Draxin Induces Cognitive Impairment in Mice via Wnt/β-Catenin-Mediated Tau Phosphorylation

Abstract

Background:

Dorsal repellent axon guidance protein (draxin) is a secreted protein that plays an establishment role in the formation of proper connections between neurons. Although draxin is known to regulate the elongation of axons from various types of neurons in vitro, its specific role in mature neurons remains unclear. Draxin expression in the hippocampal region of patients with Alzheimer’s disease (AD) has been reported to be higher than in normal subjects. The present study investigated the effect of draxin on the expression of microtubule-associated protein 2 (MAP2) and neuronal nuclear antigen (NeuN), and on tau protein phosphorylation in mouse hippocampal neurons (HT22 cells) and AD cellular models. In addition, stereotactic techniques were used to inject neuronal-targeted adeno-associated virus (AAV) into the hippocampus of C57BL/6 mice to assess the effects of draxin overexpression on hippocampal neurons, as well as on behavioral and pathological features.

Methods:

In vitro experiments were conducted using mouse hippocampal neuronal cells (HT22 cells) and established AD cellular models, focusing on evaluating draxin’s effects on the expression of neuronal markers (MAP2 and NeuN) and the phosphorylation status of tau protein. For in vivo validation, neuron-targeted AAV was delivered into the hippocampus of C57BL/6 mice via brain stereotactic injection to achieve draxin overexpression. Subsequent assessments included analyses of hippocampal neuronal integrity, behavioral tests (Y-maze and Morris water maze, to evaluate spatial learning and memory), and detection of AD-related pathological markers.

Results:

In vitro experiments revealed that draxin overexpression decreased the cell survival rate, increased the apoptosis rate, decreased the expression of MAP2 and NeuN, and showed a trend for increased phosphorylation of tau protein compared with the control group. Notably, the spatial learning memory ability of mice with draxin overexpression in the brain, as determined by the Y-maze and Morris water maze tests, was significantly diminished compared with the control group. These mice also showed elevated tau protein phosphorylation and altered expression of wingless-related integration site (Wnt)/β-catenin/glycogen Synthase Kinase 3 beta (GSK-3β) pathway components.

Conclusions:

Our results suggest for the first time that neuronal overexpression of draxin may induce neuronal damage via the Wnt/β-catenin/GSK-3β signaling pathway, leading to AD-like neuropathological damage and cognitive dysfunction.

Graphical abstract

Keywords

draxin / hippocampal neurons / cognitive dysfunction / Alzheimer’s disease

1. Introduction

Alzheimer’s disease (AD) is a neurodegenerative disease characterized by progressive cognitive dysfunction. The main characteristic pathological changes of AD are senile plaques formed by extracellular amyloid-\beta (A\beta) and neurofibrillary tangles (NFTs) formed by aggregates of phosphorylated tau (p-tau) [1], leading to neuronal dysfunction and ultimately to cognitive decline. The rate of neurogenesis declines during adulthood and aging [2, 3, 4]. Moreover, adult hippocampal neurogenesis is significantly reduced in AD patients and murine models of AD [5, 6]. The expression of neurogenesis-associated genes, including doublecortin (DCX) and sex-determining region Y-box 2 (Sox2), is reduced in patients with severe AD compared with early AD patients and non-demented controls, possibly impairing hippocampal neurogenesis [3, 7, 8, 9].

Dorsal inhibitory axon guidance protein (Draxin) is a secreted protein involved in axon guidance and the development of the forebrain and dorsal spinal cord [10, 11, 12]. Recent studies suggest that draxin may also be involved in the regulation of mature neurons. For example, draxin protein [13] can be detected in the peripheral blood of the elderly, and its expression level is elevated in patients with post-traumatic stress disorder [14]. Draxin expression is increased in the intestinal tissues of patients with inflammatory bowel disease and correlates with a poor response to treatment [15]. Furthermore, high draxin expression in gliomas correlates with a poor prognosis [16]. These findings suggest that draxin may exert a regulatory effect on the nervous system in adulthood and may have detrimental effects on mature neurons. Although direct evidence linking draxin to AD is limited, our previous bioinformatics analysis of the Alzheimer’s disease database (AlzData) database found that draxin expression in the hippocampus of AD patients is elevated compared with controls (p = 0.044). In addition, hydroxymethylation of the upstream region of the draxindraxin gene is associated with the formation of p-tau [17]. This suggests a potential role for draxin in the pathogenesis of AD, although the exact mechanism of action remains to be elucidated. In the present study, draxin was found to have injurious effects on mouse hippocampal neuronal cells. It was also found to increase the level of tau protein phosphorylation. Moreover, draxin overexpression significantly impaired the learning and memory capacities of mice and exacerbated the progression of AD-related pathology.

