Cannabidiol Protects Against 1‐Methyl‐4‐Phenylpyridinium and Manganese‐Induced Neurotoxicity via Nod‐Like Receptor Protein 3 Inflammasome Suppression

ABSTRACT

Parkinson's disease (PD) is a neurodegenerative disorder characterized by dopaminergic neurodegeneration, alpha‐synuclein (α‐Syn) accumulation, and neuroinflammation. The NOD‐Like Receptor (NLR) family pyrin domain containing 3 NLRP3 inflammasome has recently been identified as a central mediator of PD‐associated inflammatory responses. Cannabidiol (CBD), a non‐psychoactive phytocannabinoid, exhibits anti‐inflammatory and neuroprotective properties; however, its effects on NLRP3 inflammasome in PD remain insufficiently understood. This study investigated the neuroprotective effects of CBD‐rich oil against 1‐methyl‐4‐phenylpyridinium (MPP+) and manganese‐induced neurotoxicity in SH‐SY5Y cells. Cells were exposed to these substances with or without CBD co‐treatment, and cell viability, α‐Syn, dopamine, inflammatory markers [C reactive protein (CRP) and interleukin 18 (IL‐18)], and NLRP3 expressions were evaluated. MPP+ and manganese exposures significantly decreased cell viability and dopamine levels while increasing α‐Syn accumulation and inflammatory markers. Manganese induced an approximately twofold upregulation in NLRP3 mRNA and 1.5‐fold increase in protein expression. CBD co‐treatment preserved dopamine levels, attenuated α‐Syn accumulation, reduced IL‐18 and CRP concentrations, and attenuated NLRP3 expression. These findings demonstrate that CBD‐rich oil exerts neuroprotective effects in a PD cellular model by attenuating α‐Syn accumulation, preserving dopamine homeostasis, which is associated with reduced NLRP3 expression and potential modulation of inflammasome‐related signaling, supporting further investigation of CBD as a potential therapeutic strategy for PD.

CBD oil exhibits neuroprotective effects in Parkinson's disease models by inhibiting NLRP3 inflammasome activation. It preserves dopamine levels and reduces α‐synuclein accumulation, suggesting its potential as a therapeutic strategy against neuroinflammation and neurodegeneration.

1. Introduction

Parkinson's disease (PD) is a chronic, progressive neurodegenerative disorder and is considered as the second most common neurodegenerative disease after Alzheimer's disease. It is characterized by a combination of motor symptoms, including resting tremor, rigidity, bradykinesia, and postural instability as well as prominent non‑motor symptoms such as sleep disturbance, neuropsychiatric changes, fatigue, and cognitive impairment. Together, these features contribute substantially to disability and reduced quality of life [1, 2, 3].

Neuropathologically, PD is defined by degeneration of dopaminergic neurons within the substantia nigra pars compacta and the intraneuronal accumulation of misfolded α‑synuclein (α‐Syn) in Lewy bodies and Lewy neurites [4]. Beyond these hallmark features, converging evidence implicates a broader network of pathobiological processes including mitochondrial dysfunction, oxidative stress, dysregulated intracellular signaling, and immune activation, in the initiation and progression of PD [5, 6]. Despite major advances in symptomatic management, effective disease‑modifying therapies remain limited, highlighting the need to identify therapeutically targetable pathways and to evaluate potential interventions using experimentally tractable models [3, 5].

Neuroinflammation is increasingly recognized as an active contributor to PD pathogenesis rather than merely a secondary consequence of neuronal loss. In the central nervous system (CNS), inflammatory activation can be triggered by diverse pathological stimuli, including toxic exposures, and is characterized by the release of pro‐inflammatory mediators such as interleukin‐1β (IL‐1β), interleukin‐6 (IL‐6), interleukin‐18 (IL‐18), tumor necrosis factor (TNF), and reactive oxygen species (ROS) [7]. Clinical and experimental data indicate that inflammatory signatures are detectable in PD and may participate in a feed‐forward cycle in which neuronal injury promotes immune activation, which in turn amplifies neurodegeneration [6, 8].

Among inflammatory mediators, IL‐18 has emerged as a potential mechanistic contributor to PD‐associated inflammation. Elevated circulating IL‐18 levels and links between IL‐18 signaling and α‐Syn‐related inflammatory pathways have been reported in PD, and polymorphisms within IL‐18 regulatory regions have been investigated in relation to disease susceptibility [9, 10]. C‐reactive protein (CRP), an acute‐phase pentraxin synthesized during systemic inflammation, has likewise been evaluated as an accessible biomarker of inflammatory burden [11]. Meta‐analytic evidence indicates that individuals with PD exhibit significantly higher CRP concentrations in both peripheral blood and cerebrospinal fluid compared with matched controls [12]. Together, these findings support the use of IL‐18 and CRP as informative markers of PD‐related inflammatory activity in clinical and experimental contexts [9, 12].

