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Commun Biol
Commun Biol
Communications Biology
2399-3642
Nature Publishing Group UK London

39223249
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10.1038/s42003-024-06768-4
Article
Activated tissue-resident macrophages contribute to hair cell insults in noise-induced hearing loss in mice
http://orcid.org/0000-0002-8203-8082
Pan Jing 12
Wang Kaiye 12
Qu Jiaxi 12
Chen Dongxiu 12
http://orcid.org/0000-0002-1707-4482
Chen Anning 12
You Yunyou 12
http://orcid.org/0000-0001-7882-4462
Tang Jie jietang@smu.edu.cn

12345
http://orcid.org/0000-0001-8105-2256
Zhang Hongzheng redtrue@smu.edu.cn

12
1 grid.284723.8 0000 0000 8877 7471 Department of Otolaryngology Head & Neck Surgery, Zhujiang Hospital, Southern Medical University, Guangzhou, 510282 China
2 grid.284723.8 0000 0000 8877 7471 Ear Research Institute, Zhujiang Hospital, Southern Medical University, Guangzhou, 510282 China
3 https://ror.org/01vjw4z39 grid.284723.8 0000 0000 8877 7471 Department of Physiology, School of Basic Medical Sciences, Southern Medical University, Guangzhou, 510515 China
4 https://ror.org/01vjw4z39 grid.284723.8 0000 0000 8877 7471 Key Laboratory of Mental Health of the Ministry of Education, Southern Medical University, Guangzhou, 510515 China
5 https://ror.org/01vjw4z39 grid.284723.8 0000 0000 8877 7471 Guangdong-Hong Kong Joint Laboratory for Psychiatric Disorders, Southern Medical University, Guangzhou, 510515 China
2 9 2024
2 9 2024
2024
7 107810 1 2024
21 8 2024
© The Author(s) 2024
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.
Macrophages serve as the primary immune cell population and assume a pivotal role in the immune response within the damaged cochleae. Yet, the origin and role of macrophages in response to noise exposure remain controversial. Here, we take advantage of Ccr2RFP/+ Cx3cr1GFP/+ dual-reporter mice to identify the infiltrated and tissue-resident macrophages. After noise exposure, we reveal that activated resident macrophages change in morphology, increase in abundance, and migrate to the region of hair cells, leading to the loss of outer hair cells and the damage of ribbon synapses. Meanwhile, peripheral monocytes are not implicated in the noise-induced hair cell insults. These noise-induced activities of macrophages are abolished by inhibiting TLR4 signaling, resulting in alleviated insults of hair cells and partial recovery of hearing. Our findings indicate cochlear resident macrophages are pro-inflammatory and detrimental players in acoustic trauma and introduce a potential therapeutic target in noise-induced hearing loss.

Cochlear tissue-resident macrophages, not infiltrated peripheral monocytes, respond to noise exposure in mice. The pro-inflammatory effects of these macrophages contribute to insults of ribbon synapses of hair cell in noise-induced hearing loss

Subject terms

Diseases
Neurology
Auditory system
"Technology Innovation 2030-Major Projects" on brain science and brain-like computing of the Ministry of Science and Technology of China(2021ZD0202603)issue-copyright-statement© Springer Nature Limited 2024
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pmcIntroduction

The cochlea was long considered immune privileged due to the presence of blood-labyrinth barriers (BLB). However, the activities of Immune cells and inflammation have been described in acute and chronic insults to the cochlea, such as noise, ototoxic injury, aging, or a cochlear implant insertion1–8. In response to these insults, macrophages are the primary immune cell population in the cochlea and play a major role in immune activity9. High-level noise exposure (NE) induces the damage of cochlear sensory cells, resulting in permanent hearing loss. Numerous of researches have shown dramatic alteration in the abundance, morphology, and mediator expression of cochlear macrophages after NE, indicating the activity of macrophages in response to acoustic trauma. However, before we conclude that cochlear macrophages would be a therapeutic target in noise-induced hearing loss (NIHL), pivotal questions need to be clarified. What is the origin of activated macrophages in the cochlea, i.e., are they cochlear tissue-resident or infiltrated macrophages? Whether the activated cochlear macrophages are injurious or protective to cochlear function?

Tissue-resident macrophages, with specific expression signatures of CX3CR1, CD163, and CD11b, distribute across modiolus, spiral ganglion neurons (SGNs), and spiral ligament of the cochlea. Under physiological conditions, they contribute to the maintenance of homeostasis and maintain their population with a turnover time of several months10. Infiltrated macrophages, with specific expression signatures of CCR2 and Ly6C, are recruited from the bone marrow monocytes. They infiltrate into tissues and differentiate into macrophages almost instantly following acute insults. Generally, both activated resident and infiltrated macrophages contribute to the immune response to insults by producing pro-inflammatory and anti-inflammatory mediators, phagocytosis, and antigen presentation. In NIHL, the evidence regarding the origin of activated macrophages in the cochlea is conflicting. It is unclear which macrophage population is activated and initiates the response to NE.

The role of activated macrophages in NIHL also remains to be determined. Evidenced by the increasing pro-inflammatory cytokines and chemokines in the cochlea following acoustic trauma, macrophages are proposed to initiate the inflammation and may result in the subsequent loss of cochlear sensory cells. Meanwhile, macrophages are involved in reparative response by producing anti-inflammatory mediators, clearing cellular debris and pathogens from tissues. These reparative functions are evidenced by the activity of macrophages in tissue repair after NE and cochlear implantation1,5,7. Whether and when macrophages are protective or injurious is critical to understanding the role of macrophages in NIHL.

To address the above questions, in the present study, we employed the CX3CR1 and CCR2 fluorescence labeling mice to identify the tissue-resident and infiltrated macrophages in the cochlea. We determined the changes in the abundance, distribution, and morphology of these macrophages across different regions of the cochlea after NE. The roles of macrophage activity in NIHL and the relevance of the TLR4-NLRP3 pathway were also examined.