2. Materials and Methods

2.1 Mice, Brain Stereotactic Injection, and Experimental Groups

Male C57BL/6 mice (15 weeks old) weighing approximately 23–28 g were purchased from Jiangsu Jicui Pharmachem Biotechnology Co., Ltd. (Nanjing, Jiangsu, China). All mice were housed (five animals per cage) in specific pathogen-free facilities at the Guangdong Medical University Laboratory Animal Center. They were maintained at a constant temperature of 23 \pm 2 °C, a relative humidity of 60 \pm 5%, a 12-h light/12-h dark cycle, and with free access to food and water. Bedding and fresh water were replaced weekly, and every 5 days post-modeling. All animal-related experiments in this study were approved by the Ethics Committee of Guangdong Medical University (Ethics Approval No. GDMU-2023-000109) and adhered to international practices for animal experiments.

Phosphate-buffered saline (PBS), empty adeno-associated virus (AAV), and draxin-expressing AAV (GOSV0403972; Shanghai Genechem Co., Ltd., Shanghai, China) were delivered into the hippocampus of 15-week-old male C57BL/6 mice using stereotactic techniques. The animals were securely positioned in a stereotactic frame (018.116; NEUROSTAR, Malvern, PA, USA) with the head maintained in a flat skull position. The anterior fontanelle was exposed following a midline scalp incision with ophthalmic scissors. According to the mouse brain stereotactic atlas, the Cornu Ammonis 1 (CA1) region was targeted with the following coordinates relative to the anterior fontanelle: 2.1 mm posterior, 1.8 mm lateral from the midline, and 2.0 mm ventral from the skull surface. A total volume of 1 µL of each solution was slowly infused using a micro-syringe (980-02918-00; Ningbo Zhenhai Glass Instrument Factory, Ningbo, Zhejiang, China) at a rate of 0.1 µL/min. The needle was kept in place for an additional 5 min to facilitate adequate diffusion of the solution and minimize backflow. Subsequently, the needle was gradually withdrawn to prevent reflux. The wild-type C57BL/6 mice were assigned to three experimental groups based on the injected material. The wild-type (WT) +PBS group was injected with PBS, the WT+AAV-NC (negative control) group was injected with empty AAV, and the WT+AAV-draxin group was injected with draxin-expressing AAV. At the end of all experimental procedures (including behavioral tests), mice were euthanized via an intraperitoneal injection of sodium pentobarbital (150 mg/kg, Sigma-Aldrich, St. Louis, MO, USA) to ensure a humane death. Euthanasia was confirmed by the absence of vital signs (cessation of breathing, loss of corneal reflex, and no response to paw pinch) for at least 5 min. The brains were then rapidly dissected for subsequent histological and molecular analyses.

2.2 Cell Culture

The HT22 ( iCell-m020) mouse hippocampal neuronal cell line used in the experiments was purchased from Cellverse (iCell) Bioscience Technology Co., Ltd. (Shanghai, China). All cells were authenticated by the vendor and tested negative for mycoplasma prior to use. Cell experiments were conducted in a sterile, ultra-clean facility within the cell room of the Institute of Neurology at Guangdong Medical University, adhering strictly to aseptic requirements. The cells were cultured in an incubator under optimal conditions, featuring a gas phase composition of 95% air and 5% carbon dioxide, a constant temperature of 37 °C, and a humidity range of 70%–80%.

2.3 Viruses Used for Overexpression

(1) Lentivirus: pHBLV-CMV-MCS-3FLAG-EF1-ZsGreen-T2A-PURO, an empty load virus, and a draxin-overexpressing virus were purchased from Hanheng Biotechnology Co., Ltd. (HH20221222GYQ-LV01, Shanghai, China).

(2) Adeno-associated virus: GV753-hsyn promoter-EGFP-FT2A-MCS-WPRE-SV40 PolyA, empty load virus, and draxin-overexpressing virus were purchased from GeneChem Co., Ltd. (GOSV0403972, Shanghai, China).

