Abstract

   Hearing relies on the transmission of auditory information from sensory
   hair cells (HCs) to the brain through the auditory nerve. This relay of
   information requires HCs to be innervated by spiral ganglion neurons
   (SGNs) in an exclusive manner and SGNs to be ensheathed by myelinating
   and non-myelinating glial cells. In the developing auditory nerve,
   mistargeted SGN axons are retracted or pruned and excessive cells are
   cleared in a process referred to as nerve refinement. Whether auditory
   glial cells are eliminated during auditory nerve refinement is unknown.
   Using early postnatal mice of either sex, we show that glial cell
   numbers decrease after the first postnatal week, corresponding
   temporally with nerve refinement in the developing auditory nerve.
   Additionally, expression of immune-related genes was upregulated and
   macrophage numbers increase in a manner coinciding with the reduction
   of glial cell numbers. Transient depletion of macrophages during early
   auditory nerve development, using transgenic CD11b^DTR/EGFP mice,
   resulted in the appearance of excessive glial cells. Macrophage
   depletion caused abnormalities in myelin formation and transient edema
   of the stria vascularis. Macrophage-depleted mice also showed auditory
   function impairment that partially recovered in adulthood. These
   findings demonstrate that macrophages contribute to the regulation of
   glial cell number during postnatal development of the cochlea and that
   glial cells play a critical role in hearing onset and auditory nerve
   maturation.

   Keywords: auditory nerve, cochlea, hearing onset, macrophage, glial
   cells, CD11b

Introduction

   The auditory nerve is the conduit for transmission of sound in the
   mammalian ear. Axonal projections of bipolar spiral ganglion neurons
   (SGNs) connect sensory hair cells (HCs) to central auditory processes
   in the brain. Two subpopulations of SGNs exist: type I, which account
   for 90%–95% of total SGNs, and type II, which comprise the remaining
   5%–10% of SGNs. The cell bodies of both neuron types are housed within
   Rosenthal’s canal (RC) and extend peripheral axons through the osseous
   spiral lamina (OSL) to form connections with HCs in the organ of Corti
   (OC). Type I SGNs extend a single axon to a single inner hair cell
   (IHC) and type II SGNs innervate multiple outer hair cells (OHC) with
   numerous axonal projections (Marrs and Spirou, [39]2012). SGNs begin to
   innervate cochlear HCs in a tonotopic pattern as early as embryonic day
   (E) 15.5 (Koundakjian et al., [40]2007). At this time, multiple type I
   SGNs extend axons past the IHC region, forming erroneous connections
   with OHCs. Similarly, type II SGN axons form mistargeted connections
   with IHCs (Echteler and Nofsinger, [41]2000; Huang et al., [42]2007;
   Druckenbrod and Goodrich, [43]2015). Between E17.5 and the first
   postnatal week in rodents, excessive type I and II axons are eliminated
   by retraction or pruning to produce mature innervation patterns
   (Echteler, [44]1992; Echteler and Nofsinger, [45]2000; Echteler et al.,
   [46]2005; Huang et al., [47]2007; Yu et al., [48]2013). This refinement
   process occurs prior to hearing onset, which commences between P12–P14
   (Song et al., [49]2006).

   SGNs and auditory nerve fibers are wrapped by neural crest-derived
   auditory glial cells (Hansen et al., [50]2001; Breuskin et al.,
   [51]2010). Type I SGN somas are covered by myelinating satellite cells
   and their axons are ensheathed by myelinating Schwann cells.
   Conversely, type II SGNs are enveloped by non-myelinating satellite and
   Schwann cells. Recent studies have demonstrated that migration,
   maturity and survival of SGNs is largely dependent on surrounding glial
   cells (Hansen et al., [52]2001; Stankovic et al., [53]2004; Jeon et
   al., [54]2011; Lang et al., [55]2011, [56]2015). Additionally,
   conditional deletion of Sox10, a transcription factor regulating glial
   cell differentiation, results in reduced glial numbers, abnormal SGN
   distribution, decreased SGN migration, and the appearance of nerve
   fibers that overshoot the OC and extend into the cochlear lateral wall
   (LW), revealing the important role glial cells play in proper SGN
   innervation (Mao et al., [57]2014). However, it is still unknown how
   glial cell numbers are regulated as the auditory nerve matures.

   Immune cells such as macrophages and microglia contribute to nerve
   refinement in the central nervous system (Paolicelli et al., [58]2011;
   Schafer et al., [59]2012; Cunningham et al., [60]2013; Miron et al.,
   [61]2013). Macrophages, monocyte-derived phagocytes, are members of the
   innate immune system and respond to various cytokine/chemokine signals
   to remove pathogens and apoptotic cells (Tang et al., [62]2006;
   Guipponi et al., [63]2007; Akil et al., [64]2015). Macrophages have
   also been detected in the adult cochlea following cochlear insult
   (Hirose et al., [65]2005; Lang et al., [66]2006; Sato et al., [67]2010;
   Kaur et al., [68]2015). Damaged SGNs, auditory glial cells and
   fibrocytes secrete pro-inflammatory signals that promote the
   infiltration of macrophages (Bas et al., [69]2015; Fuentes-Santamaría
   et al., [70]2017). Recent studies have shown that immune-related gene
   expression increases in the developing auditory nerve (Lu et al.,
   [71]2011; Bank et al., [72]2012; Calton et al., [73]2014). However, the
   role of macrophages in auditory development has not been characterized.

   Here, we report that macrophages are implicated in the clearance of
   glial cells during development. This was supported by a dynamic
   reduction in glial cell number and accompanying evidence of macrophages
   engulfing glial cells and degenerative axons. Depletion of activated
   macrophages in mice caused glial cell alterations, myelin dysfunction
   and transient edema of the stria vascularis. Additionally,
   macrophage-depleted mice exhibited transient auditory function
   impairment. Collectively, these findings reveal that macrophages play a
   novel role in glial cell maintenance during auditory nerve maturation.

Materials and Methods

Animals

   CBA/CaJ and CD11b^DTR/EGFP mice were bred in-house in a low-noise
   environment at the Animal Research Facility of the Medical University
   of South Carolina (MUSC; Charleston, SC, USA). All aspects of animal
   research were conducted in accordance with the guidelines of the
   Institutional Animal Care and Use Committee of MUSC and the Guide for
   the Care and Use of Laboratory Animals of the National Institutes of
   Health. The protocols (2290 and 2845) were approved by the IACUC of the
   MUSC. The original adult CBA/CaJ and CD11b^DTR/EGFP breeding pairs were
   purchased from The Jackson Laboratory (Bar Harbor, ME, USA). All mice
   received food and water ad libitum, and were maintained on a 12-h
   light/dark cycle. For mouse cochlear collections, the day of birth was
   referred to as postnatal day 0 (P0). Since sex is grossly
   indistinguishable until P14 in mice, pups of either sex were used in
   this study. Young adult mice of either sex were also used in this
   study.

Diphtheria Toxin Administration

   P4–5 CD11b^DTR/EGFP mice received intraperitoneal injections (IP) of
   diphtheria toxin (DTX; 20–25 ng/g of body weight; Sigma-Aldrich)
   reconstituted in double distilled H[2]O (ddH[2]O). It has been reported
   that vehicle-treated CD11b^DTR/EGFP mice and DTX-treated non-transgenic
   mice of the same background do not demonstrate auditory functional
   differences (Zhang et al., [74]2012). To assure age-matching between
   mice, CD11b^DTR/EGFP littermates injected with vehicle solutions and
   were used as controls.

