Migraine-induced cochlear injury triggers ZBP1-mediated PANoptosis via CGRP signaling

Abstract

Objective

Consequences of hearing abnormalities in migraine remain largely unexplored, particularly regarding peripheral outcomes. This study aims to explore how migraine induces peripheral auditory injury and what role calcitonin gene–related peptide (CGRP) plays in the inner ear, which may contribute to the precise subtyping of migraine and inform targeted auditory-protective therapies.

Methods

A mouse model of chronic migraine (CM) was established by intermittent injections of nitroglycerin and validated via mechanical and thermal nociceptive assays. CGRP signaling was assessed by Western blotting and immunofluorescence. Auditory function was evaluated using auditory brainstem response (ABR) and distortion product otoacoustic emissions (DPOAE) in CM and CGRP blockade (Rimegepant-treated) groups. RNA sequencing of cochlear tissue identified dysregulated signaling pathways, which were subsequently validated in vitro using HEI-OC1 auditory cells and in vivo within the inner ear.

Results

Repeated nitroglycerin injections induced persistent mechanical and thermal hyperalgesia, confirming successful model establishment. A significant upregulation of CGRP and its receptors, particularly in outer hair cells (OHCs) were confirmed. The CM mouse exhibited elevated ABR, DPOAE thresholds and OHC injury, all of which were mitigated by Rimegepant treatment. CGRP overstimulation triggered mitochondrial stress and mtDNA leakage in HEI-OC1 cells. Cytosolic mtDNA leakage activated the ZBP1-mediated PANoptosis pathway, leading to OHC injury. As proof of concept, ZBP1 knockdown and PANoptosis inhibition mitigated cellular damage and hearing deficits both in vivo and in vitro.

Conclusions

We demonstrated OHCs as the key targets in CM-induced, CGRP-mediated peripheral hearing impairment. CGRP exerts its pathogenic role by triggering a cascade of intracellular stress and inflammatory pathways, ultimately leading to cell death. These findings provide a phenotypic basis for a subtype of migraine-induced hearing loss, paving the way for auditory protection strategies.

Graphical Abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s10194-026-02286-2.

Introduction

Migraine, a common neurological disorder characterized by recurrent headaches [1], affects over one billion people globally and profoundly impairs life quality [2]. Beyond the primary cerebral dysfunction, migraine frequently involves sensory disturbances, including auditory, visual, and somatosensory, which constitute integral components of migraine aura or accompanying symptoms [3–5]. Among these sensory processing abnormalities, auditory abnormalities remain an important, yet largely unexplored aspect of migraine pathophysiology. Accumulating evidence substantiates that migraine may raise the risk of auditory dysfunction, with reported comorbidity rates between 15% and 49% [6–8]. Its manifestations include tinnitus [9–11], hearing loss [10], and aural fullness [12, 13]. Patients demonstrate cochlear dysfunction including higher thresholds in pure tone audiometry [14, 15], prolonged latency in auditory brainstem response (ABR) [14, 16, 17], and reduced amplitude in distortion product otoacoustic emissions (DPOAE) [16, 18]. However, the true prevalence of hearing loss in migraine may be underestimated, as a large proportion of patients may suffer from high and ultra-high frequency hearing loss [19]. These frequencies occur beyond the conventional speech range, often go unnoticed by patients and undetected by standard clinical audiometry. Furthermore, epidemiological and clinical evidence demonstrates a potential etiological association between migraine and certain cochlear disorders [20], notably Ménière’s disease [21, 22], and sudden sensorineural hearing loss [23]. In vivo evidence from nitroglycerin (NTG)-induced migraine animal models also supports auditory dysfunction, showing prolonged peak latencies [24] and elevated ABR thresholds [25]. Despite the evidence above, it remains unclear whether functional and especially whether structural abnormalities occur in the peripheral auditory organ during migraine attacks. The potential for shared underlying pathophysiology between these auditory manifestations and migraine warrants further investigation.

Converging evidence has established calcitonin gene-related peptide (CGRP) as the pivotal pathogenic mediator in migraine [3, 26, 27]. Its actions include promoting the release of proinflammatory mediators [28, 29], and facilitating glutamatergic neurotransmission through both pre- and postsynaptic mechanisms [30–32], thereby reinforcing nociceptive circuitry and promoting sensitization [33]. In the peripheral auditory system, CGRP expression shifts dynamically during development: it is present between type II afferents and outer hair cells (OHCs) pre-hearing [34], however it becomes concentrated in the inner hair cell after maturation [35]. Malfunction of this efferent CGRP signaling reduces sound-evoked cochlear nerve activity [36], while maturation of suprathreshold auditory responses depends on proper formation of the cochlear CGRP receptor complex [37]. The trigeminal nerve, a source of CGRP, also innervates cochlear vasculature, such as the stria vascularis [38], suggesting that CGRP from trigeminal ganglia terminals may also modulate cochlear vascular function [39, 40]. Despite these anatomical connections and functional observations, the specific contribution of cochlear CGRP to the auditory dysfunction experienced by individuals with migraine remains a crucial unresolved question.

In this research, we investigate the functional and especially the structural injuries of the peripheral auditory system in a mouse model exhibiting migraine-like features. We further investigate the role of CGRP elevation in the cochlea injuries in the context of migraine, and provide its pathophysiology of inflammatory cell death, aiming for the subclassification of migraine and its therapeutic strategies.

Materials and methods

Animals

C57BL/6J mice (6–8 weeks old, weighing between 18 and 22 g) were group-housed under a 12-hour light/dark cycle at a controlled room temperature of 23–25 °C with free access to food and water. All experimental procedures were approved by the Animal Research Committee of Huazhong University of Science and Technology and conducted in accordance with the guidelines to minimize animal suffering and reduce the number of animals used. Mice were randomly assigned to the experimental groups, with the sample size for each group detailed in Table 1.

Table 1.

Sample size for experimental group

Experimental objective	Group	Sample size (n)
Model validation & CGRP expression	VEH	5
	NTG	5
Rimegepant intervention	VEH	5
	NTG	5
	RIM + NTG	5
PANoptosis mechanism	VEH	4
	NTG	4
	RIM + NTG	4
NSA intervention	NTG	6
	NSA + NTG	6

VEH vehicle, NTG nitroglycerin, RIM rimegepant, NSA necrosulfonamide

Chronic migraine (CM) mouse model

A CM model was induced using an established protocol involving intraperitoneal (i.p.) injections of NTG (10 mg/kg) every second day for a total of five injections over a nine-day period, as previously described [41]. The NTG working solution (1 mg/mL) was freshly prepared from a commercial stock (10 mg/mL; T021, Sigma-Aldrich, USA) by diluting with 0.9% saline. Control animals received an equal volume of the saline vehicle only.

Drug administration

Rimegepant (BMS-927711, MCE, USA) was prepared as a solution in a vehicle consisting of 2% DMSO, 40% PEG300, 5% Tween80 and 53% normal saline, achieving a working concentration of 1 mg/mL, via brief sonication to ensure complete solubility. The compound was administered via oral gavage at a dose of 10 mg/kg, 30 min before each NTG injection.