2.4 Cell Counting Kit-8 (CCK8) Assay

Cells with favorable viability were plated into 96-well plates for subsequent experiments, with cell numbers adjusted for the specific experimental requirements. For cell viability assays, approximately 2 \times 10³ cells were seeded per well, and the assay was performed after cells had fully adhered to the plate and reached an appropriate density. To determine the optimal A\beta42 (04010011526, Qiangyao Biotech Co., Ltd., Shanghai, China) concentration, around 1 \times 10⁴ cells were seeded into each well, followed by incubation with A\beta42 for 24 h before the assay. The CCK8 (CK04-11, DOJINDO Laboratories, Kyushu, Japan) assay was performed according to the manufacturer’s instructions.

2.5 Protein Immunoblotting

Tissues were homogenized in lysis buffer (2500070007, Solarbio, Beijing, China) containing protease and phosphatase inhibitors (A32961; Thermo Fisher Scientific, MA, USA) and centrifuged at 12,000 rpm for 20 min at 4 °C. Supernatant was collected and the protein concentrations quantified using a bicinchoninic acid assay (BCA) (3213970, ThermoFisher Scientific, Waltham, MA, USA). Proteins (45 µg) were mixed with loading buffer, electrophoresed on SDS-PAGE gel, and then transferred to a polyvinylidene difluoride membrane (PVDF IPVH00010, Millipore Corporation, Burlington, MA, USA). The membranes were blocked with TBS/0.1% Tween-20 (TBST) buffer containing 5% bovine serum albumin (V900933-100G; Sigma-Aldrich) at room temperature, then incubated overnight at 4 °C with MAP2 antibody (rabbit, 1:1000, 8707S, Cell Signaling Technology, Danvers, MA, USA), NeuN antibody (rabbit, 1:1000, 24307S, Cell Signaling Technology), phosphorylated tau (Ser396) (p-tau396) antibody (rabbit, 1:1000, AB109390, Abcam, Cambridge, Cambridgeshire, UK), phosphorylated tau (Thr231) (mouse, 1:1000, PA5-117230, Invitrogen, MA, USA), draxin (sheep, 1:1000, CDUB03240, R&D Systems, Minneapolis, MN, USA), or primary antibodies to \beta-catenin (1:1000, 8480, Cell Signaling Technology), GSK-3\beta (1:1000, 9315S; Cell Signaling Technology), and lipoprotein receptor-related protein 6 (LRP6) (1:1000, 3395S; Cell Signaling Technology). All primary antibodies were obtained from Cell Signaling Technology. The membranes were then washed with TBST (five times for 3 min each) and incubated with secondary antibodies (1:10,000 in TBST, BA1054, Boster Biological Technology Co., Ltd., Wuhan, Hubei, China) for 1 h. Signals were visualized using enhanced chemiluminescence (ECL) (BMU102-CN, Abbkine Scientific Co., Ltd., Wuhan, Hubei, China) (Azure Biosystems C600, Dublin, CA, USA) and optical densities were analyzed using ImageJ software (v1.54f, National Institutes of Health, Bethesda, MD, USA). Specifically, quantification software and settings were as follows: the “Gels” tool of ImageJ v1.54f was used and band optical density (OD) was measured after background subtraction (rolling ball radius = 50 pixels). Additionally, normalization details were verified: loading control bands were confirmed to be within the linear range of detection via serial dilution validation, ensuring accurate normalization. For averaging of replicates, data from 3–4 independent biological replicates were first normalized to the loading control [vinculin (1:1000, 13901S; Cell Signaling Technology) or (glyceraldehyde-3-phosphate dehydrogenase) GAPDH (1:1000, 5174S; Cell Signaling Technology)] for each sample, then averaged to calculate the relative expression level [mean \pm standard error of the mean (SEM)]. Furthermore, for western blot (WB) densitometry statistical analysis, one-way ANOVA (for multiple groups) or two-way ANOVA (for factorial designs) was used to compare relative expression levels, with Tukey’s post-hoc test for pairwise comparisons (significance set at p 0.05). Original Western blot images are provided in Supplementary Material-WB Original.

2.6 Quantitative Real-Time Polymerase Chain Reaction

Relative mRNA levels were measured via quantitative real-time polymerase chain reaction (qRT-PCR) with validated primer pairs (Table 1). Total RNA was extracted from cells using TRIzol reagent (15596018CN, Ambion, Austin, TX, USA) and cDNA was synthesized using a reverse transcription kit (RR820Q, TaKaRa, Shiga, Japan). The assays were performed on a LightCycler 96 quantitative PCR instrument (ABI QuantStudio 6 Pro, Thermo Fisher Scientific) and a 96-well optical plate [18].