At the molecular level, the NLRP3 inflammasome represents a key signaling platform linking cellular stress to inflammatory cytokine maturation. Accumulating evidence suggests that NLRP3 activation contributes to PD‐relevant neuroinflammatory cascades, with α‐synuclein aggregation proposed as one of several triggers enhancing inflammasome‐mediated cytokine release and neuronal vulnerability [13]. Accordingly, strategies aimed at reducing NLRP3 activation or expression have been proposed as potential approaches to limit inflammation‐driven neurodegeneration [13].

Environmental exposures also modify PD risk and progression. Heavy metals are of particular interest due to their capacity to disrupt redox homeostasis, induce oxidative injury, and impair neuronal function following systemic uptake and brain accumulation [14]. Manganese is an essential trace element, but excessive or chronic exposure leads to regional brain accumulation and extrapyramidal motor dysfunction (“manganism”) resembling parkinsonism [15, 16]. Mechanistically, manganese neurotoxicity has been associated with ROS generation, mitochondrial dysfunction, oxidative DNA damage, dysregulated autophagy‐lysosomal pathways, and activation of cell‐death signaling [17, 18]. Clinical observations of chronic manganese intoxication further support its relevance to parkinsonian syndromes [19]. In neuronal cell models, manganese (II) chloride increases oxidative stress and apoptosis and induces metabolic disturbances consistent with neurotoxic injury [20, 21, 22].

In vitro models provide controlled systems to investigate convergent mechanisms such as oxidative stress, inflammation, and apoptosis and to evaluate candidate neuroprotective agents. The human neuroblastoma SH‐SY5Y cell line is widely used in PD research due to its neuronal characteristics and responsiveness to dopaminergic stressors [23, 24, 25, 26]. PD‐related cellular alterations, including neuronal injury and α‐synuclein‐associated changes, can be induced using established neurotoxic substances [26]. One commonly used agent is 1‐methyl‐4‐phenylpyridinium (MPP⁺), the active toxic metabolite of MPTP, which disrupts mitochondrial bioenergetics and promotes oxidative stress‐mediated injury in neuronal systems [27, 28].

There is growing interest in phytocannabinoids as potential interventions for neurodegenerative disorders. Cannabidiol (CBD), a non‐intoxicating constituent of Cannabis sativa, exhibits neuromodulatory and neuroprotective properties, including anti‐inflammatory and antioxidant effects mediated through multiple molecular targets relevant to basal ganglia function and PD symptomatology [29, 30, 31]. Despite increasing clinical interest, evidence regarding CBD‐rich oils and mechanistic endpoints remains limited, highlighting the need for controlled experimental studies in disease‐relevant models [31, 32].

Accordingly, the present study investigated the neuroprotective potential of CBD‐rich oil in an in vitro PD model using SH‐SY5Y cells exposed to MPP⁺ and/or manganese. Endpoints included cell viability, dopamine levels, α‐synuclein accumulation, inflammatory markers (CRP and IL‐18), and NLRP3 expression at transcript and protein levels. By integrating biochemical and molecular readouts in a controlled MPP⁺ ‐based model, this study provides additional insight into whether CBD‐rich oil can attenuate convergent pathways of oxidative stress, neuroinflammation, and neuronal injury relevant to PD pathobiology and metal‐associated neurotoxicity, particularly by investigating its association with the NLRP3 pathway.

2. Material and Methods

2.1. Chemicals and Kits

3‐(4,5‐dimethylthiazol‐2‐yl)−2,5‐diphenyltetrazolium bromide (MTT), dimethyl sulfoxide (DMSO) and manganese (Mn) were purchased from Sigma‐Aldrich (Mannheim, Germany). Dulbecco's modified Eagle's medium (DMEM)‐Ham's F12 w/L‐Glutamine w/15‐mM HEPES, fetal bovine serum (FBS), and penicillin/streptomycin were obtained from Biowest (Riverside, MO). MPP⁺ iodide was purchased from TargetMol (Boston, MA). CBD‐rich Oil was obtained from Swiss FX(Switzerland). Dopamine and α‑Synuclein ELISA kits were obtained from Elabscience (Houston, TX). C‐reactive protein (CRP) and interleukin‐18 (IL‐18) ELISA kits are from BT‐LAB (Shanghai, China). Nucleogene Tri Reagent (cat. no: NGE023, Nucleogene). DNase I kit was obtained from ThermoFisher Scientific (USA). cDNA kit was obtained from ABM (Canada). Rotor Gene Q, MDx, 5plex, HRM system was obtained from Qiagen (Germany). SYBR Green fluorescent dye was obtained from Solis BioDyne (Estonia). RIPA lysis buffer and BCA Protein Assay kit were obtained from Thermo Fisher Scientific (USA).