Necrosulfonamide (NSA; HY-100573, MCE, USA) was dissolved in a mixture of 10% DMSO and 90% corn oil to a final concentration of 2 mg/mL. Mice received this formulation via i.p. injection at a dose of 20 mg/kg, 30 min before each NTG administration.

Behavioral tests

Mice were handled in the testing rooms for a minimum of 10 min twice daily over three consecutive days preceding behavioral assessment. Mice were returned to their home cage after each test.

Hot plate test

The hot plate test was conducted to measure reaction latencies before and after each NTG injection. Briefly, each mouse was placed on a hot plate (50 ± 1 °C), and the time until the first sign of nociception (e.g., paw licking, flinching, or jumping) was recorded [42]. A 50-second cut-off was implemented to prevent paw damage.

Von Frey test

The mechanical withdrawal threshold was assessed using the von Frey filament test, with calibrated filaments ranging from 0.04 g to 4 g. During testing, von Frey filaments were applied perpendicularly to the plantar surfaces of the hind paws. A positive response was defined as brisk paw withdrawal, flinching, lifting, or licking during stimulation or immediately after filament removal. The up-down method was employed, commencing with a 0.6 g filament. A negative response (O) prompted the use of the next stronger filament, while a positive response (X) led to the application of the next weaker one, with a minimum 30-second interval between stimuli. Withdrawal thresholds were automatically calculated using an online calculator [43].

Auditory function evaluation

ABR and DPOAE thresholds were recorded in mice anesthetized with 1.25% tribromoethanol (0.02 mL/g, i.p.) using a Tucker-Davis Technologies (TDT) System III. Upon confirmation of deep anesthesia (assessed by loss of the toe-pinch reflex), the animals were transferred to a sound-attenuated chamber, and their body temperature was maintained at 37℃ using a heating pad.

For ABR testing, subcutaneous electrodes were placed at the vertex (recording), the infra-auricular mastoid region (reference), and the back (ground). Acoustic stimuli were digitally generated by a calibrated MF1 speaker located 3 cm from the external auditory meatus. Tone bursts (32 –4 kHz) were presented in decreasing 5-dB steps from 90 dB SPL. Evoked potentials were recorded via electrodes using a Tucker-Davis Technologies (TDT) RA4LI head stage and RA16PA preamplifier. The outputs were then routed through a TDT RZ5 Multi I/O processor for additional amplification and digital conversion with responses averaged over 512 trials per condition across a 15 ms analysis window. The threshold was visually determined as the lowest sound level producing a reproducible ABR waveform.

DPOAE was assessed at center frequencies of 4, 8, 16, 24, and 32 kHz. Two primary tones, f1 and f2, were generated with a frequency ratio of f2/f1 = 1.2, geometrically centered around the audiometric frequency (where f1 = center frequency × 0.909 and f2 = center frequency×1.09). The primary tones were presented at equal intensities, decreasing from 80 dB SPL to 40 dB SPL in 10 dB decrements. The acoustic response was acquired via a single-channel microphone connected to the RZ6 In-A input. The DPOAE threshold was defined as the lowest sound level at which the amplitude of the cubic distortion product (2f1-f2) exceeded the noise floor by 5 dB. The average noise floor remained below 0 dB SPL across all tested frequencies.

Scanning electron microscope (SEM)

The organ of Corti was dissected and immediately fixed overnight at 4 °C in 2.5% glutaraldehyde in 0.1 M phosphate buffer. The samples were then post-fixed in 1% osmium tetroxide for 2 h at 4 °C, followed by dehydration using a graded ethanol series. Subsequent critical point drying was performed using a Leica EM CPD300. Dried specimens were mounted and sputter-coated with a thin layer of platinum. Imaging was carried out using a FEI Quanta250 field-emission scanning electron microscope. OHCs were identified based on their characteristic V-shaped stereociliary bundles and organized arrangement into three rows.

Cell culture and treatment

The HEI-OC1 cell line was acquired from Fuheng (Shanghai, China) and cultured in high-glucose Dulbecco’s modified Eagle’s medium (DMEM; C11960500BT, Gibco, China) supplemented with 5% fetal bovine serum (FBS; CF9901, Cybeking, China), and was maintained at 37 °C in a 5% CO₂ incubator for 24 h prior to experiments.

HEI-OC1 cells were transfected upon reaching 50–60% confluence in 6-well plates. The culture medium was replaced with 2 mL of DMEM, followed by the addition of a complex formed by mixing 5 µL of siRNA (final concentration: 20 nM) and 5 µL of LipoBooster 3000 Transfection Reagent (YEASEN, China) in 250 µL of Opti-MEM (P6038441, Gibco, USA). After a 24-hour incubation, the transfection medium was replaced with complete growth medium. The siRNA targeting Zbp1 (si-Zbp1) and the non-targeting negative control (si-NControl) were designed and synthesized by Weizhen Biosciences (China). The sequences are shown in Table 2.

Table 2.

Sequences of siRNA

Name	Sequences(5′→3′)
si-Zbp1-sense	GAGACAAUCUGGAGCAAAATT
si-Zbp1-antisense	UUUUGCUCCAGAUUGUCUCTT
si-NControl-sense	UUCUCCGAACGUGUCACGUTT
si-NControl-antisense	ACGUGACACGUUCGGAGAATT

Twenty-four hours after transfection, cells were stimulated with CGRP (1 µM; HY-F0203A, MCE, USA) for 24 h. Where applicable, treatment with Rimegepant (0.1 µM; BMS-927711, MCE, USA) was performed 30 min earlier than stimulation with CGRP.

Live cell imaging and analysis

HEI-OC1 cells were cultured in confocal dishes (BS-15-GJM, Biosharp) and stained with multiple fluorescent probes to visualize specific cellular components. Cells were sequentially stained with multiple fluorescent probes: MitoTracker Deep Red FM (200 nM; 40743ES50, Yeasen) for 30 min at 37 °C to visualize mitochondria, and MitoDNA-targeting Tracker (2 µM; MTD002, Biolight Biotechnology) for mitochondrial DNA (mtDNA) labeling for 30 min. The cell nuclei were counterstained with Hoechst 33,342 (C1022, Beyotime) for 30 min. After three washes with pre-warmed PBS, cells were imaged using a Zeiss LSM 800 laser scanning confocal microscope equipped with a 63× oil-immersion objective.

Mitochondrial morphology was quantitatively analyzed using the Mitochondria Analyzer plugin in ImageJ. After converting images to 8-bit grayscale and applying background subtraction, mitochondrial structures were isolated via automated thresholding and skeletonized. Key morphological parameters were then quantified, including the mean branch length, and the form factor (perimeter²/(4π × area), reflecting structural complexity).

Cell viability was assessed using a commercial Viability/Cytotoxicity Assay Kit (PF00008, Proteintech, China) containing Calcein-AM and ethidium homodimer-1 (EthD-1). After incubation with the dyes for 20 min according to the manufacturer’s instructions, metabolically active cells (green fluorescence) and membrane-compromised cells (red fluorescence) were simultaneously visualized by confocal microscopy.