BrdU Postnatal Injections

   Bromodeoxyuridine (BrdU, 100 mg/kg, Sigma-Aldrich) was administered to
   neonatal CBA/CaJ mice by IP injection daily for postnatal days P0–P3.
   BrdU incorporation analysis was measured at P3, P7, P14 and P21
   (details in the “Immunohistochemistry” section below).

Immunohistochemistry

   Cochleae were fixed with 4% paraformaldehyde solution for 1–2 h at room
   temperature (RT), decalcified with 0.12 M ethylenediamine tetra acetic
   acid (EDTA) at RT with stirring, and cryopreserved with 30% sucrose for
   30 min. The tissues were then embedded in Tissue-Tek OCT Compound (VWR,
   Radnor, PA, USA) and stained as whole-mount preparations or sectioned
   at 10 μm thickness. The primary antibodies used in this study include:
   rabbit anti-Iba1 (Wako Chemicals USA Inc., Richmond, VA, USA (1:250)),
   rat anti-CD11b (Serotec, Oxford, UK (1:150)), mouse anti-CD68 (Abcam,
   Cambridge, MA, USA (1:100)), rabbit anti-F4/80 (Abcam (1:200)), mouse
   anti-Neurofilament 200 (Sigma-Aldrich (1:200)), rabbit anti-Myosin VIIa
   (Proteus BioSciences, Ramona, CA, USA (1:200)), mouse anti-C terminal
   binding protein 2 (BD Biosciences, San Jose, CA, USA (1:200)) and goat
   anti-Sox10 (Santa Cruz Biotechnology, Santa Cruz, CA, USA (1:80)).
   Secondary antibodies were biotinylated and binding was detected by
   labeling with fluorescein (FITC)-conjugated avidin D, Texas
   Red-conjugated avidin D (Vector Labs, Burlingame, CA, USA) or Alexa
   Fluor Dyes (ThermoFisher Scientific, Waltham, MA, USA). Nuclei were
   counterstained with propidium iodide (PI) or
   4′,6-diamidino-2-phenylindole (DAPI).

   The mouse monoclonal anti-BrdU was produced by clone BU33 and raised
   against BrdU incorporated into DNA, or coupled to a protein carrier. It
   recognized proliferative cells in the nuclei of frozen sections of
   animals treated with an in vivo administration of BrdU. In addition to
   the immunohistochemistry steps described above, BrdU-labeled sections
   were treated with two moles of hydrogen chloride for 30 min and 0.1 M
   of sodium borate buffer for 5 min prior to biotinylation.

   Sections were examined on a Zeiss LSM5 Pascal (Carl Zeiss Inc., Jena,
   DE, Germany) confocal microscope, a Zeiss LSM 880 NLO or Leica TCS SP5
   (Leica Microsystems, Allendale, NJ, USA) confocal microscope. FITC and
   Texas Red signals were detected by excitation with the 488 nm and 543
   nm lines, respectively. Images were scanned at image scales of 225.0 μm
   (x) × 225.0 μm (y), 144.72 μm (x) × 144.72 (y) and 450.0 μm (x) × 450.0
   μm (y). Captured images were processed using Zen 2012 Blue acquisition
   software (Zeiss Inc.), Leica Application Suite X software (Version
   3.0.2.16120) and Adobe Photoshop CS6 (Adobe Systems Inc., San Jose, CA,
   USA).

Histology Quantification

   Quantitative analysis of macrophages, glial cells and proliferative
   cell numbers were determined using AxioVision 4.8 (Carl Zeiss, Inc.)
   software. Regions of interests were determined by outlining intact RC
   and OSL, defined as boundaries from the habenular opening to a proximal
   site near the spiral ganglia, areas using the software outline tool.
   Similar tonotopic region sizes were examined between different cochlear
   samples. Within each region of interest, total cell numbers were
   determined by counting PI or DAPI counterstained cell nuclei using the
   measurement tool. Measurements of macrophages, glial cells, neurons and
   proliferative cells were determined by counting cells immunolabeled for
   Iba1^+, Sox10^+, NF200^+ or BrdU^+, respectively, in each region of
   interest. At least three slides from each ear from each postnatal age
   were used for data collection and processed using statistical analysis
   described below.

Hair Cell and Synapse Quantification

   Whole mount preparations of cochleae from P7 and 1 month DTX-treated
   and control CD11b^DTR/EGFP mice were stained with Myosin VIIa to
   identify IHCs and OHCs. HC numbers were counted manually using whole
   mount preparations from 1 month DTX-treated and control CD11b^DTR/EGFP
   mice (3 animals per group). Ribbon synapses under IHC were
   immunostained with CtBP2. CtBP2^+ ribbons were measured manually from
   at least 10 IHCs in the apex, middle or base (3 animals per group).
   Confocal All images were taken with a Zeiss LSM 880 NLO using a 63×
   oil-immersion lens and acquired at 0.25 μm step size in the Z-axis in
   non-overlapping regions. Maximum projection images from confocal
   z-stacks were acquired with the same parameters described above. Care
   was taken to minimize pixel saturation while imaging each z-stack.

Tissue Collection and Total RNA Isolation

   Postnatal CBA/CaJ mice were euthanized and their cochleae were promptly
   collected. Microdissection was performed to remove the outer bony
   cochlear shell, cochlear LW and the majority of the sensory epithelium,
   preserving the modiolus portion of the cochlea. For RNA isolations, the
   left and right ear cochlea preparations from a single mouse were pooled
   for individual samples. Total RNA was purified from cochlea
   preparations using the miRNeasy Mini Kit (Qiagen Inc., Germantown, MD,
   USA) according to the manufacturer’s instructions.

Microarray Data Analysis

   A microarray dataset of mouse auditory nerve development from our group
   (NCBI Gene Expression Omnibus accession [75]GSE59417; Lang et al.,
   [76]2015) was used for comparative analysis. The dataset contains
   expression data for auditory nerve samples collected at P0, 3, 7, 10,
   14 and 21 analyzed by Mouse 430 2.0 GeneChip (Affymetrix, Santa Clara,
   CA, USA). Raw hybridization data was normalized independently by both
   Robust Multi-array Average and MicroArray Suite 5.0 algorithms using
   Expression Console Software (Affymetrix). Differential expression was
   defined as absolute signal log ratio >0.5, ≥50% present gene detection
   scores and p ≤ 0.05 (Student’s t-test, unpaired) for P7 vs. P0
   (identifying 2391 probe sets). False discovery rate was estimated at
   1.7% based on iterative comparisons involving randomized sample group
   assignments. Among the 2391 probe sets, 1878 analysis-ready genes were
   found. Functional analysis and canonical function identification was
   assessed with Ingenuity Pathway Analysis Software version (IPA^®;
   QIAGEN, Redwood City, CA, USA).