2.2. Analysis of Cannabinoid Content in CBD‐Rich Oil

A preliminary analysis was conducted to confirm the reliability and standardized quantity of the Cannabidiol (CBD) oil to be used in our study. In this context, the aim was to determine the main cannabinoid content of the commercially obtained CBD‐rich oil sample. The Liquid Chromatography‐Tandem Mass Spectrometry (LC‐MS/MS) method, known for its high selectivity and sensitivity in such analyses, was employed for the high‐precision determination of the concentrations of the oil's main components, CBD and CBN. The CBD‐rich oil used in this study was obtained commercially, and the LC‐MS/MS method was used for the determination of CBD and CBN amounts.

2.3. Cell Viability

10⁴ cells/well, 100 µL complete medium) and allowed to attach overnight. Cells were exposed for 24 h to manganese (0–2000 µM), cannabidiol (CBD; 0–50 µg/mL), or MPP⁺ (0–200 µM). After treatment, medium was replaced with MTT solution (0.5 mg/mL) and incubated at 37°C for 3 h. The medium was aspirated, formazan crystals were dissolved in 150 µL DMSO, and absorbance was measured at 570 nm using a microplate reader (AMR‐100, Allsheng, China). Viability was expressed as percentage of untreated control (100%) [33, 34].

2.4. Cell Line

The human neuroblastoma cell line SH‐SY5Y (ATCC CRL‐2266™, Manassas, VA) was used in this study. SH‐SY5Y is a thrice‐cloned subline of the SK‐N‐SH cell line (ATCC HTB‐11™) [35]. MPP⁺‐induced Parkinsonian model was established by exposing SH‐SY5Y cells to MPP⁺, with or without manganese, as described in the treatment protocols [36, 37].

2.5. Experimental Groups

Cells were assigned to the following experimental groups.

• 1. Control: Cells treated only with complete culture medium.
• 2. Manganese (Mn): Cells treated with 379 µM manganese for 24 h.
• 3. Cannabidiol (CBD): Cells treated with 20 µg/mL CBD‐rich oil for 24 h.
• 4. MPP⁺: Cells treated with 79.55 µM MPP⁺ for 24 h.
• 5. MPP⁺  + Mn: Cells treated with 79.55 µM MPP⁺ and 379 µM manganese for 24 h.
• 6. CBD + Mn: Cells treated with 20 µg/mL CBD and 379 µM manganese for 24 h.
• 7. CBD + MPP⁺: Cells treated with 20 µg/mL CBD and 79.55 µM MPP⁺ for 24 h.
• 8. CBD + MPP⁺  + Mn: Cells treated with 20 µg/mL CBD, 79.55 µM MPP⁺, and 379 µM manganese for 24 h.

2.6. Quantification of Biomarkers by ELISA

The levels of α‑Synuclein (Cat. No. E‐EL‐H0983), CRP (Cat. No. E‐EL‐H0043), Dopamine (Cat. No. E‐EL‐0046), and IL‐18 (Cat. No. E0147Hu) were determined using commercial ELISA kits (Elabscience, Houston, TX; and BT Lab, Shanghai, China) according to the manufacturers' instructions. Briefly, samples and reagents were equilibrated to room temperature, and standard curves were generated using serial dilutions for each analyte. Standards and samples were incubated on pre‐coated plates, followed by treatment with biotinylated detection antibodies and HRP conjugates at 37°C. After the addition of the substrate and a 10‐15 min dark incubation, the reactions were terminated using a stop solution. Absorbance was measured at 450 nm using a microplate reader, and final concentrations were calculated using a four‐parameter logistic (4PL) regression model or linear regression analysis as specified by the kit protocols.

2.7. RNA Isolation

Total RNA was isolated from cells using a modified TRIzol method [38]. Cells were washed with PBS, transferred to sterile and nuclease free tubes, and resuspended in 1 mL Nucleogene Tri Reagent (cat. no: NGE023, Nucleogene). The suspension was homogenized by pipetting, transferred to sterile 2 mL nuclease‐free tubes, and incubated at room temperature for 15 min. Then, 250 µL chloroform was added, samples were mixed by inversion, incubated at room temperature for 10 min, and centrifuged at 12,000 × g for 15 min at +4°C. Following centrifugation, 400 µL of the aqueous phase was transferred to new tubes and mixed with an equal volume of isopropanol. The mixtures were homogenized by inversion and incubated at room temperature for 10 min. Samples were then centrifuged at 12,000 × g for 10 min at +4°C to obtain RNA pellets. The pellets were washed with 70% ethanol and centrifuged at 7500 × g for 5 min at +4°C and the obtained supernatant was discarded. This step was repeated twice. Subsequently, pellets were washed with 99% ethanol and centrifuged at 7500 × g for 5 min at +4°C to complete the wash. After ethanol removal, pellets were air‐dried for 5 min at room temperature and each sample was dissolved in 40–60 µL nuclease‐free water. RNA concentration and purity were measured using a nucleic acid spectrophotometer (SMA100, Merinton, China), and RNA integrity was evaluated by agarose gel electrophoresis.