Flow cytometry

HEI-OC1 cells were analyzed by flow cytometry according to the instructions of the Annexin V-FITC/ Propidium iodide (PI) Apoptosis Kit (C1062S, Beyotime, China). Harvested HEI-OC1 cells were washed, resuspended in binding buffer (1 × 10⁵ cells/195 µL), and incubated with 5 µL Annexin V-FITC and 10 µL PI for 15 min at room temperature in the dark. Samples were analyzed within 1 h of staining. The stained cells were analyzed by flow cytometry (Sony ID7000, Japan), and the apoptosis rate was analyzed using FlowJo software (Version 10.8.1).

Western blot

The cochleae were isolated after the mice were deeply anesthetized. Total protein from cochleae and cell lines was extracted using cold RIPA lysis buffer (P0013B, Beyotime) supplemented with phenylmethylsulfonyl fluoride (PMSF, ST506, Beyotime), protease and phosphatase inhibitors (P1045, Beyotime), followed by incubation for 30 min on ice and centrifugation at 12,000 × g for 20 min at 4 °C. Protein concentration was determined using a BCA Protein Assay Kit (P0010, Beyotime). Equal amounts of protein samples were denatured at 95 °C for 10 min and separated by 12.5% SDS-PAGE. Subsequently, proteins were transferred onto polyvinylidene difluoride (PVDF) membranes (Millipore). The membranes were blocked with 5% bovine serum albumin (BSA) for 1 h at room temperature and then incubated overnight at 4 °C with the primary antibodies listed in Table 3. After washing, the membranes were probed with HRP-conjugated secondary antibodies (listed in Table 3) for 1 h at room temperature. Protein signals were visualized using an enhanced chemiluminescence (SuperFemto ECL Chemiluminescence Kit, E423-02, Vazyme) and imaged with a chemiluminescence detection system. Quantification of band intensities was performed via ImageJ software.

Table 3.

Antibodies used in western blot

Antibody	Manufacturer	Catalog number	Host	Dilution
RAMP1	Proteintech	10327-1-AP	Rabbit	1:500
CALCRL	Sigma-Aldrich	HPA008070	Rabbit	1:2000
CGRP	Proteintech	68774-1-Ig	Mouse	1:1000
ZBP1	Santa Cruz	sc-271,483	Mouse	1:100
RIPK3	Santa Cruz	sc-374,639	Mouse	1:100
MLKL	Santa Cruz	sc-293,201	Mouse	1:100
Phospho-RIPK3 (S232)	Huabio	HA721428	Rabbit	1:1000
Phospho-MLKL(S345)	Huabio	ET1705-51	Rabbit	1:1000
Gasdermin D (N terminal)	Huabio	HA721144	Rabbit	1:1000
IL-1 beta	Huabio	HA601002	Mouse	1:2000
Caspase-8 (p18)	Huabio	HA722482	Mouse	1:2000
Cleaved & pro Caspase-3	Huabio	ET1608-64	Rabbit	1:1000
GAPDH	Proteintech	81640-5-RR	Rabbit	1:20000
HRP-conjugated Goat Anti-Rabbit IgG (H + L)	Proteintech	SA00001-2	Goat	1:10000
HRP-conjugated Goat Anti-Mouse IgG (H + L)	Proteintech	SA00001-1	Goat	1:10000

Immunofluorescence

Mice were anesthetized using 1.25% tribromoethanol, followed by transcardial perfusion with 0.01 M phosphate-buffered saline (PBS) and ice-cold 4% paraformaldehyde (PFA) for fixation. Following overnight fixation in 4% PFA at 4 °C, the tissues were dehydrated through a graded sucrose series. The fully dehydrated tissues were embedded in optimum cutting temperature compound (OCT; 4583, SAKURA) and rapidly frozen on dry ice. Serial coronal sections were then obtained and mounted onto glass slides. Frozen sections of the brain or cochlea, and the flat-mount of the organ of Corti were permeabilized with 1% Triton X-100 for 15 min and subsequently blocked with 5% BSA for 1 h at room temperature. For cell samples, cells grown on coverslips were fixed with 4% PFA for 10 min, permeabilized with 1% Triton X-100 for 10 min, and blocked with 5% BSA for 30 min. All specimens were incubated overnight at 4 °C with the primary antibodies specified in Table 4. The following day, samples were incubated with fluorescent secondary antibodies (listed in Table 4) for 1 h at room temperature. Following three washes with PBST, the sections were mounted onto glass slides with the application of a mounting medium containing DAPI (BL739A, Biosharp). Specimens were then visualized using a laser scanning confocal microscope (Zeiss LSM 800).

Table 4.

Antibodies used in Immunofluorescence staining

Antibody	Manufacturer	Catalog number	Host	Dilution
RAMP1	Proteintech	10327-1-AP	Rabbit	1:50
CGRP	CST	14959	Rabbit	1:200
CGRP	Proteintech	68774-1-Ig	Mouse	1:100
ZBP1	Santa Cruz	sc-271,483	Mouse	1:50
Phospho-MLKL(S345)	Huabio	ET1705-51	Rabbit	1:100
Gasdermin D (N terminal)	Huabio	HA721144	Rabbit	1:100
IL-1 beta	Huabio	HA601002	Mouse	1:200
Caspase-8 (p18)	Huabio	HA722482	Mouse	1:100
Cleaved & pro Caspase-3	Huabio	ET1608-64	Rabbit	1:1000
Z-DNA	AbinScience	YP373023	Mouse	1:100
Phalloidin	Sigma-Aldrich	P5282	/	1:500
c-Fos	Synaptic Systems	226 017	Rat	1:2000
Sox2	Huabio	HA601396	Rat	1:200
IBA1	Immunoway	ym4765	Mouse	1:200
Myosin Ⅶa	Proteus BioScience	25-6790	Rabbit	1:200
Alexa Fluor 488 Donkey anti Rabbit IgG	AntGene	ANT024	Donkey	1:500
Alexa Fluor 647 Donkey anti Mouse IgG	AntGene	ANT034	Donkey	1:500
FITC Goat anti Rat IgG	ABclonal	AS019	Goat	1:500

RNA-seq

Cochleae were rapidly dissected from euthanized mice and immediately stabilized in RNAlater (R0118, Beyotime). Each group consisted of 4 or 5 mice. RNA sequencing was performed at Chi-Biotech (Shenzhen, China). Total RNA was extracted from the collected tissues using the TRIzol Reagent protocol. Fragments of 200–300 bp were size-selected using magnetic beads and PCR amplified to generate the final sequencing libraries. Library quality was assessed using a Qubit 2.0 Fluorometer and an Agilent 4150 system. The qualified libraries were sequenced on an Illumina platform with a PE150 strategy. Gene expression levels were normalized and quantified as transcripts per kilobase of exon model per million mapped reads (TPM). Genes with an adjusted p-value < 0.05 and |log2 (fold change)| > 1 were considered differentially expressed genes (DEGs). Functional categorization was carried out using the KEGG pathways and GO databases between different groups.