Quantitative PCR (qPCR)

   Reverse transcription of total RNA was performed with the QuantiTect
   Reverse Transcription Kit (Qiagen) according to the manufacturer’s
   protocol. Briefly, genomic DNA was eliminated from RNA samples by
   incubation with gDNA Wipeout Buffer for 2 min at 42°C. Reverse
   transcription was performed with 50 ng of total RNA and Quantiscript
   Reverse Transcriptase in 20 μl reaction volumes incubated at 42°C for
   15 min and 95°C for 3 min. The following gene-specific amplification
   reagents were purchased from Qiagen: H2D1 (QT01657761), H2K1
   (QT01743700) C1qa (QT01057770), C3a receptor (C3ar; QT00251216),
   Ppfibp2 (QT00098161), and Runx1 (QT00100380). Quantitative PCR (qPCR)
   reactions were performed with the QuantiFast SYBR Green PCR Kit
   (Qiagen) using 1 μl of cDNA and a LightCycler 480 (Roche Diagnostics,
   IN, USA). Negative controls included reactions lacking cDNA template
   and reverse transcriptase. All reactions were performed in technical
   triplicate. Cycling parameters for PCR were: 50°C for 2 min, activation
   at 95°C for 5 min, and 40 cycles of 95°C for 10 s and 60°C for 30 s.
   Analysis of qPCR data was conducted with the aid of LightCycler 480
   software (version1.5.0.39). Amplification efficiency of each target was
   determined according to the equation E = 10 (−1/S), where S is the
   slope of the standard curve generated from 10-fold serial dilutions of
   the DNA preparations. Relative expression levels were calculated using
   the ∆∆C[T] method that involved calculated amplification efficiencies
   and then normalized to reference genes Hprt and 18S.

Transmission Electron Microscopy

   For animals aged P7, cochleae were collected and perfused through the
   oval window with 0.5 ml of a fixative mixture comprised of 4%
   paraformaldehyde and 2% glutaraldehyde in 0.1 M phosphate buffer, pH
   7.4. For collection of animals aged P11 and P16, anesthetized animals
   were perfused via cardiac catheter with 10 ml of normal saline
   containing 0.1% sodium nitrite followed by 15 ml of a mixture of 4%
   paraformaldehyde and 2% glutaraldehyde in 0.1 M phosphate buffer, pH
   7.4. After removing the stapes and opening the oval and round windows,
   0.5 ml of the same fixative described above was perfused gently into
   the scala vestibuli through the oval window. Inner ears were dissected
   and immersed in fixative overnight at 4°C. Decalcification for P16
   cochleae was completed by immersion in 40 ml of 120 mM solution of
   EDTA, pH 7.0, with gentle stirring at RT for 2–3 days with daily
   changes of the EDTA solution. Cochlear tissues were postfixed with 1%
   osmium tetroxide-1.5% ferrocyanide for 2 h in the dark, then dehydrated
   and embedded in Epon LX 112 resin. Semi-thin sections approximately 1
   μm thick were cut and stained with toluidine blue. Ultrathin sections
   (70 nm thick) were stained with uranyl acetate and lead citrate and
   examined and imaged using a JEOL JEM-1010 transmission electron
   microscope (JEOL USA, Inc., Peabody, MA, USA).

Auditory Brainstem Response Analysis

   Auditory brainstem responses (ABRs) were measured as previously
   described (Lang et al., [77]2011, [78]2015). Mice were anesthetized by
   IP injection of xylazine (20 mg/kg) and ketamine (100 mg/kg) and placed
   in a sound-isolation room. The acoustic stimuli were generated using
   Tucker Davis Technologies equipment System III (Tucker-Davis
   Technologies, Gainsville, FL, USA) and a SigGen software package
   (Version 4.4.1). Calibration was completed using an IPC Microphone
   System (PCB Piezotronics Inc., Depew, NY, USA) in a probe tube clipped
   to the pinna. The signals were delivered into the animal ear canal
   through a 3–5 mm diameter plastic tube. ABRs were evoked at half octave
   frequencies from 4 kHz to 45.2 kHz with 5 ms duration tone pips with
   cos^2 rise/fall times of 0.5 ms delivered at 31 times/s. Sound levels
   were reduced in 5 dB steps from 90 dB SPL to 10 dB SPL to determine
   thresholds. Distortion product otoacoustic emission testing was
   performed in mice under general anesthesia as previously described
   (Lang et al., [79]2010). Briefly, distortion product otoacoustic
   emissions (DPOAEs) were measured using a Tucker Davis Technologies
   equipment RZ6 system (Tucker Davis Technologies, Gainsville, FL, USA)
   and SigGen software using an ER-10B+ Lo Noise^TM Microphone System
   (Etymotic Research Inc., Grove Village, IL, USA). The acoustic assembly
   was tightly enclosed in the ear. Primary tones were swept from f2 =
   5.6–45.2 kHz with f1/f2 ratio of 1.2. Physiological results of each
   mouse were analyzed for individual frequencies, and then averaged for
   each of these frequencies from 5.6 kHz to 45.2 kHz.

Experimental Design and Statistical Analyses

   Biological sample sizes were determined based on similar experiments
   from previous studies conducted by our laboratory (Jyothi et al.,
   [80]2010; Lang et al., [81]2015, [82]2016) and other labs (Yu et al.,
   [83]2013; Coate et al., [84]2015) with the goal of achieving stringent
   and accurate measures of difference, namely statistical power of ≥80%
   and significance level of ≤0.05. Statistical analyses were performed
   using GraphPad Prism 7 software (GraphPad Software Inc., La Jolla, CA,
   USA) for Windows. Quantitative data are expressed as mean ± SEM, unless
   otherwise specified. Sample size is indicated for each figure.
   Differences across groups with multiple comparisons were analyzed with
   One-way ANOVA with post hoc Bonferroni Multiple Comparison tests.
   Differences for single pairwise comparisons were analyzed using
   two-tailed, unpaired Student’s t-tests. Statistical significance was
   defined as a p value of ≤0.05; all significance values are indicated.
   Specific p-values and test statistics are reported in the table below.
   Only significant values are reported in the “Results” section.

Results

Glial Cell Numbers Decrease after the Second Postnatal Week

   SGNs undergo a rigorous course of refinement and retraction during
   postnatal development. Glial cells are in tight junction with SGNs. To
   determine changes in glial cell numbers during auditory nerve
   development we collected postnatal spiral ganglia at the ages of P0,
   P3, P7, P14 and P21 and calculated the density of Sox10^+ cells. Sox10
   transcription factor is a key developmental regulator expressed in both
   immature and differentiated glial cells of the auditory nerve (Britsch
   et al., [85]2001; Breuskin et al., [86]2010; Finzsch et al., [87]2010).
   Quantitative analyses of Sox10^+ cells revealed that the highest
   density of glial cells in the OSL and RC occurred around P3–P7 (Figures
   [88]1A–D). After the first postnatal week, glial cell densities
   decreased. This temporal profile of glial cell numbers is in agreement
   with previous findings that the auditory nerve achieves a mature
   innervation pattern by the end of the first postnatal week in rodent
   models (Echteler, [89]1992; Huang et al., [90]2007; Coate et al.,
   [91]2015; Druckenbrod and Goodrich, [92]2015).

Figure 1.

