Chemokine Receptor CCR2 Is Protective toward Outer Hair Cells in Chronic Suppurative Otitis Media Open Access

Chronic suppurative otitis media (CSOM) is a significant global health burden, affecting up to an estimated 330 million people worldwide. It disproportionately impacts children in developing regions, where it is a leading cause of permanent hearing loss (1, 2). Although conductive hearing loss associated with CSOM is well documented, the impact on sensorineural hearing loss (SNHL) in the inner ear remains poorly understood. This limited understanding of the mechanisms driving SNHL development in CSOM poses a significant obstacle in the creation of effective treatments for millions of patients (3–5).

Previously, our laboratory developed and validated a CSOM animal model using a bioluminescent strain of Pseudomonas aeruginosa (6). Using this model, we were able to determine the precise timing of hearing loss and discovered macrophages associated with SNHL in CSOM (7). Furthermore, we found that certain inflammatory cytokines are elevated in CSOM. Especially, CCL2, an important member of the MCP family, is elevated over time following middle ear infections.

CCR2 is a common receptor of the MCP family and the unique receptor of CCL2. MCP binding to CCR2 has been the subject of prior research examining this interaction and the downstream signaling cascade that influences immunity, especially in monocyte chemotaxis in vivo (8, 9). Previous research indicated that both CCL2 and CCR2 contribute to macrophage migration and accumulation within the CNS following mouse hepatitis virus infection (10).

CCR2 is expressed on endothelial cells in the brain microvasculature, enhancing permeability of the blood–brain barrier (BBB) and increasing monocyte migration to the CNS in response to neuroinflammation (11–14). To date, only one study has looked at CCL2/CCR2’s role in the inner ear. Interestingly, this study showed CCR2 plays a neuroprotective role in the cochlea in noise-induced hair cell death, with a dramatic increase in outer hair cell (OHC) damage in CCR2^−/− relative to control but no differences in monocyte migration (15).

In the present study, we used the CCR2^−/− mouse model to study the immune response of the MCP-CCR2 signaling pathway in CSOM cochlea. We hypothesized that MCP-CCR2 signaling could protect OHC through regulation of macrophage migration in CSOM.

All animal procedures were conducted with the approval of the Stanford University Institutional Animal Ethics Committee. To study the physiological role of CCR2, we employed CCR2-deficient mice expressing an RFP reporter (CCR2^−/−RFP) [B6.129(Cg)- Ccr2^tm2.1Ifc/J, The Jackson Laboratory, stock no. 017586] (16). These mice, including males and females, were backcrossed onto a C57BL/6J background to generate study littermates. Mice aged 6–8 wk were either purchased from or bred from the original stock acquired from The Jackson Laboratory (Bar Harbor, ME). Animals were housed in the Stanford University animal care facility with ad libitum access to food and water. All procedures were performed with the animals under anesthesia induced by ketamine (80–100 mg/kg) and xylazine (8–10 mg/kg). Euthanasia was achieved by CO₂ inhalation followed by secondary cervical dislocation.

Our approach was based on the validated model of CSOM described by Xia et al. (7). Briefly, mice were anesthetized and positioned on the surgical stage under the microscope. A subtotal tympanic membrane perforation of the left ear was performed, followed by inoculation of 5 µl of persister cells into the middle ear cavity at a concentration of 1.63 × 10⁴ CFU/ml. These inoculations were consistently administered between 9:00 and 10:00 a.m. in all experiments. Following inoculation, mice were maintained in a prone position for 30 min and returned to the animal facility.

The cochleae were dissected and fixed in 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA) at 4°C overnight at 10 d following middle ear infection. Samples were then decalcified in 0.5 M EDTA (VWR, Radnor, PA) for 48 h at 4°C and washed three times in PBS (Fisher Scientific). For whole-mount preparations, the organs of Corti were dissected out from the cochleae under a stereo microscope. The cochlear epithelium was divided into three parts: apex (70–100% from the base), middle (30–70% from the base), and base (0–30% from the base). For cryosection preparation, cochleae were immersed in a sucrose gradient (10–30%) and embedded in OCT. Samples were collected in 10-μm sections.