Results

Noise exposure induces permanent hearing loss and insults to hair cells

In the present study, WT, Cx3cr1GFP/+, and Ccr2RFP/+ Cx3cr1GFP/+ mice were exposed to white noise at 110 dB SPL for 3 h. Such high-level NE was reported to induce a permanent threshold shift in CBA/CaJ and C57BL/6 mice1,10–13. As shown in Fig. 1A, an average of 40 dB of the free-field auditory brainstem response (ABR) threshold shift in 16 kHz was observed at 1 day (D1) and 7 days (D7) post-NE for animals, suggesting severe hearing loss in both WT and reporter mice. The monaural ABR measurements in WT mice showed that the threshold lifting lasted for at least 14 days (D14) after exposure at all frequencies and clicks examined (Fig. 1B), indicating it should be a permanent hearing loss in our experimental design. More evidence comes from the morphological examination of cochlear sensory hair cells. Compared with the control group, the mice in D14 showed a significant loss of outer hair cells (OHCs) at apical, middle, and basal turns of the cochlea, whereas the inner hair cells (IHCs) showed no significant decrease (Fig. 1C, D). However, the severe hearing loss in our experiment may not be attributed to such mild loss in OHCs completely. Such level of hair cell loss typically results in frequency-dependent hearing loss, predominantly affecting high frequencies from the basal turns, and is expected not to surpass 40 dB conferred by the amplification role of outer hair cells. These data implied that factors beyond hair cell loss contributing to the observed severe hearing loss. Therefore, we consequently examined the ribbon synapses that transfer signals from IHCs to SGNs. As shown by the immunostaining of pre- (CtBP2) and post-synaptic (GluR2) puncta of ribbon synapses, the density of ribbon synapses was significantly decreased at all cochlear locations compared with the controls (Fig. 1E, F). These data indicate that insults of hair cells, especially the loss of ribbon synapses, are responsible for the NIHL in our experiment.Fig. 1 Noise exposure induces hearing loss and impairment in hair cells.

A Open-field ABR thresholds at 16 kHz in WT, Cx3cr1GFP/+, and Ccr2RFP/+Cx3cr1GFP/+ mice before, 1 day (D1), and 7 days (D7) post noise exposure (n = 4 for each group. **P < 0.01 and ***P < 0.001, two-way ANOVA). B Closed-field ABR thresholds in response to tone burst and click before, D1, D7, and 14 days (D14) post noise exposure in WT (n = 16 for each time point. ***P < 0.001, two-way ANOVA). C Representative immunofluorescent staining images show the hair cells in apical, middle, and basal turns before (control) and D14. Hair cells were labeled with myosin 7a (red). Scale bar = 20 μm. D The number of hair cells from apical, middle, and basal turns before and D14 (n > 5 from independent images for each group. *P < 0.05 and ***P < 0.001, Student’s t test). E Representative immunofluorescent staining images show the ribbon synapse before and D14. HCs were labeled with myosin 7a (blue), and the pre-and postsynaptic segment of ribbon synapse was labeled by CtBP2 (red) and GluR2 (green), respectively. Scale bar = 10 μm. F Statistical analysis of synaptic puncta density before and after noise exposure on D14 (n > 6 IHCs for each group. **P < 0.01 and ***P < 0.001, Student’s t test). The data were presented as mean ± SEM.

No infiltration of peripheral monocytes into the cochlea after noise exposure

To distinguish the origin of cochlear macrophage populations after NE, we used the Ccr2 RFP/+Cx3cr1GFP/+ dual-reporter mice in our experiment. In this mouse model, the CCR2-positive peripheral monocytes are labeled with red fluoresce (CCR2-RFP), while the CX3CR1-positive tissue-resident macrophages are labeled with green fluoresce (CX3CR1-GFP). This mouse model has been widely used for tracking monocytes recruitment in the inflammation response in brain injury9,14,15. Under control conditions, as shown by the cochlear cross-section, CX3CR1-GFP cells were found in multiple regions, including the Rosenthal’s canal (SGNs), basilar membrane (BM), spiral ligament (SL), and stria vascularis (SV). CCR2-RFP cells were only found sporadically among SGNs and SL (Fig. 2A). As shown by the representative images of a cochlea at D7, NE increased the number of CX3CR1-GFP cells. However, CCR2-RFP cells were still rarely observed after NE (Fig. 2A). The dynamics of the CX3CR1-GFP and CCR2-RFP cell abundance in SGNs, SV, and SL regions are shown in Fig. 2B. As CX3CR1-GFP cells were increased in all regions, no significant increase of CCR2-RFP cells was detected at different times after NE. To confirm the reliability of CCR2-RFP labeling, a silicone strip, the primary component of cochlear implants, was inserted into the cochlea of a Ccr2 RFP/+Cx3cr1GFP/+ dual-reporter mouse, mimicking the procedure of cochlear implantation. One day post-implantation, a significant presence of CCR2-RFP cells was observed throughout the cochlea, along with an increase in CX3CR1-GFP cells (Supplementary Fig. 1). This CCR2-RFP positive result of invasive injury was different from the CX3CR1-dominant responses after NE.Fig. 2 The dynamics of macrophage populations in the cochleae post noise exposure.

A Confocal images show the distribution of CX3CR1-GFP (green) and CCR2-RFP (red) cells in cochlea cross-section of Ccr2RFP/+Cx3cr1GFP/+ mice before and D7 after noise exposure. The hair cells were labeled with myosin 7a (gray) and the nuclei were labeled with DAPI (blue). SGNs: spiral ganglion neurons; BM: basilar membrane; SV: stria vascularis; SL: stria ligament. B Quantitative analysis of CX3CR1-GFP and CCR2-RFP cells across the different regions of cochlea before (control), D1, D3, and D7 after noise exposure. (n > 4 cochleae for each group. ***P < 0.001, two-way ANOVA). C Mean values of endolymph potential (EP) recorded from the basal cochlear turn in WT mice before, immediately (D0), D1, and D7 after noise exposure. (n > 3 mice for each group. ***P < 0.001, one-way ANOVA). D Fluorescence shows the distribution of CX3CR1-GFP (green) and CCR2-RFP (red) cells in the basilar membrane of Ccr2RFP/+Cx3cr1GFP/+mice before and after noise exposure on D1. The hair cells were labeled with myosin 7a (blue). Scale bars = 100 μm. Error bars represent the SEM in (B) and the SD in (C).