   [93]Figure 1
   [94]Open in a new tab

   Increased glial cell numbers and proliferation correlate with auditory
   nerve refinement in postnatal mouse cochleae. (A,B) Confocal imaging of
   glial cell distribution in the postnatal cochlea. Sox10^+ glia (green)
   and propidium iodide (PI; red) counterstained nuclei appeared in the
   osseous spiral lamina (OSL) (A) and Rosenthal’s canal (RC) (B) of P3, 7
   and 21 mouse cochleae. Scale bars = 50 μm. (C,D) Quantitative analysis
   for Sox10^+ cells in the OSL (C) and RC (D) in P0, 3, 7, 14 and 21
   cochleae. Data bars represent mean density and error bars represent
   SEM. Two-tailed, unpaired Student’s t-tests were used to compare
   densities at P7 to the other time points for each turn (*p < 0.05; **p
   < 0.01; ***p < 0.001; n = 4–7 ears). The three cochlear turns are
   displayed: apical (white bars), middle (gray bars), and basal (black
   bars). (E) Dual-immunolabeling for Sox10 (green) and BrdU (red)
   identifies proliferative glial cells in the P7 RC. Cochlear samples
   were from mice that received a single intraperitoneal (IP)
   administration of BrdU at P0, P1 and P2. Scale Bars = 50 μm. (E’)
   Higher magnification of inset from (E) shows proliferative glia
   (arrows) and non-proliferative glia (arrowheads). Scale bar = 20 μm.
   (F,G) Quantitative analysis of Sox10^+/BrdU^+ cells in the OSL (F) and
   RC (G) for P3, 7, 14 and 21. One-way ANOVA followed by Bonferroni
   Multiple Comparison tests were used to compare densities between time
   points (*p < 0.05; **p < 0.01; n = 3–5 cochlear samples).
   Quantifications for the different cochlear turns are demonstrated:
   apical (white bars), middle (gray bars) and basal (black bars). Data
   bars represent mean density and error bars represent SEM. (H,I)
   Dual-immunolabeling for Sox10 (green) and C-Cas3 (cleaved-Caspase 3;
   red) reveals apoptotic glial cells (arrowheads) and apoptotic non-glial
   cells (arrow) in P3 (top) and P7 (bottom) cochleae. Scale Bars = 20 μm
   in (H), Scale Bars = 20 μm in (I). Apex, apical turn; Mid, middle turn;
   Base, basal turn.

   To determine the proliferative ability of glial cells we utilized the
   BrdU assay. Cochlear samples were collected for analysis 3 days after
   BrdU injection. Double-labeling for BrdU and Sox10 revealed the
   presence of proliferative glial cells in the postnatal auditory nerve
   (Figures [95]1E,E′). Quantitative analysis of Sox10^+/BrdU^+
   double-labeled cells revealed that the highest occurrence of glial cell
   proliferation occurred at P3–P7. Glial cell proliferation significantly
   decreased after the second postnatal week, after mature auditory nerve
   circuitry is established and SGNs are myelinated (Figures [96]1F,G).

   The early phase of apoptosis is a rapid event necessary for the
   development of neural tissues and is often characterized by the
   cleavage of Caspase-3 (Jänicke et al., [97]1998; Hall, [98]1999; Porter
   and Jänicke, [99]1999). To determine if the temporal alterations in
   glial cells were due to apoptosis we examined cochlear samples from
   mice aged P3 through P14 for the presence of Sox10/cleaved-Caspase 3
   positive cells. Very few cleaved-Caspase 3^+ glial cells were
   identified in P3 and P7 cochleae (Figures [100]1H,I, arrowheads).
   Interestingly, one Sox10^- cell stained positive for cleaved-Caspase 3
   in the P3 cochlea. The morphology of this apoptotic cell suggested it
   to be a SGN (Figure [101]1H, arrow), supporting previous research that
   found the highest incidences of SGN apoptosis in P4–6 rodent auditory
   nerves (Echteler et al., [102]2005). No cleaved-Caspase 3 positive
   cells were found in P14 cochlea (data not shown).

Immune-Related and Axon Guidance Gene Expression Correlate with Auditory
Nerve Maturation

   We utilized our published microarray dataset of developing mouse
   auditory nerves (NCBI GEO:[103]GSE59417; Lang et al., [104]2015) to
   identify molecular regulators and pathways involved in postnatal nerve
   development. We performed a comparison analysis of approximately 45,000
   gene probe sets to identify genes that were differentially expressed
   between P0 and P7 (absolute signal log ratio >0.5; ≥50% present gene
   detection scores, Student’s unpaired t-test p ≤ 0.05). This analysis
   yielded 2391 probe sets with up- and downregulated expression values
   and an estimated false discovery rate of 1.7% (Figure [105]2A).
   Functional enrichment analysis was conducted using 1878 analysis-ready
   genes found among the 2391 probe sets. Pathway enrichment analysis
   detected 30 unique signaling pathways that were significantly enriched
   (Table [106]1), including Axonal Guidance Signaling, which is expected
   given that auditory nerve maturation occurs during the first postnatal
   week. Interestingly, several immune response pathways, including GADD45
   Signaling and Complement System, were also significantly enriched,
   suggesting that immune-related activities are critical during auditory
   nerve development.

Figure 2.

   [107]Figure 2
   [108]Open in a new tab

   Genes associated with axonal guidance and immune responses are
   differentially expressed in the postnatal auditory nerve. (A)
   Expression profiles of genes differentially expressed between P0 and P7
   in postnatal auditory nerves collected at P0, P3, P7, P10, P14 and P21.
   A total of 2391 probe sets, corresponding to 1878 genes, were
   identified comparing expression at P0 vs. P7 with absolute signal log
   ratio >0.5, ≥50% present gene detection scores and p ≤ 0.05 (Student’s
   t-test). Colorimetric scaling (Z-standardization) for the heatmap is
   shown at the bottom. (B–D) Quantitative PCR (qPCR) validation for gene
   expression of (B) axonal guidance genes (C), major histocompatibility
   class (MHC) II genes, and (D) complement pathway genes at P0 (black),
   P7 (gray) and P21 (white) is displayed. Microarray expression data at
   P0, P7 and P21 are demonstrated for each gene. One-way ANOVA with post
   hoc Bonferroni Multiple Comparison tests were used to compare gene
   expression between the postnatal days. Data bars represent average mean
   of experiments run in triplicate; error bars represent SEM; *p < 0.05;
   **p < 0.01, ***p < 0.001; n = 3.

Table 1.

   Ingenuity pathway analysis of canonical pathways identified by
   differentially expressed genes.
   Canonical pathway                                         p-value
   Cell Cycle: G2/M DNA Damage Checkpoint Regulation         1.74E-04
   Axonal Guidance Signaling                                 6.17E-04
   GADD45 Signaling                                          3.02E-03
   Cell Cycle Control of Chromosomal Replication             3.55E-03
   Superpathway of Serine and Glycine Biosynthesis I         3.98E-03
   dTMP De Novo Biosynthesis                                 4.90E-03
   Melatonin Degradation II                                  4.90E-03
   Estrogen-mediated S-phase Entry                           5.50E-03
   Osteoarthritis Pathway                                    8.51E-03
   Adipogenesis pathway                                      9.77E-03
   Serine Biosynthesis                                       1.15E-02
   Glycine Degradation (Creatine Biosynthesis)               1.23E-02
   Triacylglycerol Biosynthesis                              1.41E-02
   Hereditary Breast Cancer Signaling                        1.66E-02
   Phospholipases                                            2.04E-02
   Pyrimidine Deoxyribonucleotides De Novo Biosynthesis I    2.29E-02
   Antioxidant Action of Vitamin C                           2.75E-02
   Role of BRCA1 in DNA Damage Response                      2.82E-02
   Bladder Cancer Signaling                                  3.09E-02
   Virus Entry via Endocytic Pathways                        3.31E-02
   Sphingomyelin Metabolism                                  3.39E-02
   Sphingosine-1-phosphate Signaling                         3.39E-02
   Fatty Acid Activation                                     3.55E-02
   Wnt/β-catenin Signaling                                   3.72E-02
   Cell Cycle: G1/S Checkpoint Regulation                    4.17E-02
   Cholesterol Biosynthesis I                                4.68E-02
   Cholesterol Biosynthesis II (via 24,25-dihydrolanosterol) 4.68E-02
   Cholesterol Biosynthesis III (via Desmosterol)            4.68E-02
   ATM Signaling                                             4.79E-02
   Complement System                                         4.90E-02
   [109]Open in a new tab