Whole-mount tissues or cryosections were blocked with 5% donkey serum, 0.1% Triton X-100, 1% BSA, and 0.02% sodium azide (NaN₃) in PBS at pH 7.4 for 1 h at room temperature. Samples were then incubated in primary Abs overnight at 4°C. The following primary Abs were employed: rabbit anti-myosin VIIa (1:200; 25-6790; Proteus BioSciences) for hair cell labeling and rat anti-F4/80 (1:150, ab6640, Abcam Inc., Cambridge, MA) for macrophage labeling (17, 18). The specimens were incubated with secondary Abs diluted in 0.1% Triton X-100, 0.1% BSA, and 0.02% NaN₃ in PBS for 1 h at room temperature. The secondary Abs were conjugated with Alexa Fluor 546 (1:500; A10040, Life Technologies, Carlsbad, CA). After washing with PBS, specimens were mounted in ProLong Gold Antifade Reagent with DAPI (8961, Cell Signaling Technology, Danvers, MA) and placed under a coverslip. Images were captured using an LSM700 confocal microscope (Zeiss, Oberkochen, Germany) at 10× magnification.

Cells were analyzed using ImageJ software (NIH, Bethesda, MD). Hair cells (myosin VIIa positively labeled cells) were counted in whole-mount samples of the cochlear basal turn. In each sample, the surviving outer and inner hair cells were assessed along the basilar membrane. Macrophages (F4/80 positively labeled cells) were counted in cryosections. Each cochlear turn (base, middle, apex) was imaged and counted separately. Macrophages were counted in the regions of the spiral ganglion, spiral ligament, stria vascularis, and spiral limbus of each cochlea in all three cochlear turns, as described in our previous publication (7). The number of macrophages, summed over all regions and turns per cochlea, was compared between groups.

The cochleae from both the CCR2^+/+ and CCR2^−/− CSOM groups were dissected out at 10 d following middle ear infection. Samples were homogenized mechanically in lysis buffer. This buffer included 1% Triton X-100 (9002-93, Sigma-Aldrich, St. Louis, MO), 0.5% NP-40 (FNN0021, Thermo Fisher Scientific, Waltham, MA), 25 mM Tris HCl, pH 7.5 (1185-53-1, MilliporeSigma, Burlington, MA), 100 mM NaCl (7647-14-5, Sigma-Aldrich), Halt protease inhibitor mixture (78430, Thermo Fisher Scientific), and phenylmethanesulfonylfluoride (329-98-6, MilliporeSigma). Samples were stored at −80°C, and cytokine analysis was performed at the Human Immune Monitoring Core (Stanford University) as previously described (16). Briefly, Mouse 48-plex Procarta kits (EPX480-20834-901, Thermo Fisher Scientific) were employed, and plates were read using the FM3D FlexMap instrument with a lower bound of 50 beads per sample per cytokine. Custom Assay Chex control beads were added to all samples. The median fluorescence intensity (MFI) was averaged over duplicate wells for each cytokine per sample on each plate. The MFI for sample media, serving as the background, was averaged and then adjusted by adding 2 SD. The presented data represent the sample MFI minus the media MFI.

Dextran has been used to evaluate the permeability of the BBB under various conditions, including neurologic diseases, trauma, and drug treatments (19). It has also been used to assess blood–labyrinth barrier (BLB) permeability (20). To check BLB permeability in CCR2^−/− CSOM, FITC-dextran (MilliporeSigma, 46944-100MG-F) was injected i.v. Each mouse was administrated 200 μl at the concentration of 5 µg/µl. The cochleae were dissected and processed for cryosections 1 h after dextran-FITC administration. The sections were washed with PBS and a coverslip was placed after adding the mounting media. Images were captured using an LSM700 confocal microscope (Zeiss) at 10× magnification.