2.7 Tissue Immunofluorescence

Mouse brain sections were washed three times with PBS. They were blocked in PBS containing 10% normal goat serum (240528; Affinity Biosciences Ltd., Changzhou, Jiangsu, China) for 1 h at room temperature, then incubated overnight at 4 °C with the primary antibody. After washing with PBS, sections were incubated in blocking buffer with secondary antibodies for 1 h. Cell nuclei were stained with Hoechst 33342 (C0031-01, Solarbio) for 8 min at room temperature. Following three PBS washes, images were acquired using a confocal microscope (FV3000, Olympus, Tokyo, Japan). Primary antibodies used for immunostaining were MAP2 (rabbit; 1:200, Cell Signaling Technology) and NeuN (rabbit, 1:200, Cell Signaling Technology). Secondary antibodies were Alexa Fluor 488 streptavidin anti-biotin (1:400, 702410ES03, Yeasen Biotechnology, Shanghai, China) and Alexa Fluor 488/555 coupled goat anti-rabbit IgG (1:800, cr3358443-5, Abcam).

2.8 Water Maze Experiments

The Morris water maze (MWM) was used to evaluate spatial learning and memory in mice. Briefly, a circular pool (diameter 120 cm, height 50 cm) filled with water (22 \pm 1 °C, water depth 30 cm) was rendered opaque with non-toxic white pigment. A camera system was mounted vertically approximately 3 m above the pool. The pool was divided into four quadrants: east, west, south, and north, with a platform (diameter 12 cm, height 29 cm) placed centrally in the north quadrant. Distinctive visual cues (different colors/shapes) were fixed around the pool to aid platform localization. The 7-day test included three phases: visible platform training, hidden platform training, and probe trial (no platform). On day 1 (visible platform training), the platform was exposed 1 cm above the water, and each mouse was released into the water from the center of each quadrant (north, east, west, south). Latency to find the platform was recorded. Mice failing to locate the platform within 60 s were guided to the platform and allowed to stay for 15 s. A submerged platform (1 cm below the water) was used on days 2–6 (hidden platform training). Each mouse was trained four times per day and released from different quadrants in varying order. Escape latency (time to find the platform) was recorded, with training ending at the 60th second, and a 15-s temporary stay on the platform for mice failing to locate it. On day 7 (probe trial), the platform was removed. Spatial exploration was assessed by releasing mice from the quadrant opposite the original platform location. Data were analyzed by two-way repeated measures ANOVA.

2.9 Y-Maze Experiment

Spatial memory was assessed using a Y-maze with removable arm partition baffles, as previously described [19]. Mice were habituated to the experimental environment for 1 week before the formal experiment. After habituation, distinct colored and shaped stickers were affixed to the end of each arm. The formal experiments commenced with each mouse tested for 10 min per session. The Y-maze experiment consisted of two phases, spaced 1 h apart. During the first phase of training, the novel arm was blocked with a partition baffle, and mice were placed at the end of the starting arm. After 10 min of free exploration at the starting arm and other accessible arms, mice were returned to their home cages. The second phase of the test was conducted 1 h later, with the partition baffle removed from the novel arm, and each mouse repositioned at the end of the starting arm. The time spent, distance traveled, and number of arm entries were recorded during 10 min of free exploration across all three arms.

2.10 AD Database Mining and Analysis

Draxin expression data were acquired from the AD-dedicated database AlzData (http://www.alzdata.org/) by integrating five publicly available gene expression datasets (GSE28146, GSE29378, GSE36980, GSE48350, GSE5281). Samples were selected based on three stringent criteria: human hippocampal tissue, clear grouping into AD patients or age-matched normal controls, and \ge90% data integrity. The final cohort comprised 68 AD and 52 normal controls, with all AD cases conforming to the National Institute on Aging and Alzheimer’s Association (NIA-AA) diagnostic criteria. Dataset-specific sample sizes were as follows: GSE28146 (AD n = 12, control n = 8), GSE29378 (AD n = 15, control n = 10), GSE36980 (AD n = 14, control n = 9), GSE48350 (AD n = 13, control n = 12), and GSE5281 (AD n = 14, control n = 13). Data analysis was performed using R software (v4.3.0, R Core Team, Vienna, Austria). Quantile normalization was applied to eliminate inter-dataset batch effects. The limma package (3.54.1, Gordon K. Smyth, Walter and Eliza Hall Institute of Medical Research, Melbourne, VIC, Australia) was used for linear model fitting to calculate draxin’s Log2 fold change (Log2FC) between the AD and control groups, two-tailed t-tests generated raw p-values, and the Benjamini-Hochberg method was employed to adjust for false discovery rate (FDR), with significance thresholds set at p 0.05 and FDR 0.2. To ensure the robustness and reliability of the integrated analysis, we performed additional reproducibility and sensitivity checks. Reproducibility check: we re-analyzed each dataset independently (rather than only integrating them) and confirmed consistent upregulation of draxin in AD hippocampi across all five datasets (p 0.05 for GSE28146, GSE29378, and GSE5281; p 0.1 for GSE36980 and GSE48350), supporting the robustness of the finding. Sensitivity analysis: exclusion of any single dataset did not alter the overall result (pooled p 0.05), indicating the integrated analysis was not biased by individual datasets.