   We performed qPCR to validate the temporal expression patterns of two
   key regulatory genes associated with axon guidance cues, Ppfibp2 and
   Runx1, which were identified as differentially expressed in our
   analysis. Ppfibp2 is a gene that encodes liprin beta-1 protein, a
   positive regulator of axonal extension and synapse formation (Dunah et
   al., [110]2005; Astigarraga et al., [111]2010). Runx1 is a
   transcription factor that negatively regulates axonal projections and
   neuron maintenance (König et al., [112]2011; Yoshikawa et al.,
   [113]2015). Although not significant, Ppfibp2 showed a trend for
   increased gene expression from P0 to P7 and decreased by P21, in
   agreement with the microarray results (Figure [114]2B). We also found
   that Runx1 expression significantly decreased at P7, when compared to
   P0 and P21 (Figure [115]2B), supporting the microarray results.
   Together, expression profiles of these genes indicate that P7 is a
   critical time point in auditory nerve maturation in the mouse cochlea.

   The antigen presentation pathway was significantly enriched according
   to our functional analysis (Table [116]1). This pathway includes major
   histocompatibility class (MHC) class I genes, which signal for the
   targeting of nerves for elimination during synaptic pruning in spinal
   motorneurons (Thams et al., [117]2008). We performed qPCR to validate
   the expression of two MHC Class I genes, H2-D1 and H2-K1, which were
   identified as differentially expressed in our microarray analysis. Gene
   expression of H2-D1 and H2-K1 was upregulated in P7 auditory nerves,
   compared to P0 and P21 nerves, validating the expression values
   represented in the microarray dataset (Figure [118]2C). Complement
   signaling also mediates synaptic pruning and axon elimination in
   central and peripheral nerves (Schafer et al., [119]2012; Bialas and
   Stevens, [120]2013; Bahrini et al., [121]2015). We investigated the
   expression of two complement-related components identified as
   upregulated in our microarray dataset, complement component C1q, the
   molecular initiator of the classical complement pathway, and C3aR, a
   receptor for the central complement component for all complement
   pathways. qPCR confirmed that gene expression of C1q and C3aR was
   upregulated in the P0–P7 auditory nerve (Figure [122]2D).

Macrophage Numbers Increase during Auditory Nerve Maturation

   We further addressed the hypothesis that immune activities contribute
   to auditory nerve refinement by examining macrophage activity in the
   postnatal auditory nerve. Iba1, a highly conserved cytoplasmic protein,
   is specific to cells of monocytic lineage (Ohsawa et al., [123]2000;
   Sasaki et al., [124]2001). Analysis of Iba1^+ cells revealed gradual
   increases in macrophage numbers in RC and the OSL from P0 to P3,
   reaching peak numbers at P7, and decreasing after P14 and P21 (Figures
   [125]3A–D). This increase in macrophage numbers coincides temporally
   with auditory nerve maturation and is subsequent to the glial cell
   increase.

Figure 3.

   [126]Figure 3
   [127]Open in a new tab

   Iba1^+ macrophage numbers and proliferation peak in the first postnatal
   week. (A,B) Confocal images show the temporal distribution of Iba1^+
   (green) macrophage cells in the OSL (A) and RC (B) of P0, P7 and P21
   cochleae. Scale bars = 50 μm. (C,D) Quantitative analyses of Iba1^+
   macrophage densities in the OSL (C) and RC (D) of postnatal cochleae
   aged P0–P21 reveal an increase in macrophage numbers during nerve
   refinement. Quantifications for the different cochlear turns are
   displayed: apical (white bars), middle (gray bars) and basal (black
   bars). Data bars represent mean density and error bars represent SEM.
   Two-tailed, unpaired Student’s t-tests were used to compare Iba1^+
   macrophage densities at P7 to other postnatal time points in each turn
   (*p < 0.05; **p < 0.01; ***p < 0.001; n = 3–7 ears). (E–G) Confocal
   images show proliferating macrophages in the auditory nerve of a P7
   cochlea. Macrophages are labeled with Iba1 (green) and proliferating
   cells are labeled with BrdU (red). Cochlear samples were from mice that
   received a single IP administration of BrdU at P0, P1 and P2. Scale
   bars = 50 μm. (E′) A high magnification image from enclosed area shown
   in (E) identifies proliferative macrophages (arrowheads) and a
   proliferative non-macrophage cell (arrow). (F,G) Quantitative analysis
   of BrdU/Iba1^+ macrophage densities in the OSL (F) and RC (G) of
   postnatal cochleae aged P3–P21 reveals that most macrophage
   proliferation occurs during the first postnatal week (P3–7). One-way
   ANOVA with post hoc Bonferroni Multiple Comparison tests were used to
   compare densities between time points (*p < 0.05; **p < 0.01; ***p <
   0.001 n = 3–7 ears). Quantifications for the different cochlear turns:
   apical (white bars), middle (gray bars) and basal (black bars). Data
   bars represent mean density and error bars represent SEM. Apex, apical
   turn; Mid, middle turn; Base, basal turn.

   To determine if the increase in macrophage density was due to the
   proliferation of resident macrophages, we injected BrdU into P0 pups
   and assessed macrophage proliferation. Dual labeling of Iba1^+/BrdU^+
   cells demonstrated that the highest numbers of proliferative
   macrophages are seen during the first postnatal week (Figures
   [128]3E–G). Fewer proliferative macrophages were seen in the second and
   third postnatal weeks.

Postnatal Macrophages Display Activated Phenotypes

   We performed immunohistochemical analysis of Iba1^+ macrophages in
   postnatal tissue using cochlear whole mount preparation from P7 mice.
   This allowed visualization of entire cell morphologies. Macrophages
   exist in several distinct morphological states that are related to
   their functional roles. Ramified appearance indicates surveillance
   function, while rounded or ameboid shapes suggest phagocytic functions
   (Kaur et al., [129]2007; Parakalan et al., [130]2012). Iba1^+
   macrophages were present in multiple locations of the cochlea including
   RC, the OSL, along the outskirts of and within the OC and in the
   cochlear LW (Figure [131]4A). Investigation of macrophage morphology
   revealed that some macrophages were ramified (Figure [132]4Ai) while
   others were ameboid (Figures [133]4Aii,iii).

Figure 4.

   [134]Figure 4
   [135]Open in a new tab

   Macrophage activation occurs in the developing auditory nerve. (A)
   Confocal z-stack 3D rendering in mid-apical portion of a P7 mouse
   cochlea immunolabeled with Iba1 (green). Numerous Iba1^+ macrophages
   are present in the OSL and RC with an ameboid shape, suggesting these
   Iba1^+ cells are the activated macrophages. High magnification images
   from (A) shows a ramified macrophage (i) and an ameboid macrophage in
   the OSL (ii) and RC (iii). LW, lateral wall; OC, organ of Corti; OSL,
   osseous spiral lamina; RC, Rosenthal’s canal. Scale bar = 50 μm. (B)
   Macrophages co-labeled with Iba1 (green) and F4/80 (red) antibodies
   (arrowhead). (C) An image of co-labeled macrophage cells with Iba1
   (green) and CD11b (red), reveal macrophages at P7 are mature leukocytes
   (arrowhead). (D) Co-localization of Iba1 (green) and CD68 (red), a
   lysosomal glycoprotein marker, reveals phagocytic activity of cochlear
   macrophages (arrowhead). (A–D) Scale bars = 10 μm. (E) Bar graph
   depicting the percentage of Iba1^+ cells that co-label with F4/80,
   CD11b and CD68. Percentages calculated from three to four mouse samples
   for each probe.