Auditory brainstem responses (ABRs) and distortion product otoacoustic emissions (DPOAEs) were serially measured in a cohort of mice as previously described (21, 22). Briefly, the ABR potentials were measured from needle electrodes positioned at the bottom of the tympanic bulla and at the vertex of the head, with a ground electrode placed in the rear leg. A bioamplifier (DP-311, Warner Instruments, Hamden, CT) was used to amplify the signal 10,000 times. The sound intensity level was raised in 10-dB steps from 10- to 80-dB sound pressure level (SPL), and the sound frequency was varied between 4 and 46 kHz. At each SPL, 260 responses were sampled and averaged. The peak-to-peak value of the ABR was measured, and the threshold at each frequency was calculated to be when this value was 5 SD above the noise floor. If no ABR response was detected, even at our equipment limits of 80-dB SPL, we arbitrarily defined the threshold to be 80-dB SPL. DPOAEs were measured by a probe tip microphone (type 4182, Brüel & Kjær, Nærum, Denmark) in the external auditory canal. The frequency response of this microphone was measured using a free field microphone with a flat frequency response out to 100 kHz (type 4939, Brüel & Kjær). This calibration curve was then used to adjust the DPOAE amplitudes we measured during the experiments. The sound stimuli for eliciting DPOAEs were two 1-s sine-wave tones of differing frequencies (F2 = 1.226F1). We varied the range of F2 from 4 to 46 kHz. The two tones were of equal intensities and stepped from 20- to 80-dB SPL in 10-dB increments. The amplitude of the cubic distortion product was measured at 2*F1–F2. The threshold at each frequency was calculated to be when the DPOAE was 0.5-dB SPL and 2 SD above the noise floor. If no DPOAE response was detected, even at our equipment limits of 80-dB SPL, we arbitrarily defined the threshold to be 80-dB SPL.

Statistical analyses were performed using GraphPad Prism version 9.5.1 (GraphPad Software Inc., La Jolla, CA). Two-tailed t tests were used to determine p values, with data considered statistically significant at p < 0.05. All data in figures are presented as the mean ± SD.

We conducted ABR and DPOAE tests, which showed no hearing changes in the CCR2^−/− group compared with the control group (Supplemental Fig. 1). These data demonstrate that CCR2^−/− retains normal hearing and can be accurately compared with our control group, CCR2^+/+.

Second, to study the effect of CCR2 deletion on cochlear injury in CSOM, we generated CSOM in CCR2^−/− and CCR2^+/+ mice and assessed OHC survival at 10 d following middle ear infection. The OHC survival rate was 84 ± 12.5% in the basal turn of CCR2^+/+ CSOM cochleae (Fig. 1A), whereas it was 63 ± 19.9% in the basal turn of CCR2^−/− CSOM cochleae (Fig. 1C). CCR2^−/− mice demonstrated increased OHC loss in the basal region relative to CCR2^+/+ mice (p = 0.036) (Fig. 1E). No OHC loss was observed in the middle region of the cochleae for both groups (Fig. 1B, 1D). No inner hair cell (IHC) loss was noted in the base, middle, or apical region in both CCR2^−/− and CCR2^+/+ mice.

To assess if CCR2 deletion affects macrophage migration in CSOM, we counted the number of macrophages using cryosection. The average number of macrophages per turn was 35.4 ± 8.8 in the CCR2^+/+ group (Fig. 2A–2C), whereas it was 22.3 ± 7.2 in the CCR2^−/−group (Fig. 2D–2F). Macrophage numbers were significantly reduced in CCR2^−/− CSOM (Fig. 2G. p < 0.001). These data revealed that CCR2 deletion resulted in a significant reduction of macrophages.

Next, to evaluate if CCR2 deletion affects the cytokine profile, we examined cochlear cytokines using a Luminex Bead Array. Of the 44 cytokines measured, 31 were reduced in the CCR2^−/− CSOM group, whereas 13 were elevated, compared with the CCR2^+/+ group (Fig. 3). Specifically, the relative CCL7 level was 8152 ± 5034 in the CCR2^−/− group, whereas it was 2253 ± 1414 in the CCR2^+/+ group. The relative IL-33 level was 187 ± 60 in the CCR2^−/− group, whereas it was 364 ± 81 in the CCR2^+/+ group (Fig. 3, dotted box). Both cytokines showed significant differences (p = 0.036 for CCL7 and p = 0.0047 for IL-33).

We checked if BLB permeability was altered by CCR2 deletion. Dextran was injected i.v. into the mice at 10 d following middle ear infection, and the cochleae were dissected 1 h after the injection. Dextran-GFP was not observed inside the cochleae in the CCR2^+/+, CCR2^+/+ CSOM, and CCR2^−/− CSOM groups (Fig. 4A–4C). The BLB permeability in the stria vascularis remained unchanged, even in the CCR2^+/+ group.