To examine the integrity of the intrastrial fluid–blood barrier, the endolymphatic potential (EP) was measured at different times after NE. A ~40% reduction in EP value was recorded immediately after NE, and then fully recovered to control value within 24 h (Fig. 2C). This drop in EP indicates the temporary breakdown of the intrastrial fluid–blood barrier, which is essential for hearing function and cochlear immune homeostasis. The loss of the integrity of the intrastrial fluid–blood barrier may result in the leakage of molecules and cells from the capillaries. Interestingly, our data showed that no peripheral monocytes were allowed to infiltrate into the cochlear at any time after NE, even during this period.

The macrophages were further examined in the whole mounts of the BM, where pathologies were found in hair cells after NE. As shown by the representative images in Fig. 2D, compared with the control condition, abundant CX3CR1-GFP cells were observed after NE, especially in the region of hair cells where CX3CR1-GFP cells were rarely present in a steady state. Consistent with other regions, little CCR2-RFP cells under control conditions and no increase of their abundance after NE were found in our experiments. These results confirmed that peripheral monocytes were not infiltrated into the cochlear, suggesting that infiltrated macrophages were not implicated in the response to noise trauma.

Resident macrophages are activated after noise exposure

A previous study showed that depletion of CCR2 may promote the loss of hair cells after NE in the CX3CR1GFP/+ mouse strain13. Although our experiments have shown no difference in the noise-induced ABR shift between the Ccr2 RFP/+Cx3cr1GFP/+ mice and the WT controls (Fig. 1A), the impact of CCR2 gene manipulation on the CX3CR1 positive macrophages is uncertain. Therefore, instead, the activity of tissue-resident macrophages in the BM was examined in the CX3CR1GFP/+ mouse strain, in which the tissue-resident macrophages are labeled with green fluorescence of the CX3CR1 biomarker.

Under control conditions, CX3CR1-GFP cells were distributed near the modiolus side along the entire BM, while only a few CX3CR1-GFP cells were located in the region of hair cells (Fig. 3A). To better illustrate the dramatic changes in macrophage bodies and processes post-NE, we performed 3D reconstructions of confocal images. As shown by right panel of Fig. 3A, before NE, macrophages at the basilar membrane were stellate in shape with a slender body and multiple long dendritic processes. (cell 1 and 2). After NE, cell bodies became rounder, and processes significantly shortened (cell 3) or even disappeared (cell 4). Such change in morphology implies immune activation of CX3CR1-GFP macrophages after NE. To quantify the noise-induced changes in abundance and distribution, the numbers of CX3CR1-GFP cells in the different regions of the BM are shown in Fig. 3B. A dramatic increase of CX3CR1-GFP cells was observed in the region of hair cells (–100 ~ 0 μm and 0–100 μm from the IHCs) at all cochlear locations, while a significant increase was also found in most other regions of the BM.Fig. 3 Changes of CX3CR1-GFP macrophages in morphology, abundance, and distribution after noise exposure.

A Left panel: fluorescence shows the CX3CR1-GFP (green) cells in the basilar membrane of Cx3cr1GFP/+ mice before (control) and after noise exposure on D7. The hair cells were labeled with myosin 7a (red) and the nuclei were labeled with DAPI (blue). Scale bars = 100 μm. Insets are the magnified images of representative macrophages in each box (Scale bars = 10 μm). Right panel: 3D reconstructions of representative macrophages in left panel. Scale bars = 10 μm. B The density of CX3CR1-GFP cells at different distances from the IHCs. The cell numbers were measured every 100 μm x 100 μm region. The left, middle, and right panels show the data from apical, middle, and basal turns of the cochlea, respectively. n > 3 cochleae for each group. The data were presented as mean ± SEM. **P < 0.01 and ***P < 0.001, Student’s t test.

Toll-like receptor 4 (TLR4) is a cellular receptor of inflammatory factors and endogenous molecules of damaged cellular debris. Upon the binding of these ligands, TLR4 activates macrophages and the innate immune system through the NF-κB signaling pathway16–18. In activated macrophages, the NLRP3 inflammasome pathway is usually associated with damaging inflammatory processes and apoptosis. Both signaling pathways have been documented to modulate the insults of cochlear sensory cells19,20. However, the interaction of these signaling pathways and their roles remained unknown in macrophages after NE. In agreement with previous studies21,22, we found that TLR4/NF-κB and NLRP3 inflammasome pathways were expressed in cochlear macrophages and activated by NE (Supplementary Fig. 2A). The up-regulation of NF-κB and expression of IL-1β suggests that cochlear macrophages were activated and may play a pro-inflammatory role after NE. To further confirm the effect of these pathways on the activation of cochlear macrophages, TAK-242, the TLR4 inhibitor was introduced into the cochlea through the trans-tympanic route. To verify the efficacy and timing of drug entry into the inner ear, as well as to ensure trans-tympanic injections did not significantly affect mouse hearing, we created a solution of FM1-43 using the same formula and administered it via trans-tympanic injection. The results showed fluorescence on hair cells within a day and no impact on hearing, demonstrating the efficacy and safety of this drug delivery method in our study (Supplementary Fig. 3). Under the administration of TAK-242, the expression levels of TLR4, MyD88, and NF-κB p65 were decreased significantly, indicating that the noise-up-regulated TLR4 pathway was inhibited. Accompanied, the activation of the NLRP3 pathway was also inhibited as reflected by a remarkable reduction of NLRP3, ASC, and IL-1β (Fig. 4A). Interestingly, the increase of CX3CR1-GFP cells and their redistribution in the BM were also abolished by TAK-242 (Fig. 4B, C). These results, together with the changes in the morphology, abundance, and distribution, indicated that cochlear resident macrophages were activated and switched to a pro-inflammatory phenotype by NE.Fig. 4 Trans-tympanic injection of TAK-242 inhibits noise-induced TLR4-NLRP3 activity and changes in CX3CR1-GFP macrophages.