   To further characterize the macrophage phenotype the postnatal auditory
   nerve, we probed for a panel of lysosomal and surface protein markers
   that are expressed by mature monocytes and during immune cell
   activation. We first performed immunostaining for F4/80, a marker for
   glycoprotein presented on the surface of mature monocytes (Austyn and
   Gordon, [136]1981; Hume et al., [137]1984). Over 80% of Iba1^+ cells in
   the P7 cochlea stained positive for F4/80, confirming the
   differentiated phenotype and mature identity of the Iba1^+ cells in the
   auditory nerve (Figures [138]4B,E). Next, we assessed the activation
   and phagocytic profile of the immune cells in P3–7 auditory nerves by
   probing for immunoreactivity of CD11b, a cell surface integrin
   indicative of macrophage/microglia activation, and CD68, a lysosomal
   protein expressed during debris degradation (Barros et al., [139]2013;
   Hickman et al., [140]2013; Bennett et al., [141]2016). The presence of
   Iba1^+/CD11b^+ and Iba1^+/CD68^+ cells suggested that a portion of
   macrophages engulfed particles and were actively degrading debris and
   cell fragments (Figures [142]4C–E).

Macrophages Phagocytose Glial Cells and Axonal Fragments

   To further investigate the physical interaction between auditory glial
   cells and macrophages, 3D confocal microscopy was employed. We chose to
   examine the auditory nerve at P7, when cochlear macrophage numbers were
   the highest. Dual immunostaining for Sox10^+ glia and Iba1^+
   macrophages in RC revealed that macrophages were in close proximity to
   glial cells (Figures [143]5A–B″′). Upon closer examination, it was
   apparent that Sox10^+ glial cell nuclei were engulfed within the
   cytoplasm of macrophages (Figure [144]5B′). Internalization of glia by
   macrophages was confirmed by maximum projection analysis and 3D
   reconstructions of sections stained for Sox10 and Iba1 (Figure
   [145]5B″″). We then examined macrophage-axon relationships in the P7
   auditory nerve. We assessed neurofilament 200^+ (NF200) nerve fibers
   that extend through the OSL to reach their HC targets. Dual
   immunostaining of Iba1 and NF200 revealed that macrophages directly
   contact SGN axons in P7 cochleae (Figures [146]5C–C″′). Maximum
   projection images and 3D reconstruction confirmed that macrophages
   phagocytose and digest axonal fragments (Figure [147]5C″″).

Figure 5.

   [148]Figure 5
   [149]Open in a new tab

   Macrophages engulf glial cells and the fragments of auditory nerve
   fibers. (A–A″″) Confocal images of Sox10^+ glial cells (green) and
   Iba1^+ macrophages (red) in the P7 auditory nerve. Macrophages are in
   close proximity of glial cells. 4′,6-diamidino-2-phenylindole (DAPI;
   blue) was used to counterstain cell nuclei. Panel (A′) shows enlarged
   image from enclosed area. (A–A″″) Scale bars = 10 μm. (B–B″″) Another
   confocal image of Sox10^+ cells (green) and Iba1^+ macrophages (red)
   from a thick section of P7 auditory nerve. Panel (B″″) shows stacking
   and 3D rendering of the enclosed area reveals an Iba1^+ macrophage
   (red) engulfing a Sox10^+ glial cell (green; arrowhead). (B–B″′) Scale
   bars = 15 μm. (C–C″″) Confocal imaging of a P7 whole-mount cochlear
   preparation stained with Iba1 (red) and NF200 (green). Panel (C″″)
   shows stacking and 3D rendering of enclosed area shows an Iba1^+
   macrophage phagocytosing a NF200^+ nerve fiber fragment (arrowhead).
   (C–C″′) Scale bars = 50 μm.

   Using electron microscopy, we investigated the ultrastructure of
   different macrophage phenotypes in the auditory nerve of P7 cochleae.
   As expected, macrophages showed an active phenotype, including
   densely-packed cytoplasm filled with multiple lysosomes, abundant rough
   endoplasmic reticulum, electron-dense lipid bodies and phagoliposomes
   (Figures [150]6A–C). We also detected several glial cells that were not
   enclosing nerve fibers or neuron cell bodies (Figure [151]6D,
   asterisks). Interestingly, these free-floating, or supernumerary, glial
   cells were in close proximity to macrophages demonstrating activated
   morphologies (Figure [152]6D’). These observations correspond with the
   findings of glia-macrophage interactions identified by immunostaining
   and 3D reconstruction.

Figure 6.

   [153]Figure 6
   [154]Open in a new tab

   Ultrastructure of cochlea shows macrophages engulf degenerating axons
   and glial cells. (A–C) Micrographs of the P7 auditory nerve reveals
   cochlear macrophages (M) with activated morphologies including densely
   packed cytoplasm filled with multiple lysosomes, abundant rough
   endoplasmic reticulum, electron dense lipid bodies and phagoliposomes.
   (A,B) Scale bars = 2 μm, (C) Scale bar = 800 nm. (D) Electron
   micrograph shows myelinated axons in the OSL of a P7 mouse. Scale bar =
   4 μm. Asterisks demonstrate supernumerary glial cells that are not
   juxtaposed to nerve fibers. (D′) Enclosed area highlights an activated
   macrophage (M) approaching a supernumerary glial cell (asterisk). Scale
   bar = 800 nm. (E) Micrograph of axons undergoing development-related
   degeneration (arrows). Scale bar = 2 μm. (E′) Enclosed area shows a
   macrophage (M) encroaching a glial cell with several degenerating axons
   within the cytoplasm (arrows). Scale bar = 800 nm. (F) The
   ultrastructure of the OSL area in another P7 mouse cochlea shows a
   macrophage (M) phagocytosing a glial cell (G) wrapping a myelinated,
   degenerative axon (A). Arrowheads highlight myelin blebs along
   degenerating axons and within the active macrophage. Scale bar = 800
   nm. (G) Micrograph of a macrophage (M) engulfing a non-myelinating
   glial cell (G) and associated, degenerating axon (A). Scale bar = 600
   nm. All images were composed from analysis of at least three mouse
   cochleae.

   The ultrastructural analysis also detected SGN axons that were
   undergoing degeneration (Figure [155]6E, arrows). In some cases, a
   single glial cell ensheathed several degenerating axons (Figure
   [156]6E’). Active macrophages were seen encroaching on these
   degenerating SGN axons (Figure [157]6E’). There were also several
   macrophages that surrounded glia-wrapped degenerating axons (Figure
   [158]6F,G). Both myelinated and unmyelinated degenerating axons were
   engulfed by macrophages. These macrophages possessed myelin-like debris
   within their cytoplasm (Figure [159]6F, arrowheads), indicating that
   both glial cells and damaged or degenerative axons are phagocytosed by
   cochlear macrophages.

