulation of neuritin, we pretreated dissociated sensory neurons with either a MEK-1/2 inhibitor (U0126), a negative control for U0126 (U0124), or a PI-3K inhibitor (LY294002) before stimulation with NGF (Fig. 4). It can clearly be seen from the photomicrographs that inhibition of either MEK-1/2 or PI-3K abolishes NGF-mediated neurite outgrowth (Fig. 4C and D). Inhibition of TrkA with k252A (10 μmol/l) also reduced neuritin upregulation (data not shown). U0124 had no effect on neurite outgrowth (data not shown). The levels of neuritin in neurons treated with these inhibitors was determined using Western blotting (Fig. 4E) and quantified using densitometric analysis (Fig. 4F). The NGF-mediated upregulation of neuritin was significantly reduced to below control values by U1026 (while the negative control, U1024, had no effect). The PI-3K inhibitor also prevented the NGF-mediated upregulation of neuritin but did not reduce the basal level of neuritin in sensory neurons (Fig. 4E and F).

These results indicate that NGF-mediated neurite outgrowth can be prevented using inhibitors of both MEK-1/2 and PI-3K. These inhibitors prevent the NGF-mediated upregulation of neuritin, a factor we have shown to be crucial in mediating NGF-induced neurite outgrowth.

Since diabetes is associated with a decrease in NGF levels, we hypothesized that neuritin levels may also be compromised in diabetes. We compared neuritin gene expression levels in DRG of control rats with those of rats with 12 weeks of STZ-induced diabetes (nine rats, three chips per condition) and found a significant reduction in mRNA from 2,756 to 1,548 (Affymetrix units, P < 0.05; Affymetrix accession number U88958_at). This decrease in diabetes was also seen at the protein level using Western blot analysis (Fig. 5). This deficit in neuritin was reversed by treatment with NGF (P < 0.02 for the higher NGF dose, Fig. 5). Diabetic rats lost weight and were hyperglycemic (blood glucose: control 6.0 ± 0.6 mmol/l versus diabetic untreated 33.0 ± 3.2 mmol/l). Treatment with NGF did not affect the raised blood glucose in diabetic rats, though the higher NGF dose exacerbated body weight loss (data not shown). NGF was assumed not to attenuate the severity of the diabetic state.

As illustrated in Fig. 2, neuritin is axonally transported in control sciatic nerve. We compared neuritin levels in sections proximal and distal to a ligature (and an equivalent section from contralateral unconstricted nerve) in control and diabetic rats using Western blotting and densitometric analysis (Fig. 6). In control rats the proximal accumulation (P1+P2) was 65.9 ± 3.5 (arbitrary units, mean ± SEM) and distal accumulation (D1+D2) was 34.1 ± 3.5 (Fig. 6A). In untreated diabetic rats the proximal accumulation (13.2 ± 2.5) was significantly reduced (P < 0.001), though the distal accumulation (23.3 ± 7.3) was not (Fig. 6B). NGF treatment of diabetic rats (0.5 mg/kg) increased proximal accumulation (38.5 ± 4.2) such that it was significantly higher (P < 0.003) than that seen in untreated diabetic rats but still significantly lower (P < 0.003) than that seen in controls. Distal accumulation was also increased numerically by NGF (41.7 ± 11.0) such that it did not differ significantly from the equivalent values for the other two groups (Fig. 6C).

To determine the functional role of neuritin, we used a gene silencing approach. Sensory neurons were transfected with siRNA using electroporation with AMAXA Nucleofector (31). Adult rat sensory neurons were transfected with either a cocktail of three neuritin siRNAs or a scrambled sequence siRNA with no homology to any known genes or untransfected to act as control. Neurons were treated for 48 h with NGF (10 ng/ml) or control media alone.

Transfection with a scrambled sequence siRNA did not affect the NGF-induced upregulation of neuritin mRNA (Fig. 7A) or protein (Fig. 7B and C). However, transfection with neuritin siRNA caused a significant reduction in both neuritin message (Fig. 7A) and protein (Fig. 7B and C) to below control neuronal values, indicating the specificity of the siRNA.

To assess the contribution of neuritin to the regenerative response, we used an in vitro culture system of dissociated adult sensory neurons treated with NGF to promote neurite outgrowth (30,32,33). Following 48 h in culture, neurons began to extend neurites in the absence of exogenous neurotrophins (Fig. 8A). Neurons typically extended only a few neurites, which were unbranched and relatively short (316 ± 29 μm; Fig. 8E).

Exogenous NGF produced robust neurite outgrowth, and this outgrowth was not affected by transfection of neurons with scrambled siRNA (Fig. 8B and D–F). A greater number of neurons extended neurites in the presence of NGF compared with control neurons (NGF: 76% vs. control 59% neurons with neurites). Furthermore, neurites were significantly longer (533 ± 36 μm, P < 0.05; Fig. 4B and D) and more complex (Fig. 8B E, and F). In contrast, transfection of neurons with neuritin siRNA reduced the number of neurite-bearing cells to 48% (in the presence of NGF). The length of the longest neurite extended in response to NGF was significantly reduced (281 ± 22 μm), and neurite density, as measured by total crosspoints, was significantly inhibited compared with neurons transfected with the scrambled siRNA (Fig. 8E and F). Silencing neuritin, therefore, reduced neurite outgrowth to a level equivalent to that observed in the absence of exogenous NGF.