A Representative immune bands (upper panels) and quantitative analysis (lower panels) of western blotting show the expression level of TLR4, MyD88, NF-κB, NLRP3, ASC, and IL-1β from control and D3 cochlea. TAK-242: trans-tympanic injection of TAK-242, an inhibitor of TLR4. NE: noise exposure. GAPDH and β-actin were applied as the loading control. (n > 5 mice for each group. *P < 0.05; **P < 0.01 and ***P < 0.001, one-way ANOVA). B Representative confocal images of the basilar membrane after noise exposure (NE). Macrophages were labeled by GFP (green). The hair cells were labeled with myosin 7a (red) and the nuclei were labeled with DAPI (blue). C The density of CX3CR1-GFP cells at different distances from the IHCs. The cell numbers were measured every 100 μm x100 μm region. The upper, middle, and bottom panels show the data from the apical, middle, and basal turns of the cochlea, respectively. (n > 3 cochleae for each group. *P < 0.05 and ***P < 0.001, Student’s t test). Scale bar: 100 μm. The data were presented as mean ± SEM.

TAK-242 attenuates ribbon synapse reduction and hearing loss in noise trauma

To determine the roles of activated macrophages in NIHL, TAK-242 was applied to inhibit the activation of cochlear macrophages via the TLR4-NLRP3 pathway. As shown in Fig. 5, the noise-induced loss of CtBP2 puncta and paired ribbon synapses were significantly reduced in the IHCs treated with TAK-242. Considering the insult of ribbon synapse was the major pathology of noise trauma for hair cells in our experiment, this result suggests that the inhibition of macrophage activation may have a protective effect in noise-exposed cochlea. This conclusion was further confirmed by the in vivo measurements of hearing function. Immediately after NE, one ear of the animal was trans-tympanic treated with TAK-242 while the other ear was treated with the vector (Fig. 6A). For the vector controls, monaural ABR thresholds were elevated significantly one day after NE (D1). Such NIHL was persistent, without any recovery observed at D7 and D14 (Fig. 6B). For those ears treated with TAK-242, although NE induced comparative acute hearing loss at D1, the ABR thresholds were recovered partially at most frequencies tested at D7 and D14 (Fig. 6B, C). The recovery of hearing function was accompanied by our morphological finding, indicating that activated macrophages promote the insults of hair cells and contribute to permanent hearing loss in noise trauma.Fig. 5 Inhibition of TLR4 prevents noise-induced ribbon synapse loss.

A Representative immunofluorescent staining images show the ribbon synapse after noise exposure (NE) with or without trans-tympanic injection of TAK-242. Hair cells were labeled with myosin 7a (blue), and the pre-and post-synaptic segment of ribbon synapse was labeled by CtBP2 (red) and GluR2 (green), respectively. Scale bar = 10 μm. B Quantitative analysis of synaptic puncta density for each group. Circles indicate the mean density of controls without noise exposure. (n > 12 inner hair cells for each group. ***P < 0.001, Student’s t test). The data were presented as mean ± SEM.

Fig. 6 Inhibition of TLR4 reduces noise-induced hearing loss.

A Experimental design. Immediately after noise exposure, TAK-242 was administrated via trans-tympanic injection into one ear, and vector was injected into the other ear of the same mouse as control. B Monaural ABR thresholds and (C) ABR threshold shifts in response to tone bursts and clicks before, D1, D7, and D14 post-noise exposure. Stars (*): comparison between before and NE+ vector group; hashes (#): comparison between NE + vector and NE + TAK242 group. (n > 6 mice for each group. *P < 0.05; **P < 0.01; and ***P < 0.001; #P < 0.05; ##P < 0.01; and ###P < 0.001, two-way ANOVA). The data were presented as mean ± SEM.

Discussion

Our experiments show that the abundance, distribution, and morphology of cochlear resident macrophages are dramatically changed by high-level NE. Together with these changes, the noise-induced macrophage activation was further confirmed by the up-regulated TLR4-NLRP3 pathway. Meanwhile, no conspicuous presence of infiltrated macrophages under control conditions or distinct changes in this population was observed after NE in the cochlea. Through the TLR4-NLRP3 pathway, the inhibition of resident macrophage activity alleviates the trauma of cochlear hair cells and the accompanying hearing loss after NE. Our finding in this study reveals that the activity of tissue-resident macrophages, but not infiltrated macrophages, contributes to the insults of hair cells in NIHL.

CCR2 and CX3CR1 dual-reporter mice have been widely used in studies of heart, retina, and brain ischemic injuries to distinguish subpopulations of macrophages, namely infiltrated and resident macrophages15,23,24. After NE, a significant increase in the number of CX3CR1-GFP cells was observed, whereas changes in CCR2-RFP cells were not pronounced (Fig. 2), indicating a CX3CR1-dominated immune response to NE. Previous studies have shown that CCR2 depletion exacerbates hair cell loss in CX3CR1GFP/+ mice after NE, suggesting a possible interaction between CCR2 and CX3CR1 cells13. Ccr2RFP is a knock-in/knock-out allele. In heterozygous Ccr2RFP/+ mice, which showed same hearing phenotype as wild-types in our study, one allele of Ccr2 was replaced by RFP. To avoid the potential impact of CCR2 deficiency, CX3CR1GFP/+ mice were employed instead in subsequent experiments to examine the resident macrophage activity. This allowed us to observe CX3CR1-GFP cell responses with unaffected CCR2 cells, providing more straightforward evidence of noise-activated resident macrophages. The widely used C57BL/6 J mouse strain is known to exhibit early age-related hearing loss (AHL), which is associated with a mutation in the Cdh23 gene of this mouse. This mouse strain begins to show degeneration of cochlear hair cells around the age of 6 months, leading to hearing impairment25. The mice used in our study, including CX3CR1-GFP mice and CCR2 and CX3CR1 dual-reporter mice, are all developed from the C57BL/6 J mouse strain. To mitigate the effects of early AHL, we utilized young mice aged 1–2 months for our experiments. During the experimental period, the mice did not exhibit AHL, nor did we observe age-related morphological changes in the cochlea. However, we did not examine the cochlear macrophages or expose older mice (older than 6 months) from the C57BL/6 J strain to noise. Consequently, our data did not ascertain whether the impact of noise on macrophages and the resulting hearing loss were consistent across other life stages.