Statistical analysis

Data are presented as the mean ± standard deviation (SD) from at least three independent experiments, unless otherwise stated. Statistical analyses were performed using GraphPad Prism software (Version 8.0). For comparisons between two groups, a two-tailed, unpaired Student’s t-test was applied. For comparisons among more than two groups, one-way or two-way analysis of variance (ANOVA) was employed, followed by Tukey’s post hoc test for pairwise comparisons between all groups. A p-value of less than 0.05 was considered statistically significant.

Results

CGRP signaling is elevated in cochlear OHCs in the NTG-induced CM model

A CM mouse model was established by intraperitoneal injections of NTG once every other day over a nine-day period. Hind paw mechanical and thermal pain thresholds were measured before (basal response) and two hours after each injection (post-treatment response; Fig. 1A). To validate the successful establishment of the CM model, we first assessed its core behavioral phenotype: mechanical (Fig. 1B) and thermal (Fig. 1E) pain thresholds. In the vehicle (VEH) group, basal and post-treatment pain thresholds did not change during a nine-day period. In the NTG injection group, the basal mechanical and thermal pain thresholds on the first day were normal, but gradually decreased over the following days, suggesting the development of chronic hypersensitivity (Fig. 1C, F). An acute hypersensitivity response was also observed two hours after NTG administration on day 1 and day 3, evidenced by a pronounced decrease in mechanical and thermal pain thresholds (Fig. 1D, G). Given the established role of the trigeminal nucleus caudalis (TNC) in processing sensation and its central role in CM-related sensitization [44], we further investigated markers of neuronal activation and sensitization in this region. Immunofluorescence analysis confirmed a significant upregulation of c-Fos-positive cells and CGRP expression in TNC tissue on day 9 after NTG injections (Fig. 1H, I).

Fig. 1.

CGRP signaling is elevated in cochlear OHCs in the NTG-induced CM model. A. The schematic diagram showing CM mouse model establishment and behavior studies’ protocol. Cochlea and TNC tissue were collected after the last behavior tests on day nine. B-G. The hind paw mechanical (B-D) and thermal (E-G) thresholds before (basal) and two hours after each NTG injection (post-treatment) were gradually decreased during nine-day NTG injection procedure, evaluated by Von Frey test and hot plate test, respectively (n = 5 per group). H-I. C-Fos-positive neurons and CGRP expression were increased in TNC after nine-day NTG treatment, revealed by immunostaining of c-Fos and CGRP. Scale bar: 500 μm (left panel), 20 μm (right panel; n = 4 per group). J. CGRP expression in the OHCs was increased after nine-day NTG treatment. Representative immunostaining images showing CGRP, phalloidin and DAPI in the organ of Corti. Scale bar: 20 μm. K-L. CGRP signaling was increased in the cochlea after nine-day NTG treatment, revealed by western blotting images of proteins (K) and densitometric quantification (L) for CGRP and its receptors, RAMP1 and CALCRL, GAPDH was used as the loading control. (n = 3 per group). Statistical analysis was carried out via two-way ANOVA for B-G, two-tailed Student’s t-test for I and L, *p < 0.05, **p < 0.01, ***p < 0.001

To investigate the role of CGRP signaling in the cochleae, we first observed the local expression of CGRP and its receptors in the inner ear. Western blotting analysis revealed that CGRP and its receptors (receptor activity-modifying protein 1, RAMP1; calcitonin receptor-like receptor, CALCRL) were constitutively expressed in cochlea, and their expression levels were upregulated after NTG treatment (Fig. 1K, L). Further immunostaining localized CGRP immunoreactivity to the peri-inner hair cell region. Notably, NTG treatment induced a marked increase in CGRP signaling intensity, especially within the OHCs (Fig. 1J). The predominance of CGRP signaling activation in OHCs was confirmed using an alternative CGRP antibody from a different host species (Supplementary Fig. 1A), which verified the co-localization of RAMP1 and CGRP in OHCs of NTG-treated mice. Furthermore, a modest increase in the expression of both CGRP and RAMP1 was also detected within spiral ganglia in NTG-treated mice, though baseline expression of RAMP1 was observed in the VEH group (Supplementary Fig. 1B). These results demonstrate a specific elevation of CGRP signaling in cochlea OHCs in CM model, implying a potential role for CGRP-mediated mechanisms in peripheral auditory processing in migraine pathophysiology.

The CGRP antagonist Rimegepant ameliorates auditory dysfunction and OHC injury in the CM model

To evaluate the function of CGRP signaling within the inner ear in the context of migraine, a potent and selective CGRP receptor antagonist Rimegepant [45], was administered thirty minutes prior to each NTG injection (NTG + RIM group, Fig. 2A). CM-associated mechanical and thermal pain hypersensitivities were partially reversed by Rimegepant (Supplementary Fig. 1C-F). Furthermore, NTG-induced auditory dysfunction, evidenced by elevated ABR thresholds measured on the last day of NTG injection (Fig. 2B-D), was effectively reversed by Rimegepant (Fig. 2C, D). Consistent with the ABR threshold shifts, the NTG group exhibited a significant elevation in DPOAE thresholds compared to the VEH group, confirming dysfunction of the outer hair cells (Fig. 2E, F). Subsequent morphological integrity analysis by SEM revealed that OHC pathologies in the NTG group, including stereocilia degeneration, structural disarray, and hair cell loss (Fig. 2G), were also partially rescued by Rimegepant treatment. Together, our findings suggest functional hearing impairment and structural damages of hair cells in this CM model might be mediated by overactivation of CGRP signaling in the inner ear.

Fig. 2.

CGRP antagonist Rimegepant ameliorates auditory dysfunction and OHC injury in the CM model. A-B. Schematic illustration of Rimegepant intervention and hearing detection protocol. Rimegepant was administered via oral gavage thirty minutes prior to each NTG injection. Auditory function was detected on the day 9. C-D. NTG-induced hearing threshold elevation rescued by Rimegepant administration, evaluated by pure tone ABR across 4, 8, 12, 16, 24, and 32 kHz (D). Representative ABR wave recordings (C) to 24 kHz were shown, and the bold blue, red, and green lines represent the thresholds of each group (n = 5 mice per group). E-F. NTG-induced OHC dysfunction rescued by Rimegepant administration, evaluated by DPOAE across 4, 8, 16, 24, and 32 kHz (F). Representative DPOAE amplitude spectrum (E) measured at center frequency of 24 kHz were shown, and the red lines indicate the distortion product peak at 2f1-f2 (n = 6 mice per group). Statistical analysis was carried out via two-way ANOVA, *p < 0.05, **p < 0.01, ***p < 0.001 compared between NTG versus NTG + RIM groups; ^#p < 0.05, ^##p < 0.01, ^###p < 0.001 compared between NTG and VEH groups). G. NTG-induced OHC injury rescued by Rimegepant, revealed by scanning electron microscope imaging of the organ of Corti. Scale bar: 20 μm (upper panel), and 10 μm (lower panel)