2.8. Dnase I Treatment and cDNA Synthesis

To prevent potential genomic DNA contamination, total RNA was treated with DNase I according to the manufacturer's instructions. For each sample, 1000 ng RNA was brought to a final volume of 8 µL with nuclease‐free water, followed by addition of 1 µL 10× reaction buffer with MgCl₂ and 1 µL RNase‐free DNase I (final volume: 10 µL). After brief centrifugation, samples were incubated at 37°C for 30 min. Enzyme inactivation was performed by adding 1 µL EDTA (50 mM) to each tube, followed by incubation at 65°C for 10 min.

After DNase I treatment, cDNA synthesis was performed using the OneScript Plus cDNA kit. The reaction mixture contained 4 µL 5× RT Buffer, 3 µL nuclease‐free water, 1 µL dNTP (10 mM), 1 µL oligo(dT) (10 µM), and 1 µL RTase enzyme. Following brief centrifugation, reactions were incubated at 50°C for 15 min to synthesize cDNA, then at 85°C for 5 min to inactivate the enzyme. The resulting cDNA was 10‐fold diluted and stored at −20°C until qPCR analysis.

2.9. qPCR Application

The expression profile of NLRP3 in cells was analyzed using the Rotor‐Gene Q, MDx, 5plex, HRM system. Commercial kit containing SYBR Green fluorescent dye was used. Primers were selected based on previously published sequences, and specificity was confirmed using Primer‐BLAST (NCBI). Primer sequences, amplicon lengths, and annealing temperatures are shown in Table 1.

Table 1.

Sequences of Amplified genes.

Genes	Primer sequences	Amplicon lengths (bp)	Annealing temperature (°C)	References
NLRP3	F:5′‐GAGGAAAAGGAAGGCCGACA‐3′	92	58°C	Matias et al., 2021 [39]
	R: 5′‐TGGCTGTTCACCAATCCATGA‐3′
GAPDH	F: 5′‐ TGCACCACCAACTGCTTAGC‐3′	87	61°C	Dikmen et al., 2023 [40]
	R: 5′‐GGCATGGACTGTGGTCATGAG‐3′

Abbreviation: bp, base pair.

qPCR cycling conditions consisted of an initial denaturation at 95°C for 15 min, followed by 40 cycles of denaturation at 95°C for 15 s, annealing at 57°C–61°C for 60 s, and extension at 72°C for 30 s. Product specificity was confirmed by melt curve analysis performed from 66°C to 99°C with 1°C increments.

2.10. Western Blot Analysis

Total protein was isolated from cells using RIPA lysis buffer containing protease inhibitors. Protein concentration was determined using a BCA Protein Assay kit, and 10 µg protein from each sample was mixed with 4× LDS sample buffer. Samples were incubated at 70°C for 10 min and then loaded onto gels. Proteins were separated using 4%‐12% Bis‐Tris Plus Mini Protein Gels (Thermo Fisher Scientific, USA). SDS‐PAGE was performed in MES SDS Running Buffer at 200 V and 300 mA for 25 min. Separated proteins were transferred to PVDF membranes using a semi‐dry transfer system at 25 V and 1.3 A for 15 min. Membranes were blocked for 1 h in TBS‐T (20 mM Tris‐HCl, 150 mM NaCl, 0.1% Tween‐20) containing 5% non‐fat milk.

For NLRP3 detection, the primary antibody from Elabscience (E‐AB‐93112; 1:1000 in 5% milk) and a secondary antibody (anti‐rabbit; 1:2000) were used. For the internal control, β‐actin was detected using the primary antibody from Affinity (AF7018; 1:3000) and a secondary antibody (anti‐rabbit; 1:5000 in 5% milk). Primary antibody incubation was performed overnight at +4°C, and secondary antibody incubation was performed for 1 h at room temperature. Membranes were washed 6 times with TBS‐T for 5 min each to remove unbound antibodies, followed by a final incubation in 1× TBS for 10 min.

Protein bands were visualized using a chemiluminescent substrate (SuperSignal™ West Pico PLUS, Thermo Fisher Scientific, USA) and imaged with the ChemiDoc MP Imaging System (Bio‐Rad, USA). Band intensities were quantified using ImageJ software (version 1.53k).

2.11. Total Protein Measurement

Total protein content was determined using the modified Lowry assay. A fresh copper reagent was prepared containing 1% Cu₂SO₄, 2% Na⁺/K⁺ tartrate, and 10% Na₂CO₃. For each well, 40 µL of copper reagent was mixed with 40 µL of distilled water (blank), standard, or sample. The plate was incubated at room temperature for 10 min. Then, 120 µL of phenol solution was added to each well, and the plate was incubated in a 50°C water bath for 10 min to allow color development. Absorbance was measured at 450 nm using a spectrophotometer. Protein concentration was calculated from the standard curve using the calibration (linear regression) equation and expressed as mg/mL.

2.12. Statistical Analysis

All statistical analyses were performed using IBM SPSS (v22.0). Differences between groups were tested by one‐way ANOVA. Tukey's HSD test was used for post‐hoc comparisons. Data are presented as mean ± standard error, and p < 0.05 was considered statistically significant. For quantification of NLRP3 expression, Ct values were normalized to the GAPDH, and relative expression was calculated as fold change using the 2^‐ΔΔCt method [41]. Moreover, NLRP3 protein expression levels were normalized to β‐actin and expressed as fold change.