Macrophages are found present in the cochlea of humans and mice9,26–28. Derived from the yolk sac and the fetal liver, these macrophages are long persisting in the cochlea from the embryonic stage29,30. Consistent with existing reports9, our results reveal that macrophages reside in the Rosenthal’s canal (SGN), organ of Corti, and lateral wall of the adult cochlea (Figs. 2, 3). Cochlear supporting cells are traditionally known for their vital role in maintaining the structure and ionic balance of the sensory epithelium. Interestingly, recent studies have shown that these cells exhibit macrophage-like features, including phagocytosis, cytokine secretion, and participation in cochlear immune responses31–33. In the stria vascularis (SV), processes of perivascular-resident macrophage-like melanocytes (PVM/Ms) contact with endothelial cells and ensheath strial capillaries, which is essential for forming of BLB and maintaining the EP34. The integrity of the intrastrial fluid–blood barrier prevents the leakage of molecules and cells from the blood vessels, implying that tissue-resident macrophages are the major population in healthy cochlea. This opinion is supported by our result that most cochlear macrophages are CX3CR1 positive before NE (Fig. 2). Under steady state, resident macrophages contribute to the maintenance of immune homeostasis with a relatively slow renewal through the infiltration of bone marrow-derived monocytes to the cochlea3,35,36. This may explain the origin of sporadic CCR2-positive macrophages observed in the cochlea under control conditions (Fig. 2).

NE was widely reported to cause temporary (reversible) or permanent threshold shifts of hearing, depending on the intensity and duration of NE used in different studies. In the present study, high-level NE was designed to induce acute and irreversible hearing loss, evidenced by the loss of hair cells and the trauma of ribbon synapse (Fig. 1). Our data show that noise increased the number of macrophages in the regions of SGN, basilar membrane, SV, and SL (Fig. 2). The change in macrophage number is accompanied by its re-distribution in the basilar membrane, particularly in the region adjacent to sensory hair cells (Fig. 3). Interestingly, the changes in the distribution and number of cochlear macrophages were carried out by CX3CR1 labeled cells, indicating that cochlear resident macrophages were the major population in response to noise trauma. One possible approach to of circulation macrophages infiltrated in the cochlea is through the insulted intrastrial fluid–blood barrier after NE. Activated by NE, PVM/Ms detached from capillary walls, resulting in the increased permeability of the blood-labyrinth barrier and the drop in EP37. Our experiments show that EP decreased immediately and recovered fully one day after NE, suggesting that the loss of integrity of the blood-labyrinth barrier is transient. This temporary increase in permeability allowed the penetration of immune-active elements into the cochlear, resulting in the upregulation seen at day 1 in Fig. 2B. Nonetheless, CCR2-RFP cells were not allowed to infiltrate into the inner ear, even the permeability of BLB was temporary increased by NE.

Besides the changes in distribution and abundance, the change of macrophage morphology after NE provides more evidence for the activation of cochlear macrophages. As previously reported, our data show that under homeostatic conditions, cochlear macrophages have branched morphology with ramified long processes9,28. However, after NE, macrophages acquired a small and rounded ameboid shape with less ramified extensions (Fig. 3). As an indicator of activated immune state, this change in morphology indicates the activation of macrophages after NE5,27,38,39. This conclusion is also supported by the up-regulation of TLR4 signaling, which has been reported to modulate the cochlear immune response after NE19.

TLR4 is known to be expressed in cochlear macrophages and has been implicated in the pathogenesis of acoustic trauma19. Interacting with its ligands leaked from the temporally break-down BLB, or endogenous molecules of damaged cellular debris, TLR4 can activate cochlear macrophages through the NF-κB signaling pathway16–18. Although the TLR4 signaling was confirmed to be associated with the activation of macrophages in our experiment, the role of activated cochlear macrophages in acoustic injury still needs to be determined. By producing inflammatory molecules and recruiting more immune cells, TLR4 signaling-activated macrophages have been documented to be implicated in the loss of hearing and hair cells after acoustic injury19. In a lower-level NE mouse model, however, the hearing was reported to fully recover without any hair cell loss, even though cochlear macrophages were activated comparable39. Meanwhile, activated macrophages are also linked to the protective and reparative processes after strong acoustic injury40. Thus, clarifying the role of macrophages in cochlear damage is crucial for understanding the mechanisms behind hearing loss. Regarding cochlear damage and hearing loss due to high-level NE, the conclusions from our present study were contrary to those of Manickam et al., who believe macrophages protecting cochlea against NIHL40. They employed an indirect approach, using the macrophage depletion agent PLX5622, and found that damage induced by NE was exacerbated after the removal of cochlear macrophages. However, macrophages play a vital role in maintaining immune homeostasis within the cochlea. Removing these cells would disrupts this balance, leading to cochlea degeneration and hearing loss34. Additionally, the depletion agent likely affects PVM/Ms, disrupting the integrity of the intrastrial fluid–blood barrier, thus increasing susceptibility to hearing loss and NE. Consequently, the more severe decline in hearing following PLX5622 treatment was a combined effect of the loss of macrophages homeostatic functions and acoustic trauma. In the absence of control data and functional experiment, it was reckless to conclude that macrophages play a protective role in noise-induced damage. Our research adopted a more direct approach, mitigating hair cell damage and hearing loss from NE by pharmacologically inhibiting macrophage pro-inflammatory activity, leading us to conclude that macrophages activity indeed aggravates acoustic trauma.