2.11 Data Analysis

Images were analyzed using ImageJ. All data were expressed as the mean \pm SEM and analyzed using one-way ANOVA or two-way ANOVA for multiple comparisons. The significance level was set at p 0.05. The statistical software used in the study was GraphPad Prism 9.0 (GraphPad Software, Inc., San Diego, CA, USA) for Windows.

3. Results

3.1 Elevated Draxin Expression in the Hippocampus of AD Patients, and Construction of Draxin-Overexpressing Mouse and AD Cell Models

Following a search of the AD database AlzData—encompassing the publicly available datasets GSE28146, GSE29378, GSE36980, GSE48350, and GSE5281—draxin expression in the hippocampal region of AD patients was found to be higher than in normal subjects (p = 0.044; Fig. 1A). After considering data from multiple brain regions, no difference in the FDR was found (Fig. 1B). To investigate the role of draxin overexpression in the pathogenesis and progression of AD, we constructed a mouse model of draxin overexpression by injecting draxin-AAV into the bilateral hippocampus of WT mice. The spontaneous green fluorescent protein (GFP) green signal of the virus was visualized using fluorescence microscopy (Fig. 1C). A qRT-PCRassay showed that the level of draxin expression in the WT+AAV-draxin group was significantly increased relative to the WT+PBS and WT+AAV-NC groups (p 0.05; Fig. 1D).

An AD cell model with overexpression of draxin was also constructed with HT22 cells. After mapping the optimal Multiplicity of Infection (MOI), draxin lentivirus was successfully transfected into HT22 cells, with the spontaneous GFP green signal of the virus visible by fluorescence microscopy (Fig. 1E). qRT-PCR revealed that draxin expression was significantly higher in the lentivirus (LV)-draxin group compared with the negative control (NC) and lentivirus-negative control (LV-NC) groups (p 0.0001; Fig. 1F), indicating successful construction of the HT22 cell model with draxin overexpression. To select the optimal A\beta concentration, HT22 cells were treated with A\beta42 (0–50 µM) for 24 h, and their viability was then quantified using a CCK8 assay. A dose-dependent reduction in viability (p 0.05 vs control) was observed starting at 20 µM (p 0.01; Fig. 1G). After the treatment of cells with A\beta42 at a concentration of 20 µM, western blotting showed significantly increased tau protein phosphorylation (p 0.05; Fig. 1H,I). Therefore, a 20 µM concentration was selected to treat HT22 cells and AD cell model constructs.

3.2 Draxin Overexpression Reduces Cell Viability and the Expression of Neuronal Marker Proteins MAP2 and NeuN in HT22 Cells and AD Cell Models

To investigate whether draxin has a detrimental impact on hippocampal neuronal cells, the viability of cells in the NC, LV-NC, and LV-draxin groups was evaluated by CCK8 assay, while the apoptosis rate was assessed by Hoechst staining. The survival rate of cells in the LV-draxin group was significantly decreased compared with the LV-NC group (p 0.001; Fig. 2A), while the apoptosis rate was significantly increased (p 0.01; Fig. 2B). Similar results were observed in the AD cell model constructed with HT22 cells and A\beta42, with the survival rate of cells in the LV-draxin+A\beta42 group being significantly decreased, while the apoptosis rate was significantly increased (p 0.05; Fig. 2C,D). The expression levels of MAP2 and NeuN are shown in Fig. 2E. The expression of MAP2 and NeuN was lower in the LV-draxin group compared with the LV-NC+A\beta42 group, reaching significance for MAP2 (p 0.05; Fig. 2F).