CGRP signaling activation in OHCs induces mitochondrial dysfunction and subsequent immune response

To explore the potential downstream mediators of CGRP signaling, we employed RNA sequencing to map the molecular landscape in the whole cochlea tissue (Fig. 3A). Multi-group volcano plot illustrates the comparative analysis of differentially expressed genes (DEGs) in the cochleae of NTG-induced CM model, compared with both the VEH and RIM intervention groups (Fig. 3B). The multi-group VENN analysis revealed distinct sets of co-regulated genes (Fig. 3C). A core set of 46 genes was consistently upregulated in the NTG group compared to both the VEH and the NTG + RIM groups. This upregulated gene set was predominantly enriched in the interferon-inducible gene (IFI) family, interferon regulatory factor (IRF) family, key inflammatory chemokines such as Ccl8 and Gzmb, and the innate immune sensor Zbp1. Conversely, 21 genes were consistently downregulated, which were primarily associated with ciliary structure and function, such as Spag16, Dnah3 and Tmc5. The heatmap details the expression patterns of 46 upregulated and 21 downregulated genes in the NTG group (Supplementary Fig. 2A, B).

Fig. 3.

CGRP signaling activation in OHCs induces mitochondrial dysfunction and subsequent immune response. A. Schematic diagram of RNA-seq and further experiment design. B. Volcano plot displaying mRNA expression differences between the vehicle, NTG, and RIM + NTG groups. Genes with an adjusted p-value < 0.05 and |log2 (fold change)| > 1 were shown. C. Venn diagrams showing the overlap upregulated and downregulated genes between NTG vs. VEH and NTG vs. RIM + NTG groups. D-E. Bubble diagram of biological process among overlapping upregulated genes (D) and downregulated genes (E). Red arrowheads showed immune and stress processes, green arrowheads showed cilium-related process. F. NTG-induced macrophage activation in the cochleae rescued by Rimegepant administration, revealed by immunostaining of IBA1 in mid-modiolar cochlear sections. White arrows show infiltration of macrophages. Scale bar: 100 μm. G-K. Rimegepant alleviated CGRP-induced HEI-OC1 cell mitochondria fragmentation and mtDNA leakage. Mitochondrial morphology was visualized by MitoTracker staining (G), mtDNA leakage was assessed by co-immunostaining of mtDNA and MitoTracker (H); white arrows indicate extramitochondrial mtDNA, and an arrowhead indicates a dead cell. Scale bars: 10 μm. Quantitative analyses of mitochondrial form factor (I), branch length (J), and the mtDNA leakage ratio (K) are shown. For I and J, n = 13 (Control), 17 (CGRP), 13 (CGRP + RIM); for K, n = 10 per group. One-way ANOVA was used for statistical analysis, *p < 0.05, **p < 0.01, ***p < 0.001

To functionally characterize the 46-core upregulated gene set, a GO biological process enrichment analysis was conducted. The results indicated that these genes are primarily involved in immune and inflammatory processes, notably including ‘cellular response to cytokine stimulus’ and ‘immune response’, along with processes related to cellular stress (e.g. response to stress and response to biotic stimulus), thereby underscoring the pivotal processes in the pathogenesis of NTG-induced, CGRP-related cochlea injury in CM model (Fig. 3D). Intriguingly, the 21 overlapping downregulated genes were predominantly enriched in cilium-related processes (Fig. 3E), such as cilium assembly and cilium movement. This molecular profile is consistent with our previous SEM observations, which revealed disorganized cilium architecture in the cochleae of NTG mice. KEGG pathway analysis of the 67 overlapping genes revealed significant enrichment in pathways related to neurodegeneration and inflammation, notably the NOD-like receptor signaling pathway (Supplementary Fig. 2C).

Macrophages play a pivotal role in the immune responses and inflammatory processes of the inner ear [46, 47]. To confirm the elevation of immune responses, IBA1 immunofluorescence staining was performed on cochlear sections (Fig. 3F). Compared to the VEH and RIM groups, the NTG group exhibited an increase in the number of IBA1-positive macrophages and an infiltration pattern as the macrophages penetrated the basilar membrane and accumulated around the stria vascularis.

Mitochondria, as central mediators of the cellular stress response, link morphological integrity directly to metabolic health and viability [48, 49]. To validate the stress-response pathways identified in our GO analysis, we employed an in vitro model of HEI-OC1 cells. Mitochondrial status was visually assessed using MitoTracker staining (Fig. 3G). Subsequent to CGRP stimulation, we observed a pronounced mitochondrial injury, characterized by a rounded, shrunken (Fig. 3I), and fragmented morphology (Fig. 3J). These CGRP-induced morphological alterations, indicative of extensive mitochondrial fission—a process known to facilitate programmed cell death signaling pathways, were markedly alleviated by pretreatment with Rimegepant. Further corroborating the induction of a severe cellular stress state, we detected the leakage of mitochondrial DNA (mtDNA) into the cytosol (Fig. 3H, K). Cytosolic mtDNA acts as a potent damage-associated molecular pattern that can initiate innate immune and inflammatory signaling pathways [50]. Notably, CGRP-induced leakage of mtDNA was all reversed by Rimegepant. Taken together, our findings demonstrate that cochlear CGRP signaling activation initiates a cellular stress response, which leads to mitochondrial fragmentation and the subsequent release of mtDNA into the cytosol, thereby triggering a downstream innate immune cascade.

ZBP1 expression was significantly upregulated in both the OHCs of CM model and CGRP-stimulated HEI-OC1 cells

To systematically investigate the molecular mechanism of CGRP-mediated hair cell damage, we performed Gene Set Enrichment Analysis (GSEA). Based on our evidence of CGRP-triggered mtDNA leakage and innate immunity, we prioritized the nucleic acid binding gene set, reasoning that it functionally links cytosolic nucleic acid sensing to immune pathology (Fig. 4A). Among the upregulated DEGs related to nucleic acid binding in the NTG group, ZBP1, a well-known regulator of both an innate immune sensor and programmed cell death [51, 52], emerged as a noteworthy candidate. Oxidative stress and mitochondrial genome instability potentiate the conversion of mtDNA into its Z-form, which can be recognized by ZBP1 [53, 54]. CGRP stimulation elevated cytosolic Z-form DNA in HEI-OC1 cells, an effect attenuated by Rimegepant co-treatment (Fig. 4B). Consistently, CGRP upregulated ZBP1 expression and activated its own signaling in HEI-OC1 cells, which was normalized by Rimegepant, revealed by western blot (Fig. 4C, D) and immunofluorescence (Fig. 4E). These results indicate that mtDNA-derived Z-form DNA specifically trigger ZBP1 activation in vitro.

Fig. 4.