3. Results

3.1. Analysis of Cannabinoid Content in CBD‐Rich Oil

In the analysis of the CBD‐rich oil used in the study with LC‐MS/MS, the concentrations of CBD and CBN were determined to be 70835 mg/kg and 1635 mg/kg, respectively. The LC‐MS/MS chromatograms and mass spectra of the analysis are presented in Figures 1 and 2.

Figure 1.

LC‐MS/MS analysis data of the CBD‐rich oil sample. Chromatogram (RT: 7.96) and mass spectrum (m/z: 315.23) of CBD.

Figure 2.

Chromatogram (RT: 9.97) and mass spectrum (m/z: 311.20) of CBN.

3.2. Cell Viability

MTT assays were conducted to evaluate the dose–response effects of manganese, CBD, and MPP⁺ on cell viability. Viability values were normalized to the untreated control (100%) and expressed as percentage of control (Figure 3A–C).

Figure 3.

Dose‐response effects of (A) manganese (MnCl₂), (B) cannabidiol (CBD), and (C) MPP⁺ on SH‐SY5Y cell viability assessed by MTT assay. Cell viability is expressed as percentage of control. Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Manganese induced a concentration‐dependent decrease in viability across the tested range (Figure 3A). The lowest concentration (250 µM) reduced viability to 83.2%, followed by progressive declines to 80.1% (375 µM), 72.5% (500 µM), 66.2% (750 µM), and 55.1% (1000 µM). More pronounced cytotoxicity was observed at higher concentrations, with viability decreasing to 46.1% at 1500 µM and 38.0% at 2000 µM.

CBD was well tolerated at lower concentrations (Figure 3B). Viability remained near control levels between 5 and 20 µg/mL, including a slight increase at 5 µg/mL (103.3%) and minor fluctuations at 10–20 µg/mL (−1.5% to −5.0%). At 25 µg/mL, viability decreased to 89.8%. Cytotoxicity became evident at higher concentrations, with viability reduced to 74.3% at 40 µg/mL (*) and 49.2% at 50 µg/mL (**). Overall, CBD produced minimal effects at ≤ 20 µg/mL, whereas significant reductions occurred at 40–50 µg/mL.

MPP⁺ exposure produced a dose‐dependent decrease in cell viability (Figure 3C). Viability was reduced to 89.4% at 3.125 µM and remained in the mid‐80% range at 6.25–25 µM. Greater reductions were observed at higher concentrations, reaching 82.4% at 50 µM, 75.9% at 100 µM, and 66.2% at 200 µM. The decrease was statistically significant at 100 µM and 200 µM (Figure 3C).

The concentrations for the final experimental design were selected based on both preliminary MTT assays and literature. Mn and MPP⁺ doses were calibrated to induce a moderate neurotoxic challenge, while 20 µg/mL of CBD‐rich oil was identified as the highest non‐cytotoxic dose for the protective intervention.

3.3. α‐Synuclein Levels

α‐Syn levels were normalized to total protein content (Figure 4A). Manganese significantly increased α‐Syn by 16.5% versus control (p < 0.01), whereas CBD alone had no effect (+0.6%). MPP⁺ produced a modest increase (6.9%), while combined MPP⁺ + MnCl₂ exposure induced the greatest accumulation (+26.1%, p < 0.001).

Figure 4.

Effects of CBD on α‐Syn and dopamine levels in SH‐SY5Y cells exposed to neurotoxicants. (A) α‐Synuclein levels (B) Dopamine concentrations levels. Data are presented as mean ± SEM (n = 3 per group). Statistical significance was determined by one‐way ANOVA with Tukey's post‐hoc test. *p < 0.05, **p < 0.01, ***p < 0.001.

CBD co‐treatment attenuated toxin‐induced elevations. CBD + MnCl₂ reduced α‐Syn by 12.5% compared with MnCl₂ alone (p < 0.05). In the CBD + MPP⁺ and CBD + MPP⁺ + MnCl₂ groups, α‐synuclein levels were near control (+2.6% and +1.7%, respectively). Relative to MPP⁺ + MnCl₂, CBD + MPP⁺ + MnCl₂ decreased α‐synuclein by 19.4% (p < 0.001), indicating prevention of the synergistic accumulation induced by combined toxin exposure.

3.4. Dopamine Levels

Intracellular dopamine levels are shown in Figure 4B. Manganese reduced dopamine by 25.4% versus control (p < 0.05), whereas CBD alone caused a non‐significant decrease of 10.6%. MPP⁺ decreased dopamine by 30.0% (p < 0.05), and combined MPP⁺ + MnCl₂ exposure produced the greatest depletion (−40.4%, p < 0.001).