3.3 Effects of Draxin Over-Expression on Tau Protein Phosphorylation and the Expression of Wnt/\beta-Catenin/GSK-3\beta Pathway Components in HT22 Cells and AD Cell Models

To determine whether draxin aggravated AD pathology, we examined the expression levels of p-tau in three groups of cells: NC, LV-NC, and LV-draxin. A trend for increased levels of phosphorylated tau proteins p-tau231 and p-tau396 was observed in the LV-draxin group in contrast to the LV-NC group (Fig. 2G). The level of phosphorylated tau protein p-tau396 in the NC group was significantly higher than in the LV-NC group (p 0.01), while p-tau231 showed a non-significant increase (p > 0.05). The expression level of tau proteins in NC+A\beta42, LV-NC+A\beta42, and LV-draxin+A\beta42 cells is shown in Fig. 2H. The expression of p-tau231 was significantly increased in the LV-draxin+A\beta42 group compared with the LV-NC+A\beta42 group (p 0.05), while the expression of p-tau396 showed a non-significant increase (p > 0.05). In the LV-NC+A\beta42 group, the expression of p-tau231 was significantly increased (p 0.01), while p-tau396 showed a non-significant increase (p > 0.05) compared with the NC+A\beta42 group. Subsequently, we also evaluated the expression level of Wnt/\beta-catenin/GSK-3\beta pathway components (Fig. 3A). The expression of Wnt3a and \beta-catenin was significantly downregulated in the LV-draxin group compared with the LV-NC group (p 0.05), while GSK-3\beta and LRP6 expression was upregulated (p 0.05). The LV-NC group showed no significant differences in the expression of Wnt3a, \beta-catenin, GSK-3\beta, and LRP6 compared with the NC group (all p > 0.05). Collectively, these findings suggest that overexpression of draxin in HT22 cells promotes the phosphorylation of tau proteins through activation of the Wnt/\beta-catenin/GSK-3\beta pathway, especially when stimulated by A\beta42.

3.4 Draxin Overexpression Impairs Spatial Learning and Memory Ability in Mice

The Y-maze test and MWM were used to evaluate the learning and memory ability in the three groups of mice: WT+PBS, WT+AAV-NC, and WT+AAV-draxin. The spontaneous alternation rate (alternation %) of mice in the WT+AAV-draxin group was significantly lower than in the WT+AAV-NC group (p 0.01; Fig. 4A,B). The spatial recognition experiments revealed no significant differences between the three groups in terms of the number of new arm entries (number of new arms) and the percentage of exploration time in the new arm (time in the new arm%) (p > 0.05; Fig. 4B).

In the MWM experiments, the time to find the platform (platform latency) of the mice decreased progressively over time during navigation training in the WT+AAV-NC and WT+PBS groups, but not in the WT+AAV-draxin group (Fig. 4C,D). In the probe experiment on day 6, mice in the WT+AAV-draxin group exhibited a significantly decreased number of platform crossings compared with the other two groups (p 0.001), as well as decreased time and distance in the target quadrant (p 0.05; Fig. 4E). These results suggest that draxin overexpression in the mouse brain leads to a decline in spatial learning and memory ability.

3.5 Neuropathological Changes in WT Mice Induced by Draxin Overexpression

Neuropathological changes in the mouse brain were examined by detecting the expression of MAP2, NeuN, and p-tau, as well as the Wnt/\beta-catenin/GSK-3\beta pathway (Fig. 3). Immunofluorescence (IF) revealed significantly decreased expression of MAP2 and NeuN in the WT+AAV-draxin group compared with the WT+AAV-NC group (p 0.05; Fig. 3B–E). Western blot results showed that MAP2 expression in the hippocampus of mice from the WT+AAV-draxin group was significantly decreased compared with the WT+AAV-NC group (Fig. 3F,H). There was no significant difference in the expression levels of MAP2 and NeuN between the WT+AAV-NC and WT+PBS groups.

The level of p-tau396 in mice from the WT+AAV-draxin group was increased compared with the WT+AAV-NC group (p 0.01; Fig. 3G,I–K), while no significant difference was observed between the other two groups (Fig. 3G,I). GSK-3\beta expression was also increased in the WT+AAV-draxin group (p 0.05), while \beta-catenin showed a non-significant decrease, and LRP6 a non-significant increase (p > 0.05). These results suggest that draxin overexpression in the mouse hippocampus promotes pathological hyperphosphorylation of tau protein, as well as neuronal damage that appears to be mediated by the Wnt/\beta-catenin/GSK-3\beta pathway.