Since this inhibition of neurite outgrowth by neuritin siRNA could have reflected simple interference with NGF signaling, we examined the expression of another NGF-responsive gene that is not part of the neurite outgrowth response to the neurotrophin. CGRP is upregulated by NGF both in vitro and in vivo and responds to the neurotrophin in intact (i.e., not regenerating) primary afferent neurons (34). Neither the scrambled siRNA nor the neuritin siRNA had any effect on NGF-mediated upregulation of CGRP mRNA (Fig. 7C), confirming that the knockdown of neuritin by siRNA was specific; other NGF-mediated signaling events were unaffected by the silencing of neuritin. Together these data indicate that neuritin is upregulated by NGF and plays a functional role in mediating neurite outgrowth of sensory neurons.

This investigation shows reduced expression and axonal transport of neuritin in sensory neurons of STZ-induced diabetic rats. These deficits were prevented by treatment with NGF. The in vitro studies show that neuritin is instrumental in the translation of NGF signals to promote neurite outgrowth in adult rat sensory neurons. Inhibition of neuritin using silencing RNA abolished NGF-mediated neuritogenesis.

A deficit in NGF expression by sensory neuron targets is well established in experimental diabetic neuropathy (13,15,35). Furthermore, replacement therapy with exogenous NGF in diabetic rats normalizes key molecular and functional aspects of the neuropathy (36,37). Hence, it is likely that the neuritin deficit reported here in diabetic rats is secondary to a deficit in endogenous NGF. It remains to be determined whether other stimuli such as hypo-insulinemia may also contribute to the deficit. The precise mechanism of action of NGF on neuritin expression has not been fully elucidated but appears to be dependent on mitogen-activated protein kinase and/or PI-3K activation.

Neurotrophins have previously been shown to regulate neuritin gene expression in a number of other cell types. Brain-derived neurotrophic factor and neurotrophin-3 caused an upregulation of neuritin in hippocampal and cortical neurons (22). It has also been demonstrated that hypoxia induces neuritin mRNA transcription in human microendothelial cells (HMEC-1) (38). Neuritin was identified in a microarray screen of NGF-regulated genes in rat pheochromocytoma cells (39) and has recently been found to promote neuritogenesis in this cell line (25). In this study, we demonstrate that NGF upregulates neuritin in adult rat sensory neurons, which mediates neurite outgrowth.

Sequencing of the promoter region of the mouse neuritin gene has found at least nine binding sites for transcription factors, including three sequences similar to cyclic AMP response element–binding protein (CREB), three sequences similar to AP-1 binding sites, a TPA-responsive element (TRE), and two sequences similar to the early growth response family binding site (40). These transcription factors have been implicated in NGF-mediated neurite outgrowth (41,42).

Stimulation of TrkA via NGF leads to activation of several intracellular signaling pathways including Ras-Raf-MEK-ERK and P1–3K–AKT (43). Inhibition of both MEK and PI-3K inhibited neurite outgrowth in this study and also prevented NGF-mediated upregulation of neuritin. The MEK-1/2 inhibitor U0126 reduced neuritin levels to below control levels—possibly indicating inhibition of an endogenously produced factor in the culture system. The PI-3K inhibitor reduced NGF upregulation of neuritin levels but not below control levels. This may indicate two pathways by which neuritin is upregulated, an NGF-mediated ERK/AKT pathway and an unidentified factor produced in culture (possibly by neurons themselves or contaminating Schwann cells or fibroblasts) maintaining a “constitutive” expression of neuritin via ERK activation.

The mechanism of the neuritogenic action of neuritin remains unknown. Neuritin is a glycosylphosphatidylinositol (GPI)-linked protein, without transmembrane or cytoplasmic domains. There are examples of such GPI-linked proteins acting as heterophilic or homophilic adhesion molecules, e.g., glypican-1, a heparin sulfate proteoglycan, which acts as a co-receptor for slit FGF, and laminin (44). Neuritin, however, lacks the immunoglobulin G domain common to GPI-linked adhesion molecules but does show some structural similarity to ephrins, a family of GPI-linked ligands that act as guidance molecules, activating their receptor by cell-cell contact (45).

There is conflicting evidence regarding the functional role of GPI cleaved soluble neuritin. Infection of Xenopus tectal cells with full-length neuritin promoted dendritic arborization, in contrast to infection with truncated neuritin lacking the GPI consensus signal (26). However, transfection of rat hippocampal neurons with truncated neuritin did promote neuritogenesis (22). Expression of full-length neuritin in Xenopus motor neurons increased the rate of axonal outgrowth by promoting axonal branching and reducing axonal retraction from presynaptic sites, indicating that neuritin modulates both axonal structure and function (24). Neuritin, in either truncated, secreted, or full-length version, was found to alter the morphology of NIH 3T3-cells producing neurite-like outgrowths and arborization (46). We found that neuritin is localized in discrete pucta within neurites, particularly at branch points and growth cones. Since neuritin promotes extension and branching of neurites, it will be of interest to determine whether this occurs through interactions of neuritin with the extracellular matrix at points of focal adhesions. It is interesting to note that a recent study of small-fiber innervation of the cornea in diabetic patients detected reduced branching as a significant change associated with the earliest stage of neuropathy (47). This may derive from changes in neuritin expression in peripheral fibers.

Axonal regeneration is defective in both experimental (1,48,49) and clinical (3,50) diabetic neuropathy. Furthermore, when measurements are focused on fibers of appropriate phenotype, NGF supplementation is shown to reverse the regeneration deficit in experimental diabetes (51). However, potential therapeutic effects of NGF treatment in diabetic neuropathy were negated due to the deleterious painful side effects observed in phase II clinical trials (52). It is likely that NGF-induced expression of neuritin in diabetes forms an important part of the normalization of nerve function by NGF; it will now be important to determine whether manipulation of neuritin levels in sensory neurons following injury or neuropathy may provide a potential target for therapeutic intervention in the management of peripheral nerve trauma or neuropathy.