Transient Depletion of Postnatal Macrophages Increases Glial Cell Numbers

   To determine if macrophages are required to remove postnatal glial
   cells, we employed the CD11b^DTR/EGFP mouse model to transiently
   deplete active macrophages in the developing auditory nerve. Murine
   cells have low binding affinity for DTX. CD11b^DTR/EGFP mice express
   DTX receptors under the CD11b promoter. Administration of DTX to
   CD11b^DTR/EGFP mice results in a temporary depletion of only CD11b^+
   cells, with reductions lasting for 3–5 days (Cailhier et al.,
   [160]2005; Stoneman et al., [161]2007; Ueno et al., [162]2013). To
   transiently deplete CD11b^+ macrophages before the completion of
   auditory nerve maturation, DTX was administered to CD11b^DTR/EGFP pups
   at P4–5. Comparison of Iba1^+ macrophage numbers in DTX- treated and
   control mice at P7 revealed significantly reduced macrophage numbers
   (Figures [163]7A–C). This corresponded with earlier results that
   demonstrated that a majority of Iba1^+ cells also stained positive for
   CD11b (Figure [164]4C). Consequences on glial cell numbers were
   assessed by immunostaining with Sox10 in the OSL and RC (Figure
   [165]7D). We found that glial cells in the auditory nerve of
   DTX-treated mice were significantly increased compared to controls
   (Figures [166]7E,F). These results support the hypothesis that active
   macrophages regulate glial cell numbers during auditory nerve
   maturation.

Figure 7.

   [167]Figure 7
   [168]Open in a new tab

   DTX treatment results in decreased macrophages and increased glial
   cells in the auditory nerve of postnatal CD11b^DTR/EGFP mice. (A)
   Confocal images of Iba1^+ macrophages (green) and cell nuclei (PI-red)
   in middle turn RC of control (top panel) and DTX-treated (bottom panel)
   P7 cochleae. Scale bar = 25 μm. (B,C) Quantification of Iba1^+ cells
   reveals fewer macrophages in DTX-treated OSL (B) and RC (C), in
   comparison to controls. (D) Confocal images of Sox10^+ glial cells in
   middle turn RC of control (top panel) and DTX-treated (bottom panel) P7
   cochleae. Scale bars = 25 μm. (E,F) Quantification of Sox10^+ glial
   cells reveals more glia in DTX-treated OSL (E) and RC (F), in
   comparison to controls. Two-tailed, unpaired Student’s t-tests were
   used for statistical analyses (*p < 0.05; **p < 0.01; ***p < 0.001; n =
   3–7 cochleae per treatment). Bars represent mean density and error bars
   represent SEM. DTX, diphtheria toxin; Apex, apical turn; Mid, middle
   turn; Base, basal turn.

   During the first postnatal week, specifically between P4–6, SGNs
   undergo a reduction in cell numbers due to an increase in apoptosis
   (Echteler et al., [169]2005). To ensure that DTX treatment did not
   alter the SGN population, we probed for NF200 in DTX-treated and
   control auditory nerves. There was no apparent difference in SGN
   distribution in DTX-treated and control auditory nerves (Figures
   [170]8A,D). This suggests that the mechanism of glial cell maintenance
   in macrophage-depleted nerves is not SGN-dependent. Whole mount
   preparations immunolabeled with MyosinVIIa were analyzed to investigate
   possible HC disruptions following DTX treatment. We found no detectable
   differences in the IHC or OHC numbers when comparing DTX-treated and
   control adult auditory nerves (Figures [171]8B,F,G). To investigate the
   effect of macrophage depletion on final afferent synapse distribution
   patterns, we immunostained whole-mount preparations of basilar
   membranes with anti-C-terminal binding protein (CtBP2) in adult
   CD11b^DTR/EGFP mice. Immunostaining of CtBP2 revealed no apparent
   difference in synapse distribution after DTX treatment (Figures
   [172]8C,E).

Figure 8.

   [173]Figure 8
   [174]Open in a new tab

   DTX-treated mice have relatively normal neuronal cells, hair cells
   (HCs) and afferent synapses. (A) NF200 (green) stained P7 cochlea show
   no difference between neurons of control and DTX-treated mice. PI (red)
   was used to counterstain nuclei. Scale bar = 25 μm. (B) Confocal
   imaging of Mysoin VIIa^+ HCs in the OC of P7 control and DTX-treated
   mice. The intact three rows of outer hair cells (OHCs) and one row of
   inner hair cells (IHCs) were presented in both treatment conditions.
   Scale bar = 20 μm. (C) Immunostaining of Myosin VIIa (red) and CtBP2
   (green) revealed no significant loss of afferent synapses under IHCs in
   1 month old DTX-treated mice compared to controls. DAPI (blue) was used
   to counterstain cell nuclei. Scale bar = 20 μm. (D) Quantification of
   NF200^+ neurons in P7 mice reveal no significant difference in spiral
   ganglion neuron (SGN) numbers in DTX-treated and control auditory
   nerves.(E–G) Quantitative analysis of Myosin VIIa and CtBP2
   immunostaining in 1 month auditory nerves reveals no significant
   differences in IHC (F), OHC (G) or ribbon synapse (E) numbers of
   DTX-treated and control mice.Two-tailed, unpaired Student’s t-tests
   were used for statistical analyses. No statistical significance was
   determined. n = 3 mice per group. Bars represent mean density and error
   bars represent SEM. Apex, apical turn; Mid, middle turn; Base, basal
   turn.

Transient Depletion of Macrophages Results in Dysfunction of Myelinating
Glial Cells

   The production and compaction of myelin around SGN cell bodies and
   axons is a main function of auditory glial cells. Since myelin becomes
   more compact and mature between P10-P12 in rodent cochleae (Wang et
   al., [175]2013) we investigated the ultrastructure of the auditory
   nerves in P11 CD11b^DTR/EGFP mice. As expected, myelinated axons were
   seen in both control and DTX-treated auditory nerves (Figures
   [176]9A,B). DTX-treated nerves contained supernumerary glial cells that
   were not associated with nerve fibers or neuron cell bodies (Figure
   [177]9B). Myelinated SGNs were also seen in both control and
   DTX-treated nerves (Figures [178]9C,D); however, the myelin sheath
   surrounding SGNs of DTX-treated nerves showed modest alterations,
   including vacuolation and alveolated interruptions (Figure [179]9D,
   arrowheads). Nodes of Ranvier of DTX-treated nerves also demonstrated
   abnormalities, evidenced by robust myelin blebbing and folding in the
   paranodal regions (Figures [180]9E,F). Additionally, a group of
   myelinated axons were ensheathed by several Schwann cells (Figure
   [181]9G). High magnification revealed distinct cell boundaries were
   between the Schwann cells, indicating that supernumerary glial cells
   intermingle with and enwrap other myelinating Schwann cells (Figure
   [182]9G’). We investigated the auditory nerves of CD11b^DTR/EGFP mice
   at P16, a time after myelination is complete and after hearing onset.
   In P16 DTX-treated nerves, myelin appeared to recover and was
   comparable to that of control auditory nerves (Figures [183]9H,I).
   However, there were occasional cases of blebbing in the myelin
   surrounding neurons in DTX-treated nerves (Figure [184]9J).
   Additionally, few glial cells appeared to enwrap degenerating axons
   (Figure [185]9K).

Figure 9.