CBD co‐treatment preserved dopamine in toxin‐exposed cells. CBD + MnCl₂ resulted in only an 11.6% reduction compared with control and was significantly improved versus MnCl₂ alone (p < 0.05). CBD + MPP⁺ nearly restored dopamine levels (−0.1% vs. control; p < 0.05 vs. MPP⁺ + MnCl₂), while CBD + MPP⁺ + MnCl₂ showed partial protection (−21.6%). Overall, CBD markedly attenuated MPP + ‐ and manganese‐induced dopamine depletion.

3.5. C‐Reactive Protein Levels

CRP levels were evaluated as a marker of inflammatory activation (Figure 5A). Manganese and CBD alone increased CRP by 24.6% and 18.3%, respectively; however, these changes were not significant. MPP⁺ produced the greatest elevation (+39.7%, p < 0.05), while combined MPP⁺ + MnCl₂ increased CRP by 23.8%.

Figure 5.

Effects of CBD on inflammatory markers in SH‐SY5Y cells exposed to neurotoxicants. (A) C‐reactive protein (CRP) levels. (B) Interleukin‐18 (IL‐18) levels. Data are presented as mean ± SEM (n = 3 per group). Statistical significance was determined by one‐way ANOVA with Tukey's post‐hoc test. *p < 0.05.

CRP levels in the CBD + MnCl₂ (24.6%) and CBD + MPP⁺ (38.1%) groups were comparable to their respective toxin‐only groups. However, the CBD + MPP⁺ + MnCl₂ group showed a smaller increase (+15.1%), significantly lower than MPP⁺ alone (p < 0.05), suggesting a modest anti‐inflammatory effect of CBD under combined MPP⁺ and manganese exposure.

3.6. Interleukin‐18 Levels

IL‐18, a pro‐inflammatory cytokine is associated with inflammasome activation (Figure 5B). Manganese and MPP⁺ significantly increased IL‐18 by 12.0% and 11.4%, respectively (p < 0.05), whereas CBD alone had no effect (+0.8%). Combined MPP⁺ + MnCl₂ exposure produced the greatest elevation (+17.1%, p < 0.01).

CBD co‐treatment attenuated these responses. The CBD + MnCl₂ group showed a smaller increase (+5.9%), while CBD + MPP⁺ reduced IL‐18 below control (−4.2%), corresponding to decreases of 14.0% versus MPP⁺ alone (p < 0.05) and 18.2% versus MPP⁺ + MnCl₂ (p < 0.01). The CBD + MPP⁺ + MnCl₂ group exhibited only a mild elevation (+6.3%). Overall, CBD attenuated IL‐18 production, particularly under MPP⁺‐induced inflammatory conditions.

Notably, CBD reduced IL‐18 levels slightly below the control, which may be attributed to its potent anti‐inflammatory properties.

3.7. NLRP3 mRNA Expression

RNA quality was confirmed prior to qPCR (A₂₆₀/A₂₈₀ = 1.85 ± 0.03), and melt‐curve analysis verified specific amplification of NLRP3 and GAPDH. NLRP3 expression was significantly upregulated (approximately twofold) in manganese‐treated cells versus control (p = 0.03) (Figure 6A), whereas CBD or MPP⁺ alone had no significant effect.

Figure 6.

Effects of CBD‐rich oil on NLRP3 inflammasome expression at the mRNA and protein levels in SH‐SY5Y cells. (A) NLRP3 mRNA expression quantified by qRT‐PCR using the 2^−ΔΔCt method. (B) NLRP3 protein expression assessed by Western blot with β‐actin as an internal loading control. Representative blots are shown above the densitometric analysis. Data are presented as mean ± SEM of three independent experiments (n = 3). Different letters (a,b,c,d) indicate statistically significant differences between groups (p < 0.05).

CBD co‐treatment attenuated manganese‐induced upregulation. MnCl₂ + CBD and MPP⁺ + CBD groups showed nearly 1.5‐fold expression, and MPP⁺ + MnCl₂ showed approximately 1.2‐fold expression relative to control. Notably, the CBD + MPP⁺ + MnCl₂ group exhibited ~30% lower expression versus control, indicating a reduction in NLRP3 transcription.

3.8. NLRP3 Protein Expression

Manganese significantly increased NLRP3 protein levels by 39% versus control (p < 0.001) (Figure 6B), representing the highest expression among groups. CBD alone and MPP⁺ + MnCl₂ produced minimal changes (+5% and +6%). In contrast, CBD co‐treatment was associated with lower NLRP3 protein expression in MnCl₂ + CBD (− 22%), MPP⁺ + CBD (− 24%), and CBD + MPP⁺ + MnCl₂ (−29%) groups relative to control. Compared with MnCl₂ alone, CBD + MPP⁺ + MnCl₂ decreased NLRP3 protein by 49%. These results indicate that CBD‐rich oil treatment is associated with reduced NLRP3 inflammasome expression by CBD at both transcriptional and protein levels.