Macrophages are pivotal in the immune response within the inner ear, with functions ranging from protection to potential damage to hair cell integrity. Our prior research indicates their dominance in the cochlear immune responses leading to OHC loss in CSOM (7). Despite no direct bacterial invasion causing hair cell damage in CSOM, elevated proinflammatory cytokines suggest a link between macrophages and SNHL. Cochlear injury prompts a significant increase in macrophage numbers, with distinct phenotypic changes (23). Many studies have shown that macrophages migrate into the cochlea in response to damage caused by noise exposure, ototoxic drug treatment, and age-related degeneration (24–28). However, the exact role of these macrophages after cochlear injury is still unclear.

In this study, we found that CCR2 does play a protective role in CSOM, but monocyte migration was limited in CCR2^−/− relative to CCR2^+/+. CCR2 has been implicated in recruiting monocytes/macrophages to sites of inflammation during inflammatory responses (29, 30). Our findings are similar to Sautter et al.’s study on CCR2 in noise-induced hearing loss that CCR2 plays a protective role in OHC damage (15). Simultaneously, our study also found that CCR2 does influence monocyte migration and, in fact, shows decreased macrophage migration in CCR2^−/− relative to CCR2^+/+. It is plausible to suggest that the mechanism of OHC damage is different between a noise-induced model relative to a CSOM infection model.

Previous studies have shown that MCP1-CCR2 is expressed in brain endothelial cells and is associated with endothelial dysfunction in response to inflammation (31). This suggests that MCP-1 may contribute to BBB permeability other than purely driving leukocyte migration into the brain during inflammation (32, 33). In the inner ear, the BLB is composed of endothelial cells and can be a possible mechanism by CCR2 to cause macrophage-independent damage to OHCs. However, our data show that the BLB permeability did not change among CCR2^+/+ wild-type, CCR2^+/+ CSOM, and CCR2^−/− CSOM groups. This suggests that the OHC damage in CSOM may not be due to a change in BLB permeability.

Of the 44 cytokines measured, 31 were reduced in the CCR2^−/− CSOM group, whereas 13 were elevated, compared with the CCR2^+/+ group. Specifically, CCL7 was upregulated, whereas IL-33 was downregulated, in the CCR2^−/− group. CCL7 is one of the ligands of CCR2 and is known for its ability to attract monocytes, eosinophils, basophils, and T lymphocytes to sites of inflammation. CCL7 possesses binding affinity for other chemokine receptors besides CCR2, including CCR1, CCR3, and CCR5 (34, 35). The upregulation of CCL7 in the CCR2-deficient group suggests a compensatory mechanism. Because CCR2 is a receptor for chemokines such as CCL2 and CCL7, its absence might trigger an increase in CCL7 production to maintain the recruitment of monocytes and macrophages. However, it is important to consider that CCL7 is not solely reliant on CCR2 for its effects. IL-33, on the other hand, is a cytokine that belongs to the IL-1 family and acts as an alarmin released upon cellular damage or stress to alert the immune system (17, 18). IL-33 plays a role in the development of asthma, atopic allergies, anaphylaxis, and various other inflammatory diseases by inducing proinflammatory cytokines and chemokines and enhancing Th2 immune responses (36). The downregulation of IL-33 in the CCR2^−/− CSOM group indicates a potential reduction in type 2 immune response and associated inflammatory processes, which might affect the overall immune response dynamics in the context of CSOM. The reduction in anti-inflammatory mechanisms and the indirect increase in proinflammatory mediators could explain the increase in OHC loss in our CSOM model. Further studies are needed to understand the relationship between IL-33 and CCR2^−/− models.

Taken together, this study investigated the role of CCR2 in CSOM. Mice lacking CCR2 had increased OHC loss despite having fewer macrophages in the inner ear than control animals. CCR2 deletion resulted in CCL7 accumulation and IL-33 reduction. Interestingly, CCR2 deletion does not change BLB permeability. These findings suggest that CCR2 may play a protective role in this type of ear infection, potentially by regulating macrophage function in the inner ear. Further investigation is needed to understand the exact mechanisms by which CCR2 protects against SNHL in CSOM.

The authors have no financial conflicts of interest.