To determine the role of macrophage activity in NIHL, TLR4 inhibitor was employed to apply to the inner ear. Two delivery approaches were considered: local delivery to the cochlea or systemic administration (such as intravenous injection). Due to the presence of the BLB, systemic administration requires higher doses to be effective in the cochlea. Given the widespread distribution of the TLR4 signaling pathway, high doses of systemic administration could trigger systemic reactions across multiple organs, especially the immune system. Such systemic effects could be complex and difficult to control. In our experimental design, we administered the TLR4 inhibitor via trans-tympanic injection to one ear and the vector to the contralateral ear of the same animal. The ear that received the TLR4 inhibitor underwent auditory measurements and histological analysis to assess the role of the TLR4 pathway in NIHL, with the vector-injected ear serving as the control. This method of local delivery was employed to highlight the effects of TLR4 inhibition on cochlea resident macrophages while avoiding systemic effects caused by intravenous injection. Although there was a possibility that the drug could affect the control ear through systemic circulation, the NIHL and ribbon synapse loss of control ear were identical to those of untreated ear. The morphological and functional changes in control ears (and untreated ears) were significantly greater than the ears with local injection (Figs. 5 and 6). Our findings suggested that local TLR4 inhibition only affected the injected cochlea and did not impact the contralateral ear, even with potential low-dose leakage through the circulation.

Our experiments show the up-regulation of NLRP3 signaling, which mediates the pro-inflammatory responses in activated macrophages41,42, is associated with the noise-induced TLR4 activity (Fig. 4, Supplementary Fig. 2). Blocking of the TLR4 activity inhibited the activation and re-distribution of cochlear resident macrophages, resulting in less trauma in hair cells and a partial reversal of hearing loss (Figs. 5, 6). Our results confirm that the activity of cochlear macrophages plays a pro-inflammatory role and aggravates the insults of hair cells in acoustic injury. It has been demonstrated that NLRP3 gene mutations in human can lead to sensorineural hearing loss, which was notably ameliorated, and in some cases, entirely reversed by the administration of Anakinra, an IL-1β inhibitor43. This significant observation underscored the critical role of cochlea immune responses, mediated by NLRP3, in influencing auditory function. Such findings aligned closely with the results of our study, which a marked upregulation of the NLRP3 inflammasome pathway in the cochlea after NE in mice. However, our data indicated that inhibition of TLR4, and consequently the partial suppression of the NLRP3 inflammasome, only yielded a partial recovery of hearing after NE. The partial recovery of hearing loss despite TLR4 inhibition hints at the existence of additional, NLRP3-independent pathways that contributed to the noise-induced cochlear damage and hearing loss. Nonetheless, the incomplete recovery of hearing loss upon TLR4 inhibition underscored the complex interplay of multiple signaling pathways in the pathogenesis of NIHL, suggesting that a multifaceted approach targeting both NLRP3-dependent and independent pathways may be necessary for the comprehensive treatment of noise-induced cochlear damage.

Beyond the inflammation-induced cell damage, another possible mechanism of the activated macrophages aggravating cell damage is the phagocytic function. For example, serving as macrophage in the central neural system, microglia in the brain have been found to phagocytose cell debris and redundant synapses in Alzheimer’s disease44,45. Activated cochlear macrophages may play a similar role in noise-induced ribbon synapse loss. The effects of NE on the cochlear macrophage and their impacts on tissue pathogenesis are summarized in Fig. 7.Fig. 7 Overview of the activity of cochlear macrophages in noise-induced hearing loss.

Under steady state, the resident macrophages reside in the Rosenthal’s canal (SGNs), SL, and basilar membrane of the adult cochlea. Perivascular macrophage-like melanocytes (PVM/Ms) are adjacent to blood vessels in the SV, maintaining the integrity of the intrastrial fluid–blood barrier. Activate PVM/MS with shorter processes are detached from capillary walls after noise exposure, resulting in temporary breakdown of the blood-labyrinth barrier. Together with the debris of cochlear cells, leaked inflammatory factors activate resident macrophages via the TLR4-NLRP3 pathway. After noise exposure, activated resident macrophages change in morphology, increase in abundance, and migrate to the region of hair cells, leading to the loss of OHCs and the damage of ribbon synapses. Meanwhile, peripheral monocytes are not implicated in the noise-induced hair cell insults. SGNs: spiral ganglion neurons; SV: stria vascularis; SL: spiral ligament; OHCs: outer hair cells; IHCs: inner hair cells.

The portion of hearing loss cannot be eliminated by the inhibition of macrophage activity revealing that other mechanisms are involved in the noise-induced trauma. For example, the violent mechanical force of vibration peels hair bundles off from the hair cells46. This cellular debris would initiate the activation of cochlear macrophage via the TLR4- NLRP3 pathway. Meanwhile, without the implication of macrophage activity, loss of hair bundles leads to the dysfunction of hair cells immediately and apoptosis of insulted cells in two weeks47. Other effectors besides cochlear immune responses, including ROS-mediated cochlear cell apoptosis and autophagy, are well-documented to contribute the acute insults to the cochlea after NE48–50. In conclusion, our results indicate that cochlear resident macrophages are activated and contribute to the insults of hair cells in NIHL.

Methods

Subjects

All animal experiments conducted in this study were approved by the Institutional Animal Care and Use Committee (IACUC) of Southern Medical University.