4. Discussion

The neurotoxicants used in this study produced dose‐dependent reductions in cell viability consistent with their known mechanisms. MPP⁺ inhibits mitochondrial complex I, leading to bioenergetic failure, oxidative stress, and cell death [28], while manganese also disrupts mitochondrial function and enhances reactive oxygen species generation [17, 20]. Huang et al. (2021) reported that manganese chloride induces dopaminergic neurotoxicity in SH‐SY5Y cells via BNIP3‐mediated mitophagy and oxidative stress [20], supporting the mechanistic basis of the observed toxicity. Combined exposure produced greater cellular injury than either agent alone, indicating additive or synergistic effects. Unlike previous studies that focus solely on single‐toxin models, our work investigates the protective role of CBD‐rich oil against the combined insult of Mn and MPP⁺, reflecting a more complex and multi‐factorial representation of PD pathology. Interestingly, our results indicated that manganese (Mn) exposure led to a more pronounced activation of the NLRP3 inflammasome compared to MPP⁺. This differential response may be explained by the distinct mitochondrial targeting of Mn; whereas MPP⁺ primarily inhibits complex I, Mn is known to cause more extensive mitochondrial fragmentation and the subsequent release of mitochondrial reactive oxygen species (ROS) and DNA (mtDNA) into the cytosol, which act as potent triggers for the NLRP3 complex [17, 20]. This observation is relevant to PD pathogenesis, as environmental factors, including metal exposure, are recognized contributors to disease risk and progression [14, 16].

A key finding of this study concerns α‐Syn, whose aggregation is a pathological hallmark of PD [4]. Manganese exposure significantly increased α‐Syn levels, with the greatest accumulation observed following combined MPP⁺ + manganese treatment. Notably, CBD‐rich oil co‐treatment prevented the induced α‐Syn accumulation, suggesting a protective effect on protein homeostasis that has not been extensively characterized in neurotoxicant‐based PD models. This cytoprotective role aligns with the multi‐target anti‐inflammatory profile of CBD, particularly its capacity to attenuate neuroinflammation and modulate stress responses [42]. These results align with previous reports showing manganese‐induced α‐Syn overexpression and aggregation in SH‐SY5Y cells. Li et al. (2010) demonstrated that manganese enhances both transcriptional and translational α‐synuclein expression and that α‐Syn overexpression exacerbates manganese‐induced apoptosis [43]. Similarly, Yan et al. (2021) reported that manganese‐induced α‐Syn elevation aggravates mitochondrial damage via repression of PINK1/Parkin‐mediated mitophagy [44]. Notably, CBD‐rich oil co‐treatment prevented toxin‐induced α‐Syn accumulation, suggesting a protective effect on protein homeostasis that has not been extensively characterized in neurotoxicant‐based PD models.

In addition to α‐Syn alterations, dopamine levels were markedly affected by neurotoxicant exposure. As dopaminergic depletion underlies the motor symptoms of PD [4], preservation of dopamine homeostasis is a key therapeutic objective. Both toxins reduced intracellular dopamine, with the greatest depletion observed following combined exposure. CBD‐rich oil co‐treatment preserved dopamine content, most prominently in the CBD + MPP⁺ group, where levels were nearly restored to control. These findings are consistent with prior studies demonstrating dopaminergic protection by CBD in PD models. Santos et al. (2015) reported increased viability and enhanced axonal and synaptic protein expression in MPP⁺‐treated PC12 and SH‐SY5Y cells via TrkA receptor involvement [45]. Gugliandolo et al. (2020) further showed that CBD counteracted MPP⁺‐induced cytotoxicity in differentiated SH‐SY5Y cells through activation of the Protein Kinase B/Mechanistic Target of Rapamycin (AKT/mTOR) pathway and modulation of autophagy via Cannabinoid Receptor Type 2 (CB2) and Transient Receptor Potential Vanilloid 1 (TRPV1) signaling [46]. Our results extend these observations by demonstrating preservation of intracellular dopamine under neurotoxic conditions.

Beyond cellular injury markers, the inflammatory responses observed here further support the therapeutic potential of CBD. Neuroinflammation is increasingly recognized as an active driver of PD progression rather than a secondary consequence of neuronal loss [6, 8], with cytokine signaling contributing to a self‐propagating cycle of neuronal damage and immune activation. Both neurotoxicants elevated IL‐18, a cytokine associated with inflammasome activation and implicated in PD [9, 10], while MPP⁺ also increased CRP, an inflammatory marker reported to be elevated in PD patients [12]. Although CRP is traditionally regarded as a systemic hepatic marker, recent evidence suggests it can also be expressed locally by neuronal cells under stress, reflecting early‐stage neuroinflammatory signaling [12]. CBD‐rich oil co‐treatment attenuated these responses, and the CBD + MPP⁺ group showed IL‐18 levels below control values. These anti‐inflammatory effects are consistent with known properties of CBD [47]. Rodrigues et al. (2024) demonstrated that CBD suppresses lipopolysaccharide (LPS)‐induced inflammation in BV2 microglial cells by inhibiting NLRP3 inflammasome activation and inducible nitric oxide synthase (iNOS) activity via CB2 receptor and peroxisome proliferator‐activated receptor gamma (PPARγ) signaling [48]. The present neuronal findings complement those microglial observations, suggesting a broader anti‐inflammatory action of CBD in PD‐relevant cellular systems.