3.6 Transcriptomic Sequencing of HT22 Cells With Draxin Overexpression

To explore the mechanisms underlying the detrimental impacts of draxin overexpression on the brain and neurons, RNA sequencing analysis was performed on HT22 cells from the LV-NC and LV-draxin groups. Significant changes were found in the expression of 66 genes, of which 31 were up-regulated and 35 were down-regulated (Fig. 5A). Heatmap clustering analysis showed different expression patterns among the samples (Fig. 5B). Gene ontology (GO) and kyoto encyclopedia of genes and genomes (KEGG) analyses revealed that the top 20 differentially expressed genes were significantly enriched in pathways for regulation of ribonuclease activity, adhesion of symbiont to host, response to protozoan, cellular response to interferon-beta, defense response to symbiont, defense response to virus, and regulation of nuclease activity. Amongst these, binding, cell part, and cellular process-related genes accounted for the highest percentage of biological processes (BP), cellular components (CC), and molecular functions (MF) (Fig. 5C).

Further analysis showed that draxin overexpression had a significant effect on hippocampal neuron glucose metabolism and alcohol metabolism, as well as regulating the stress response of aldehydes and the microvillus function of Schwann cells (Fig. 5D,E). KEGG enrichment analysis revealed that the differentially expressed genes were involved in a variety of metabolic regulations, with C5-branched-chain dicarboxylic acid metabolism (rich factor close to 1) being particularly significant. These genes were also enriched for biological processes such as the NF-\kappaB signaling pathway and extracellular matrix (ECM)-receptor interactions, and were associated with a variety of human diseases (Fig. 5F). Pathway analysis further revealed the important roles of the differentially expressed genes in glucose metabolism, lipid metabolism, amino acid metabolism, signaling, and the immune system (Fig. 5G). Finally, expression of the pro-apoptotic gene BCL2-modifying factor (BMF) was significantly elevated in the LV-draxin group (p 0.05), whereas expression of the anti-apoptotic genes aldo-keto reductase family 1 member B1 (AKR1B1) (p 0.01) and apoptosis and caspase activation inhibitor (AVEN) (p 0.05) were significantly reduced (Fig. 5H), suggesting that draxin may affect cell fate by regulating apoptosis-related genes.

4. Discussion

Elevated draxin expression in the hippocampus of AD patients has not been previously documented. The current study revealed for the first time the effects of draxin overexpression on mouse neurons and cognitive functions. This encompasses decreased expression of neuronal markers (MAP2, NeuN) and deficits in spatial learning and memory. Furthermore, draxin overexpression significantly promoted tau protein phosphorylation (p-tau231 and p-tau396) and neuronal apoptosis and inhibited the Wnt/\beta-catenin signaling pathway by activating GSK-3\beta. Collectively, these results imply a plausible pathogenic role of draxin overexpression in AD development and progression.

MAP2 is considered a useful marker for mature neuronal cells. NeuN, a protein encoded by the RBFOX3 gene, is also commonly used as a neuronal biomarker. Previous studies have reported that draxin inhibits neuronal axon growth in vitro [20, 21] and in vivo [22, 23]. This is consistent with the present findings that draxin decreased cell viability, decreased the expression of MAP2 and NeuN, and increased the apoptosis rate in HT22 cells and an A\beta42-induced HT-22 AD-like cellular model. Tau protein critically maintains the stability of microtubules by orchestrating their structural organization in neurons. Abnormally elevated levels of p-tau, such as phosphorylated tau threonine 231 (p-tau231) and serine 396 (p-tau396), constitute the pathological basis for neurofibrillary tangle formation (NTF), which represents a hallmark pathological feature of AD [23, 24]. Draxin overexpression gave rise to marked elevation of p-tau396 in HT22 cells, and p-tau231 in the A\beta42-induced HT-22 AD-like cellular model. These phosphorylation sites (p-tau231 and p-tau396) cooperatively promote NTF, with p-tau231 emerging earlier in AD progression than p-tau396 [25]. Our findings suggest that draxin overexpression causes direct neuronal injury and subsequent pathological changes.

Furthermore, draxin overexpression in WT mice resulted in spatial learning and memory dysfunction, as indicated by decreased time spent exploring the new arm in the Y-maze, prolonged escape latency, and reduced platform crossings in the MWM test. Moreover, mice with draxin overexpression showed reduced levels of hippocampal MAP2 and NeuN expression, along with elevated p-tau396. Draxin has been reported to be a competitive antagonist of Wnt proteins for binding to low-density LRP6, which in turn inhibits the classical Wnt/\beta-catenin signaling pathway [26]. The Wnt/\beta-catenin/GSK-3\beta pathway plays a pivotal role in neurogenesis, synaptic plasticity, and neuronal migration and differentiation, all of which are dysregulated in AD. Consequently, we systematically analyzed the expression profiles of key components in this signaling pathway. Decreased Wnt3a and \beta-catenin expression, and increased GSK-3\beta and LRP6 expression were found in the LV-draxin group, suggesting that draxin overexpression causes disruption of the Wnt3a/\beta-catenin/GSK-3\beta pathway.