ZBP1 expression was significantly upregulated in the OHCs of CM model and CGRP-stimulated HEI-OC1 cells. A. NTG-induced upregulation of genes related to nucleic acid binding, revealed by GSEA enrichment and heatmap. B. Rimegepant alleviated CGRP-stimulated Z-DNA elevation, showed by immunostaining of Z-DNA in HEI-OC1 cells, scale bar: 50 μm C-D. Rimegepant mitigated CGRP signaling activation and the elevated expression of ZBP1 in CGRP-stimulated HEI-OC1 cells, assessed by western blotting of proteins (C) and densitometric quantification (D) (n = 3 per group). GAPDH was used as the loading control. E. Rimegepant rescued CGRP signaling activation and ZBP1 upregulation in CGRP-stimulated HEI-OC1 cells, visualized by immunostaining of ZBP1and RAMP1, scale bar: 10 μm. F. Rimegepant alleviated NTG-induced Z-DNA accumulation in the OHCs. Representative immunofluorescence images showing the co-localization of Z-DNA and the hair cell marker Myosin VIIa in the organ of Corti, scale bar: 20 μm. G. NTG-induced ZBP1 upregulation in cochleae rescued by Rimegepant administration, revealed by western blotting of proteins and densitometric quantification for ZBP1, GAPDH was used as the loading control. H-K. Immunostaining (H) and quantitative analyses (J, K) demonstrated that Rimegepant treatment reversed the loss of OHC and the predominant upregulation of ZBP1 in OHCs in CM mouse model. OHCs corresponding to 8, 16, and 32 kHz sensations in the organ of Corti were assessed (I). Representative images of Myosin VIIa and ZBP1-positive OHCs illustrate these findings (H), with pentagrams indicating OHC loss, scale bar: 20 μm; n = 4 per group, two-way ANOVA, *p < 0.05, **p < 0.01, ***p < 0.001

To further validate the downstream of CGRP signaling in vivo, we performed immunofluorescence staining on the organ of Corti. Consistent with our in vitro findings, we observed an accumulation of Z-DNA in the cytoplasm of OHCs of NTG-treated mice (Fig. 4F). This cytosolic Z-DNA accumulation triggered upregulation of ZBP1, which was effectively reversed by Rimegepant (Fig. 4G). Importantly, ZBP1 immunoreactivity was predominantly in OHCs (Fig. 4H, K), not in neighboring supporting cells (Supplementary Fig. 3A) or the spiral ganglia (Supplementary Fig. 3B). The function of OHCs were quantified at cochlear frequency regions corresponding to approximately 8, 16, and 32 kHz (Fig. 4I). Compared to the VEH and NTG + RIM groups, the NTG group exhibited a significant loss of OHCs across these frequencies (Fig. 4H, J). This cell-type-predominant upregulation of ZBP1 in OHCs, coinciding spatially with the observed ABR and DPOAE thresholds shift and structure injury in Fig. 2, nominates ZBP1 as a crucial and specialized molecular player in the cascade of CGRP-mediated peripheral auditory deficits.

CGRP signaling blockade or ZBP1 knockdown ameliorates OHC death in vitro by inhibiting the PANoptosis pathway

Given the establishment of ZBP1 as a master regulator of programmed cell death [55], we conducted flow cytometric analysis with Annexin V/ PI staining in HEI-OC1 cells (Fig. 5A) to delineate the molecular pathway of CGRP-induced injury. The knockdown efficiency of ZBP1 siRNA was confirmed at both the mRNA and protein levels (Supplementary Fig. 2D-F). Quantitative results revealed that treatment with Rimegepant and ZBP1-siRNA attenuated CGRP-induced necroptosis and modestly reduced late apoptosis and pyroptosis. Correspondingly, TUNEL staining further confirmed that both CGRP antagonism and ZBP1 inhibition effectively suppressed apoptotic and pyroptotic pathways (Fig. 5B, D). Morphological assessment via EthD-1 staining further demonstrated a reduction in necroptotic cell death upon CGRP or ZBP1 blockade (Fig. 5C, D).

Fig. 5.

CGRP signaling blockade or ZBP1 knockdown ameliorates OHC death in vitro by inhibiting the PANoptosis pathway. A. Rimegepant or ZBP1 knockdown alleviated CGRP-stimulated cell death in HEI-OC1 cells, revealed by flow cytometry analysis of Annexin V-FITC/PI staining. B–D. Immunostaining of TUNEL (B) and EthD-1/ Calcein (C), and quantitative analyses (D) demonstrated that Rimegepant treatment or ZBP1 knockdown alleviated CGRP-triggered cell death in HEI-OC1. The arrows indicating TUNEL+ or dead cells in B and C, respectively. Scale bars: 50 μm (n = 3 biologically independent experiments). E–J. Rimegepant treatment or ZBP1 knockdown mitigated CGRP-activated PANoptosis signaling in HEI-OC1 cells, evaluated by western blotting of proteins and densitometric quantification for pyroptosis (E, H), necroptosis (F, I), and apoptosis (G, J) markers (n = 3 biologically independent experiments). K–M. CGRP stimulated PANoptosis signaling in HEI-OC1 was rescued by Rimegepant treatment or ZBP1 knockdown, evaluated by immunostaining of GSDMD-N terminal and IL-1β (K), p-MLKL (L), Caspase-8 (M), with white arrows indicating colocalization of IL-1β and GSDMD-N terminal (K), p-MLKL activation in the cell membrane (L), scale bar: 50 μm. Statistical analysis was carried out via two-way ANOVA, *p < 0.05, **p < 0.01, ***p < 0.001

Based on the concurrent activation of apoptosis, pyroptosis, and necroptosis by CGRP-ZBP1 signaling, we postulated the engagement of PANoptosis—an integrated inflammatory cell death pathway. To substantiate this hypothesis, we investigated the key molecular components of PANoptosis. Inhibition of CGRP or ZBP1 significantly suppressed the expression of key mediators across all three programmed cell death pathways: the GSDMD N-terminal fragment (GSDMD-N) and mature IL-1β for pyroptosis (Fig. 5E, H), phosphorylated RIPK3 (p-RIPK3) and phosphorylated MLKL (p-MLKL) for necroptosis (Fig. 5F, I), and cleaved Caspase-8 and Caspase-3 for apoptosis (Fig. 5G, J).

Confocal immunofluorescence imaging confirmed these results by revealing the co-localization of IL-1β and GSDMD-N in HEI-OC1 cells following CGRP challenge, indicative of pyroptotic pore formation. This effect was abolished by Rimegepant or ZBP1 knockdown (Fig. 5K). Furthermore, immunostaining for p-MLKL demonstrated that CGRP stimulation elevated p-MLKL levels and promoted its translocation to the cell membrane, a hallmark of necroptosis execution [56], which was mitigated by CGRP blockade or ZBP1 knockdown (Fig. 5L). Additionally, a pronounced decrease in cleaved Caspase-8-positive cells upon CGRP antagonism or ZBP1 knockdown indicated suppression of apoptotic initiation (Fig. 5M). Collectively, our in vitro findings demonstrate that CGRP overstimulation might initiate a ZBP1-mediated PANoptosis pathway, an integrated cell death process encompassing apoptosis, pyroptosis, and necroptosis, which ultimately leads to the injury of HEI-OC1 cells.