C57BL/6 wild-type (WT) female mice aged 1.5–2 months and weighting 15–20 g were used in our experiments. B6.129(Cg)-Cx3cr1tm1Litt Ccr2tm2.1Ifc/JernJ mice (Stock number: 032127) and B6.129P2(Cg)Cx3Cr1tm1Litt/J mice (Stock number: 005582) (The Jackson Laboratory, Bar Harbor, ME, USA) at the same age were also used in our study. B6.129(Cg)-Cx3cr1tm1Litt Ccr2tm2.1Ifc/JernJ mice, also known as Ccr2RFPCx3cr1GFP mice, express red fluorescent protein in CCR2-positive peripheral monocytes, T cells, and NK cells; and green fluorescent protein in monocytes, dendritic cells, NK cells, and brain microglia. B6.129P2(Cg)-Cx3Cr1tm1Litt/J mice, also known as Cx3cr1GFP mice, express green fluorescent protein in the place of Cx3cr1. Heterozygous mice (Ccr2RFP/+Cx3cr1GFP/+ and Cx3cr1GFP/+) were created by cross-breeding corresponding homozygous with C57BL/6 J mice. For genotyping, using One Step Mouse Genotyping Kit (Vazyme, Nanjing, China), genomic DNA was extracted from a small piece of mice tail and was used as template for PCR with specific primers. The primers used in the genotyping protocols are listed in Supplementary Table 1. Mice were housed in a room maintained at 22 ( ± 2) °C and subjected to a 12:12 light/dark cycle with lights on at 8:00 AM.

Auditory brainstem response (ABR) measurements

ABR measurements were conducted to assess the auditory function of the mice, following established protocols51. Animals were anesthetized with an intraperitoneal (i.p.) injection of sodium pentobarbital (80 mg/kg) and were maintained on 0.5% inhaled isoflurane, and placed on the center of an anti-vibration table in a soundproof room. A subdermal needle electrode (recording electrode) was located over the skull vertex, and the reference and ground electrodes were placed ventrolaterally to the external pinna. For close-field ABR recording, each ear was individually stimulated using the tube of a close-field speaker (MF1, Tucker-Davis Technology, Alachua, FL, USA). For open-field ABR recording, the speaker was positioned 10 cm away toward the head of the animal. Software SigGenRZ (Tucker-Davis Technology, Alachua, FL, USA) was used to generate click and 5 ms tone bursts with various frequencies and intensities. Click and tone burst stimulus intensities were gradually decreased by 5 dB from 95 dB sound pressure level (SPL). The ABR waveforms were amplified, recorded, averaged, and stored by the software BioSigRZ. Real-time averages of the amplified evoked responses from 256 sweeps were made. The ABR thresholds were defined as the minimum sound intensity at which averaged waveforms could be distinguished. An SPL threshold of 95 dB was assigned when there was no significant ABR wave at maximum strength.

Noise exposure

Animals were placed in a vertical acrylic cylinder (10 cm in diameter, 20 cm high) to receive noise exposure. Continuous Gaussian white noise at a level of 110 dB SPL for 3 h was generated by the software SigGenRZ. The signal was routed through a power supply (P350, Breeze Audio, China) and a power amplifier (TAS5630, Breeze Audio, China) to a loudspeaker positioned above the acrylic cylinder. The noise level was calibrated using a sound level meter (2610, Brüel & Kjær, Denmark).

Trans-tympanic administration

Animals were anesthetized with pentobarbital sodium (80 mg/kg, i.p.). A heating pad was used to maintain their body temperature during administration. Specific TLR4 inhibitor TAK-242 (0.2 mg/kg, HY-11109, MedChemExpress Company, New Jersey, USA, LogP calculated as 2.53 via ChemDraw) was dissolved in a mixture of a vehicle containing 10% DMSO, 5% Tween-80, 40% PEG300, and 45% saline. Utilizing a 33-G cannula, 5 µl of either TAK-242 or the vehicle alone was gently injected through the tympanic membrane under a surgical stereomicroscope. Animals were allowed to recover on the heating pad before being returned to the animal housing facility. To assess the effectiveness of trans-tympanic drug delivery, we used membrane dye FM1-43 (Invitrogen, USA), which shared the formulation with TAK-242, for trans-tympanic injection. Subsequently, dye uptake in hair cells was examined by confocal microscopy.

Endocochlear potential (EP) recording

EP recording followed a well-documented method52, detailed as follows. Briefly, after anesthesia, a tracheotomy was performed to exposure the cochlea under a surgical stereomicroscope. Using a fine dental drill, a small hole was opened on the lateral wall of the cochlear basal turn. Controlled by a micromanipulator, a recording microelectrode (pulled by a glass pipet and filled with 150 mM KCl) was inserted into the scala media, while the ground electrode was placed on the dorsal neck muscles. The response from the microelectrode was amplified using the Multiclamp700B amplifier (Molecular Devices, Foster City, CA, USA) and acquired using the software pClamp 10 (Molecular Devices, Foster City, CA, USA). The voltage changes during the microelectrode penetration were continuously recorded. Bilateral ears were analyzed for each animal.

Histology and immunostaining

Histology and immunostaining procedures were conducted following the methodology outlined in our earlier research51,53. In brief, the mice were deeply anesthetized and killed by decapitation. The cochleae were dissected from the temporal bone, and perfused with 4% paraformaldehyde overnight at 4 °C. After 5 days of decalcification, the organs of Corti were isolated and divided into three segments evenly (apical, middle, and basal) for subsequent immunostaining. For quantifying CX3CR1 and CCR2 positive cells, the cochlea was serially sectioned in the axial plane at a thickness of 10 μm. Cochlea tissues were permeabilized by 0.3% Triton X-100 in PBS for 30 min followed by blocking with 10% normal goat serum for 1 h at room temperature. Tissues were incubated overnight at 4°C with primary antibodies as followed: rabbit anti-myosin 7a (1:1,000, 25-6790, Protein Tech, Chicago, IL, United States), rat anti-F4/80 (1:1,000, MF48000, Invitrogen, Carlsbad, CA, United States), rabbit anti-TLR4 (1:1,000, ab13556, Abcam, United States), rabbit anti- F4/80 (1:1,000, 30325, Cell Signaling Technology, United States), rat anti-NLRP3 (1:1,000, Invitrogen, Carlsbad, CA, United States), mouse anti-CtBP2 (1:1,000, 612044, BD Biosciences, United States), mouse anti-GluR2 (1:1,000, MAB397, Millipore, United States). After incubation with the primary antibodies, the tissues were rinsed with PBS, and incubated in the dark with secondary antibodies for 1 h at room temperature. The secondary antibodies (Invitrogen, Carlsbad, CA, United States) were as follows: Myosin 7a staining: Alexa Fluor 568 goat anti-rabbit IgG, Alexa Fluor 647 goat anti-rabbit IgG or Alexa Fluor 405 goat anti-rabbit IgG; F4/80 and TLR4 staining: Alexa Fluor 488 goat anti-rat IgG and Alexa Fluor 405 donkey anti-rabbit IgG; F4/80 and NLRP3 staining: Alexa Fluor 488 goat antirabbit IgG and Alexa Fluor 405 goat anti-rat IgG; Myosin 7a, CtBP2 and GluR2 staining: Alexa Fluor 405 donkey anti-rabbit IgG, Alexa Fluor 488 goat anti-mouse IgG2a and Alexa Fluor 568 goat anti-mouse IgG1. The fluorescence images were acquired using a confocal laser scanning microscopy (Olympus, Japan).