To further investigate the anti‐inflammatory mechanisms, NLRP3 inflammasome expression was evaluated at both mRNA and protein levels. The NLRP3 inflammasome senses cellular stress and promotes maturation of inflammatory cytokines, including IL‐18 [13]. Increasing evidence implicates NLRP3 activation in PD pathogenesis, with α‐synuclein aggregates proposed as triggers for inflammasome assembly [13]. In the present study, manganese markedly activated NLRP3, producing ~2‐fold mRNA upregulation and a 39% increase in protein expression versus control. CBD‐rich oil co‐treatment significantly attenuated this response, with the CBD + MPP⁺ + MnCl₂ group showing ~49% lower NLRP3 protein levels compared with manganese alone. These findings are consistent with literature demonstrating CBD‐mediated modulation of inflammasome signaling. Liu et al. (2020) reported inhibition of NLRP3 activation by CBD in THP‐1 monocytes via purinergic receptor p2x, ligand‐gated ion channel 7 (P2X7) receptor modulation and reduced potassium efflux [49], and Chu et al. (2024) highlighted CBD as a promising anti‐inflammatory agent acting through suppression of the NLRP3 pathway [50]. The present results extend these observations by demonstrating that CBD‐rich oil treatment is associated with the suppression of neurotoxicant‐induced NLRP3 activation in a neuronal PD model.

The broad protective effects of CBD observed across multiple endpoints likely reflect its interaction with diverse molecular targets. CBD modulates cannabinoid receptors, TRPV1 channels, peroxisome proliferator‐activated receptor‐γ (PPARγ), 5‐hydroxytryptamine receptor 1A (5‐HT1A), and adenosine signaling pathways implicated in neuroprotection and inflammation [29, 30]. Patricio et al. (2020) reviewed evidence supporting CBD as a therapeutic candidate in PD, emphasizing its neuromodulatory and neuroprotective properties [29]. Such a multi‐target profile may explain its efficacy against the range of pathological alterations induced by neurotoxicant exposure in the present study. The concomitant reduction in NLRP3 expression and lower IL‐18 levels suggests that CBD‐rich oil may modulate this pathway, as the NLRP3 inflammasome activates caspase‐1, which cleaves pro‐IL‐18 into its active form [13]. By potentially suppressing NLRP3 activation, CBD appears to inhibit this pathway upstream, thereby limiting downstream inflammatory cytokine release.

Several limitations should be considered in the present study. Although widely used in PD research [24, 51, 52], SH‐SY5Y cells are an immortalized neuroblastoma line and do not fully recapitulate mature dopaminergic neurons. Consistent with this limitation, Santos et al. reported that CBD does not induce neuritogenesis in SH‐SY5Y cells due to the absence of TrkA receptor expression, indicating potential cell‐type–specific responses [45]. Use of differentiated neurons or primary cultures would strengthen the findings. Moreover, the single‐cell‐type model cannot reproduce neuron–glia interactions present in the brain; given the central role of microglia in neuroinflammation [49], co‐culture or in vivo models are warranted. Finally, in vitro concentrations may not reflect achievable in vivo exposure levels, and animal studies are necessary to confirm the observed protective effects under physiological conditions. Importantly, while we observed a clear association between CBD‐rich oil treatment and reduced NLRP3 expression, this study did not employ functional tools such as caspase‐1 activity assays, IL‐1β maturation analysis, or specific NLRP3 inhibitors to definitively prove functional inhibition. These mechanistic validations remain a target for future research to fully confirm the pathway through which CBD exerts its neuroprotective effects.

In conclusion, CBD‐rich oil mitigated multiple PD‐relevant pathological features in a neurotoxicant‐based cellular model. CBD reduced α‐synuclein accumulation, preserved dopamine content, attenuated inflammatory markers, and was associated with reduced NLRP3 expression at both mRNA and protein levels. These findings support CBD as a potential neuroprotective agent and suggest that NLRP3 modulation may be a contributing mechanism. Further validation in animal models and clinical studies is required to determine therapeutic relevance in Parkinson's disease.

Author Contributions

Göksun DEMİREL: conceptualization, methodology, writing – original draft, writing – review and editing, funding acquisition, supervision, resources. Anıl YİRÜN: conceptualization, methodology, project administration. Deniz Arca ÇAKIR: writing – original draft, software. Hüseyin ÖZKAN: validation, investigation. Berfu Şura GÜNEŞ: formal analysis, data curation. Selen SELVİ: formal analysis, data curation. Yeter EROL ÖZTÜRK: validation, investigation. Pınar ERKEKOĞLU: writing – review and editing, visualization.

Ethics Statement

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.