Pharmacological inhibition of PANoptosis signaling protects against OHC injury and hearing loss in CM model

Having established that CGRP has triggered PANoptosis signaling in HEI-OC1 cells in vitro, we next sought to determine whether this integrated cell death pathway contributes to OHC injury in CM model in vivo. Consistent with our in vitro findings, the protein levels of pyroptosis markers (GSDMD-N and mature IL-1β; Fig. 6A, D), necroptosis markers (p-RIPK3 and p-MLKL; Fig. 6B, E), and apoptosis markers (cleaved Caspase-8 and cleaved Caspase-3; Fig. 6C, F) were significantly elevated in the inner ears of NTG mice compared to both the VEH and RIM-treated groups. Immunofluorescence staining further confirmed in vivo that pharmacological antagonism of CGRP effectively attenuated the expression of these key executioner proteins specifically within outer hair cells (Fig. 6G, H), confirming the involvement of CGRP-driven PANoptosis in OHCs in the CM model.

Fig. 6.

Pharmacological inhibition of PANoptosis signaling protects against OHC injury and hearing loss in CM model. A-F. NTG-induced PANoptosis in the cochleae rescued by Rimegepant, evaluated by western blotting of proteins and densitometric quantification for pyroptosis (A, D), necroptosis (B, E), apoptosis (C, F) markers (n = 4 mice per group). G-H. Immunostaining images demonstrated that Rimegepant treatment rescued PANoptosis signaling activation in the organ of Corti of CM model. Representative images of GSDMD-N terminal and IL-1β (G), p-MLKL and Caspase-8 (H), were shown, with white arrows indicating p-MLKL activation in cell membrane, scale bar: 20 μm. I. Schematic illustration of NSA and hearing loss detection protocol. NSA was administered via intraperitoneal injection thirty minutes prior to each NTG injection, auditory function was detected on day 9 (post treatment). J-L. Hearing threshold elevation in CM mouse model relieved by NSA administration, evaluated by pure tone ABR (K) across 4, 8, 12, 16, 24, and 32 kHz (n = 5 mice per group), ABR wave recordings (J) to 24 kHz in post treatment were shown, and the bold lines represent the thresholds of two groups. DPOAE threshold (L) across 4, 8, 16, 24, and 32 kHz (n = 6 mice per group) were shown. M-N. Immunostaining of phalloidin at different frequencies (M) and quantitative analyses (N) demonstrated that NSA administration rescued the loss of OHCs in CM model. Pentagrams indicate OHCs loss, scale bar: 20 μm (n = 4 mice per group). Statistical analysis was carried out via two-way ANOVA, *p < 0.05, **p < 0.01, ***p < 0.001

Given the evidence of PANoptosis in OHCs, we hypothesized that its inhibition could alleviate cell injury and preserve hearing function. Necrosulfonamide (NSA) is a small-molecule inhibitor that directly targets the key executioners of inflammatory cell death: it blocks oligomerization of MLKL in necroptosis [57] and pore formation by the GSDMD-N fragment in pyroptosis [58]. We administered NSA via intraperitoneal injection 30 min prior to each NTG administration. After auditory measurement on day 9 to monitor hearing function, cochleae were collected for morphological analysis (Fig. 6I). NSA treatment markedly suppressed the NTG-induced ABR threshold shifts (NTG-post vs. NSA + NTG-post), particularly at the frequencies of 4, 8, and 24 kHz (Fig. 6J, K). Consistent with the ABR findings, NSA administration attenuated the NTG-induced elevation of DPOAE thresholds, with the most pronounced protective effects observed at 16 and 24 kHz (Fig. 6L). This functional rescue was corroborated by morphological analysis: Phalloidin staining of the hair cell cytoskeleton demonstrated that NSA effectively prevented OHC loss (Fig. 6M, N). These results demonstrate that partial inhibition of the PANoptosis pathway conferred a substantial therapeutic benefit against CM-induced peripheral auditory processing deficits.

Discussion

We elucidate a potential pathogenic pathway that bridges the clinical comorbidity between migraine and auditory dysfunction, from the initial migraine trigger to the final occurrence of hair cell injury. Our key findings delineate a pathogenic cascade in the CM model: (1) Aberrant CGRP signaling predominantly upregulates in OHCs; (2) CGRP overstimulation initiates mitochondrial stress, culminating in fragmentation and cytosolic release of mtDNA, and importantly, induces both structural and functional deficits in OHCs; (3) The liberated mtDNA undergoes conversion to Z-DNA, functioning as the critical ligand for ZBP1 activation; (4) Activated ZBP1 coordinates pyroptosis, apoptosis, and necroptosis, underlying OHC loss and functional hearing deficits; (5) Inhibition of CGRP-ZBP1-PANoptosis axis reversed or alleviated OHC deficits.

Aberrant CGRP upregulation in the sentinel OHCs

The comorbidity between migraine and auditory symptoms suggests the existence of shared CGRP-mediated pathological mechanisms. Previous research has established that CGRP is transiently expressed between type II afferents and OHCs in the pre-hearing cochlea [34], with its expression declining to a sparse immunoreactivity beneath IHCs in the adult murine cochlea [36]. Studies in αCGRP-null mice have establish that CGRP is critical for synaptic transmission between inner hair cells and auditory nerve fibers, as indicated by reduced ABR wave I amplitudes [37], without affecting cochlear mechanical function mediated by outer hair cells [36]. In our study, we confirm that CGRP expression in the adult mouse cochlea is normally restricted to a low basal expression level around IHCs. However, in NTG-induced CM mice, we found CGRP labeling was significantly upregulated in the OHCs, indicating CGRP signaling re-emerges as a potent modulator of OHC function under stress conditions in the mature auditory system. OHCs act as biological regulators in the auditory transduction process, responsible for fine-tuning hearing thresholds and frequency selectivity [59–61]. OHCs are vulnerable to ototoxic insults, positioning them as potential stress receptors [62]. OHCs receive dual innervation from both medial olivocochlear efferent fibers and type II afferent neurons [63, 64]. The efferent system is known to modulate OHC activity and provide protection against various ototoxic challenges [65, 66]. Type II afferents are thought to serve as cochlear nociceptors that detect damage-related metabolites like ATP to signal acoustic pain [67, 68]. In our study, we reinforce OHCs’ role as nociceptive receptors in CM-induced CGRP-mediated peripheral hearing impairment. OHCs emerge as the cochlear frontline sentinels due to their dual burden of direct contact with pathogenic mediators and high intrinsic stress.