GSK-3\beta is a key enzyme in tau protein hyperphosphorylation. Immunohistochemical analysis has revealed that GSK-3\beta activity in the cerebral cortex of AD patients is significantly higher than in healthy controls, and co-localized with A\beta plaques and neurofibrillary tangles [27]. GSK-3\beta exacerbates neuronal dysfunction by phosphorylating tau and destabilizing microtubules. Additionally, it indirectly impairs the neuroprotective effects of the Wnt/\beta-catenin pathway by suppressing \beta-catenin stability [28, 29]. The observed increase in p-tau elicited by draxin overexpression could potentially result from inhibitory effects on the Wnt3a/\beta-catenin/GSK-3\beta signaling pathway. This suggests that draxin may broadly affect signaling, metabolic regulation, organelle function, the cytoskeleton, the stress response, and the growth and differentiation of hippocampal neurons.

RNA sequence analysis revealed that relative expression of the BMF gene was significantly higher in the LV-draxin group compared with the LV-NC group, whereas the relative expression of AKR1B1 (aldo-keto reductase family 1 member B1) and AVEN (apoptosis and caspase activation inhibitor) was significantly lower. BMF is a member of the B-cell lymphoma-2 (Bcl-2) family of proteins, and promotes or inhibits cell death by regulating the mitochondrial pathway [30]. BMF protein regulates the permeability of the mitochondrial membrane by binding to other Bcl-2 family proteins, promoting destabilization of the mitochondrial membrane and the release of apoptosis-associated proteins [31]. Increased expression of BMF is usually associated with activation of the apoptotic pathway [31]. The AKR1B1 gene encodes a key enzyme that is involved in a variety of biological processes, including antioxidant defense [32] and metabolic homeostasis [33]. The downregulation of AKR1B1 in neurons may reduce cellular resistance to oxidative stress and increase the risk of cellular damage, thereby promoting apoptotic pathways. AVEN is known to inhibit apoptosis, in particular by interacting with Apaf-1, which blocks the release of cytochrome c from mitochondria to the apoptotic signaling pathway in the cytoplasm [34]. Thus, downregulation of AVEN may render neurons more susceptible to undergoing programmed cell death. Both the upregulation of BMF and the downregulation of AKR1B1 and AVEN suggest that draxin overexpression could potentially modulate AKR1B1- and AVEN-mediated apoptosis-related signaling pathways, although this requires further research.

In summary, our study demonstrated that draxin overexpression impairs learning and memory abilities in mice. This appears to be mediated by inhibition of the Wnt/\beta-catenin/GSK-3\beta signaling pathway, resulting in elevated tau phosphorylation and neuronal injury. Further studies are needed to fully elucidate the molecular mechanisms involving draxin in AD.

5. Conclusions

Our research yields strong evidence indicating that overexpressed draxin leads to cognitive dysfunction in mice through its damaging effects on hippocampal neurons. Suppression of the Wnt/\beta-catenin/GSK-3\beta signaling pathway may represent a potential underlying mechanism.

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Funding

Guangdong Provincial Basic and Applied Basic Research Fund(2024A1515220002)

Clinical + Basic Research Project of Guangdong Medical University(4SG23284G)

Clinical + Basic Research Project of Guangdong Medical University(GDMULCJC2024003)

Clinical + Basic Research Project of Guangdong Medical University(GDMULCJC2024036)

Affiliated Hospital of Guangdong Medical University(LCYJ2020B009)

Guangdong Provincial Medical Association Clinical Research Fund - Healthcare Special(2024HY-A6006)

2023 Special Project of the Songshan Lake Medical-Engineering Integration Innovation Center of Guangdong Medical University(4SG22307P)

Guangdong Medical University Affiliated Hospital High level Talent Research Launch Fund(GCC2022011)

Science and Technology Planning Project of Zhanjiang(2020B01395)

Science and Technology Planning Project of Zhanjiang(2020B01421)

Science and Technology Planning Project of GDMU(GDMUQ2021047)

Science and Technology Planning Project of GDMU(GDMU2021122)