CX3CR1-GFP cells analysis

When performing CX3CR1-GFP cells density analysis, the experimenter was blind to the treatment of animals. We performed calculations on whole-mount basilar membrane confocal image stacks captured under a 10X objective lens. Using inner hair cells as the origin (0), with the direction toward the cochlear modiolus as positive (+) and toward the outer hair cells as negative (−), we quantified the number and distribution of CX3CR1-GFP cells at various distances and directions from the inner hair cells within each 100 μm x100 μm region.

3D reconstruction

The Imaris volume rendering function (Imaris 8.1, Oxford Instruments; RRID:SCR_007370) was used for 3D reconstruction. Representative macrophages confocal image stacks were first converted to an imaris file (.ims) using Imaris FileConverter. For the 3D reconstruction of macrophages, the “surface” function was applied using the following custom settings: surface detail was defined as 0.2 mm (smooth); threshold (local contrast) as 200. After removal of all nonspecific background signal, the “mask all” function was used to create the final surface reconstruction.

Immunoblot analysis

Whole cochleae were homogenized with a glass pestle in ice-cold RIPA lysis buffer (Byotime Biotechnology, China) containing 1% PMSF (Fdbio science, China). Following a 30-min incubation on ice, the lysate underwent centrifugation at 12,000 × g at 4 °C for 15 min. Protein concentration was determined using the Bio-Rad Protein Assay BCA kit. Subsequently, 30 μg total protein from each sample underwent separation via SDS-PAGE and transfer to a PVDF membrane (Millipore, Bedford, MA, USA). The blots were blocked with 5% non-fat dry milk in TBST and subjected to an overnight incubation at 4 °C with primary antibodies targeting TLR4 (ab13556, Abcam, USA), NLRP3 (ab4207, Abcam, USA), NF-κB (8242, Cell Signaling Technology, USA), MyD88 (4283, Cell Signaling Technology, USA), ASC (67824, Cell Signaling Technology, USA), IL-1β (ab9722, Abcam, USA) and GAPHD (AG019, Byotime Biotechnology, China) diluted in primary antibody dilution buffer (Byotime Biotechnology, China). Following a PBS wash, the PVDF membrane underwent incubation with secondary antibodies conjugated to HRP (SA00001-1, SA00001-2, Proteintech, China) for 2 h. The chemiluminescence detection system (Image 600, GE Healthcare, Chicago, IL, USA) was employed to detect protein bands. Subsequently, protein grayscale analysis was conducted using the Image Quant TL software (GE Healthcare, Chicago, IL, USA).

Statistics and reproducibility

Statistical analysis was conducted using GraphPad Prism 8.4.2 (GraphPad Software). The data are presented as the mean ± SEM or SD, derived from a minimum of three independent experiments. Student’s t test was utilized for comparing two groups, while analysis of variance (ANOVA) was employed for comparisons among multiple groups. Post hoc analysis using Tukey’s test was performed after ANOVA. Experiments with two factors underwent analysis with a two-way ANOVA. A significance threshold of P < 0.05 was applied for all tests.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Supplementary information

Peer Review File

Supplementary Information

Description of Additional Supplementary Files

Supplementary Data 1

Reporting summary

Supplementary information

The online version contains supplementary material available at 10.1038/s42003-024-06768-4.

Acknowledgements

This study was supported by the “Technology Innovation 2030-Major Projects” on brain science and brain-like computing of the Ministry of Science and Technology of China (2021ZD0202603), the National Key Research and Development Program of China (2022YFC2402700), the National Natural Science Foundation of China (82271156), the Guangdong Basic and Applied Basic Research Foundation (2022A1515012036, 2023A1515012557), the Guangdong-Hong Kong Joint Laboratory for Psychiatric Disorders (2023B1212120004).

Author contributions

H.Z. and J.T. designed experiments and revised the manuscript. J.P. conducted the experiments, analyzed the data, and wrote the manuscript. K.W., J.Q., D.C., A.C., and Y.Y. conducted the experiments. All authors read and approved the final manuscript.

Peer review

Peer review information

Communications Biology thanks Yu Sun, Tracy A Newman, Si Ming Man and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Primary Handling Editors: Dr Ophelia Bu. [A peer review file is vailable].

Data availability

All data generated or analyzed during this study are included in this published article and its supplementary information files. The uncropped blot images in the figures can be found in Supplementary Figs. 4, 5 in Supplementary information. The source data behind the graphs in the paper can be found in Supplementary Data 1.

Competing interests

The authors declare no competing interests.

Ethical approval

All animal experiments conducted in this study are approved by the Institutional Animal Care and Use Committee (IACUC) of Southern Medical University. We have complied with all relevant ethical regulations for animal use.

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

These authors jointly supervised this work: Jie Tang, Hongzheng Zhang.
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