CGRP: a neural-immune bridge in cochlear pathology

Our data demonstrate that CGRP’s toxicity extends beyond vascular effects to directly instigate an inflammatory signaling cascade within the sensory hair cells themselves. In migraine, CGRP exhibits dual pathogenicity through peripheral neuroinflammatory actions and direct central neuromodulation [32]. In peripheral sensitization, CGRP serves as a critical initiator and sustainer of pro-inflammatory signaling by vasodilation and immune cell activation [69–71]. In central sensitization, CGRP enhances synaptic transmission efficacy as a neuromodulator, its receptors colocalize with post-synaptic AMPA receptors [72], potentiating their responsiveness to glutamatergic input [73]. Our data show that excessive CGRP signaling directly causes OHC stress response, mainly through mitochondrial network collapse. Moreover, we found that the subsequent release of damage-associated molecular patterns, following the CGRP-mediated mitochondrial fragmentation in hair cells, activated innate immune responses within the cochlea, ultimately driving the transition from cellular stress to peripheral auditory inflammation. This demonstrates that cochlea is implicated as a direct target organ in migraine, where CGRP functions as a bridge between neural signals and neuroinflammatory pathways within the sensory hair cells, converting neuronal activity into immune-inflammatory responses in cochlea under migraine.

ZBP1-mediated PANoptosis: an integrative mechanism for hearing loss

Our study identifies ZBP1-mediated PANoptosis as the critical mechanism integrating neuroinflammation, OHC loss, and functional auditory deficits in the CM model. ZBP1 is a key innate immune sensor that detects Z-form nucleic acids from various sources, including viruses [74], endogenous elements [75, 76], and—as highlighted by our findings—stressed mitochondria [54, 77]. It acts as a central integrator of inflammatory and cell death signaling [76], driving PANoptosis: a combined apoptosis, pyroptosis and necroptosis mechanism [78]. Accumulating evidence implicates ZBP1-mediated PANoptosis in the pathogenesis of several neuroinflammatory and degenerative diseases, such as Alzheimer’s disease [79], spinal cord injury [80], and Parkinson’s disease [81]. In our CM model, CGRP-induced mitochondrial fragmentation and subsequent mtDNA release activated ZBP1, which was predominantly upregulated in OHCs. This led to the activation of PANoptosis-associated signaling, characterized by elevated levels of phosphorylated RIPK3 and Caspase-8, GSDMD (pyroptosis), cleaved Caspase-3 (apoptosis), and p-MLKL (necroptosis) in OHCs. This mechanism is consistent with recent work implicating ZBP1 and PANoptosis in other ototoxic models, such as neomycin-induced hearing loss [82], suggesting it may represent a common final pathway in OHC damage. The partial rescue of auditory function in CM mice through administration of NSA, which concurrently inhibits key executors of pyroptosis and necroptosis, provides direct evidence for the therapeutic relevance of targeting PANoptosis in hearing preservation.

Migraine-related auditory deficits: clinical guidance

Previous study showed pharmacological management of migraine may be effective in treating various otologic symptoms, including calcium channel blockers, selective serotonin reuptake inhibitors, gepants and monoclonal antibodies targeting CGRP signaling pathway [83]. Our findings establish Rimegepant as an effective therapeutic agent that not only rescues hearing function but also ameliorates stress-related damage in OHCs within the migraine pathological context, presenting new opportunities for interdisciplinary management of auditory symptoms in otolaryngological practice. The targeted modulation of CGRP signaling appears particularly promising for alleviating the peripheral cochlear dysfunction identified in our investigation. Previous reports on migraine-induced hearing impairment have shown inconsistencies in the affected frequency ranges from low (0.5 kHz) to high frequency (16 kHz) [18, 19, 84]. Some auditory damage in migraine patients may easily go undetected in daily life [19]. In this study, we reveal moderate full-frequency hearing impairment in migraine mice, mainly focusing on peripheral auditory system. Our findings support the implementation of routine auditory screening in migraine management for patients with migraine-related auditory dysfunction, thereby highlighting the importance of hearing assessment in migraine patients. We demonstrate that the CGRP-ZBP1-PANoptosis axis is actively engaged within the cochlea, establishing OHC as a direct target of migraine-related pathology.

Limitations and future directions

Several limitations of our study should be acknowledged. First, while the NTG-induced CM model is well-validated, it may not fully recapitulate the complex etiology of human migraine. Second, the specific origin of CGRP-containing fibers innervating OHCs—whether from peripheral type II afferents or central medial olivocochlear efferents—remains unresolved. This distinction is functionally important, as these systems likely subserve different roles in OHC modulation and pathology. Future studies using neuronal tracing, selective denervation, or cell-specific knockout models will be necessary to elucidate the source of CGRP signaling and its contribution to OHC damage. Third, our study focused predominantly on OHCs; the potential vulnerability of inner hair cells or spiral ganglia neurons in migraine-related auditory deficits merits further exploration. Indeed, consistent with the multicellular complexity of the cochlea, our transcriptomic analysis reflected an aggregate stress response extending beyond the hair cells. We observed significant downregulation in Neuroactive ligand-receptor interaction, suggesting concurrent pathological alterations in spiral ganglia or synaptic transmission. Notably, the enrichment of vascular smooth muscle contraction implies compromised cochlear blood flow regulation or stria vascularis dysfunction, which aligns well with the neurovascular etiology of migraine. Finally, the translational potential of PANoptosis inhibition should be evaluated in other ototoxic and age-related hearing loss models to assess its broader therapeutic applicability.

Conclusion and clinical outlook

In summary, our results demonstrate that migraine pathophysiology can directly engage the peripheral auditory system via a CGRP-driven, ZBP1-mediated PANoptosis pathway in OHCs. This mechanism might provide a molecular basis for the clinical concept of “cochlear migraine” and argues for greater integration of neurological and otological perspectives in managing patients with comorbid migraine and auditory symptoms. Our findings support the implementation of routine auditory screening in migraine management and the exploration of CGRP-targeted therapies for patients with migraine-related auditory dysfunction. Looking forward, future research should focus on auditory biomarkers in human patients, dissecting the central-peripheral interactions in migraine-related auditory processing, and developing targeted neuroprotective strategies aimed at preserving long-term hearing function.

Supplementary Information

Below is the link to the electronic supplementary material.

Abbreviations

CM

Chronic migraine

NTG

Nitroglycerin

OHCs

Outer hair cells

CGRP

Calcitonin gene-related peptide

TNC

Trigeminal nucleus caudalis

NSA

Necrosulfonamide

ABR

Auditory brainstem response

DPOAE

Distortion product otoacoustic emissions

CALCRL

Calcitonin receptor-like receptor

RAMP1

Receptor activity-modifying protein 1

Author contributions

The study was designed by SLZ, YSL and WDX. The in vitro experiments were completed by WDX and NZ. The in vivo experiments were completed by WDX. Tissue collection of TNC and cochleae were carried out by WDX, JYC and SZ. WDX and NZ performed statistical analysis and finished writing the manuscript. XM, YXW, SLZ, YSL provided supervision and final check. All authors discussed the results and participated in manuscript preparation and editing.

Funding

This work was supported by grants from the National Key Research and Development Program of China (2023YFC2508403, 2023YFC2508002), the National Natural Science Foundation of China (82371168 and 82171152), Beijing Natural Science Foundation (7252145).

Data availability

All data generated in this study are included in this article. Source data are provided with this paper or upon request from the corresponding authors.

Declarations

Ethics approval and consent to participate

All experimental procedures were approved by the Animal Research Committee of Huazhong University of Science and Technology.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.