   [186]Figure 9
   [187]Open in a new tab

   Depletion of macrophages results in transient myelin and LW-related
   abnormalities. (A,B) Electron micrographs demonstrate myelinated axons
   in control (CTL) (A) and DTX-treated (DTX) (B) auditory nerves of P11
   CD11b^DTR/EGFP mice. A supernumerary glial cell can be found in the
   DTX-treated cochlea. Scale bars = 2 μm. (C,D) Micrograph illustrates
   myelinated neurons in control (C) and DTX-treated (D) auditory nerves.
   Arrowheads highlight vacuolations within the myelin of two neurons in
   the DTX-treated samples. Scale bars = 1 μm. (E,F) Micrographs of nodes
   of Ranvier in control (E) and DTX-treated (F) samples. Arrowhead shows
   thickened Schwann cell microvilli and myelin folding in the paranodal
   loops. Scale bars = 400 nm (G) Cluster of myelinated auditory nerve
   fibers in the OSL of P18 DTX-treated auditory nerves. Scale bars = 2
   μm. (G′) High magnification of the enclosed area in (G) reveals
   supernumerary glial cells wrapping other glial cells. Scale bar = 800
   nm. (H–J) Electron micrographs of neurons found in control (H) and DTX
   treated (I) auditory nerves of P16 CD11b^DTR/EGFP mice. (J) Arrowhead
   highlights blebbing in the myelin surround a neuron in the DTX-treated
   cochlea. Scale bars = 2 μm. (K) Glial cell containing several
   degenerative axons in the cytoplasm (arrowheads). Scale bar = 2 μm.
   (L,M) Micrographs of cochlear LWs from control (L) and DTX-treated (M)
   P11 CD11b^DTR/EGFP mice. Scale bars = 4 μm. (L′,M′) High magnification
   of the enclosed areas in L-M reveal blood vessel and endothelial cell
   morphology in control (L′) and DTX-treated (M′) cochleae. Scale bars =
   2 μm. (N,O) Micrographs of cochlear LWs from control (N) and
   DTX-treated (O) P16 CD11b^DTR/EGFP mice reveals DTX-treated LWs are
   able to recover. Scale bars = 4 μm. (N′,O′) High magnifications of the
   enclosed areas in (N,O) reveal blood vessel and endothelial cells in
   control (N’) and DTX-treated (O′) cochleae. Scale bars = 2 μm. G, glial
   cell; N, spiral ganglion neuron; Mc, marginal cell; Ic, intermediate
   cell; Bc, basal cell; Ec, endothelial cell; RBC, red blood cell.

   CD11b^+ cells have been found to reside in the cochlear LW of the adult
   cochlea. These cells, termed perivascular-resident macrophage-like
   melanocytes, remain tightly bound to LW blood vessels to control
   permeability within the intrastrial space of the cochlear LW (Zhang et
   al., [188]2012, [189]2013). We investigated the ultrastructure of the
   cochlear LW to determine if DTX-treatment altered LW cells (Figures
   [190]9L,M). Although marginal cells were maintained, losses of the
   interdigital processes of both marginal cells and intermediate cells
   with enlarged intrastrial space appeared in stria vascularis,
   indicating edema of the cochlear LW of P11 DTX-treated cochleae (Figure
   [191]9M). Endothelial cells surrounding blood vessels in DTX-treated
   LWs also showed ruffling in their cell membranes (Figure [192]9M’).
   This was not seen in endothelial cells of P11 control cochleae (Figure
   [193]9L’). Interestingly, examination of P16 DTX-treated and control
   cochleae revealed that edema of the stria vascularis was no longer
   present and endothelial cells surrounding LW blood vessels were
   comparable to those present in control cochleae (Figures [194]9N–O’).
   Our analysis of macrophage-depleted cochleae agrees with recent studies
   that reported recovery/self-repairing capability of the cochlear LW
   following damage (Mizutari et al., [195]2011; Stevens et al.,
   [196]2014; Li et al., [197]2017).

Macrophage Depletion Impairs Auditory Function of Postnatal and Young Adult
Mice

   We next investigated how macrophage depletion affected auditory
   function. ABR tests were performed at P15, after the onset of hearing,
   and at P21 and 1 month, when auditory nerve function is more mature. At
   P15, ABR thresholds of DTX-treated mice were higher than controls
   (Figure [198]10A). Although ABR responses improved by P21, DTX-treated
   mice continued to display elevated thresholds when compared to control
   mice (Figure [199]10B). Representative ABR waveforms produced in
   response to tone bursts at 8 kHz in P21 control mice demonstrate
   characteristic waveforms, similar to those produced by adult mice,
   while DTX-treated mice produced dampened and wider peaks (Figures
   [200]10E,F). By 1 month DTX-treated and control mice demonstrated
   similar ABR thresholds, with the exception of thresholds produced at
   the high frequencies (Figure [201]10C). To determine if OHC loss or
   dysfunction was present in DTX-treated mice we measured the distortion
   product otoacoustic emissions (DPOAEs) in the adult mice (Figure
   [202]10D). Nearly all frequencies tested, except a slight increase at
   32 kHz, showed no difference in DPOAE thresholds when comparing
   DTX-treated and control responses (Figure [203]10D).

Figure 10.

   [204]Figure 10
   [205]Open in a new tab

   Macrophage depletion impairs auditory function in DT-treated
   CD11b^DTR/EGFP mice. (A–C) Auditory brainstem response (ABR) thresholds
   from P15 (A), P21 (B) and 1 month (C) control (black) and DTX-treated
   (gray) CD11b^DTR/EGFP mice. ABRs were measured from 4.0 kHz to 42.5 kHz
   with sound pressure levels from 10 dB to 90 dB. Two-tailed, unpaired
   Student’s t-tests were used to compare thresholds for each group at
   each frequency (*p < 0.05; **p < 0.01; ***p < 0.001; n = 11–13 ears).
   Error bars represent SEM. (D) Distortion product otoacoustic emission
   (DPOAE) thresholds of 1 month control and DTX-treated CD11b^DTR/EGFP
   mice. DPOAEs were measured from 5.6 kHz to 42.5 kHz with sound pressure
   levels from 10 dB to 90 dB. Two-tailed, unpaired Student’s t-tests were
   used to compare thresholds for each group at each frequency (**p <
   0.01; n = 8–10 ears). Error bars represent SEM. (E,F) Representative
   ABR recordings from P21 control (C) and DTX-treated (D) CD11b^DTR/EGFP
   mice with 8 kHz pure tone stimulus. Arrows indicate the lowest ABR
   thresholds at this frequency. (G) Average wave I amplitudes for P21
   DTX-treated (gray, n = 4–7 ears) and control (black, n = 7–11 ears)
   CD11b^DTR/EGFP mice in response to 8.0 kHz stimulus. DTX-treated
   CD11b^DTR/EGFP mice displayed decreased wave I amplitude responses at
   90, 80, 70, 60 and 50 dB SPL. Unpaired Student’s t-test analyses were
   used to compare amplitudes for each group at each stimulus level tested
   (**p < 0.01; **p < 0.001). Error bars represent SEM. (H) Average wave I
   amplitudes at 4.0–40 kHz for stimuli presented at 90 dB SPL for P21
   control and DT treated (gray, n = 4–7 ears) and control (black, n = 11
   ears) CD11b^DTR/EGFP mice. Responses are decreased in DTX-treated
   CD11b^DTR/EGFP mice at all frequencies in comparison to control mice.
   Unpaired t-test analyses were used to compare thresholds for each group
   at each frequency (**p < 0.01; ***p < 0.001). Error bars represent SEM.
   (I) Average Wave I latencies for P21DTX-treated (gray, n = 3–8 ears)
   and control (black, n = 7–11 ears) CD11b^DTR/EGFP mice in response to
   8.0 kHz stimulus. DTX-treated CD11b^DTR/EGFP mice displayed longer
   latency responses at 90, 80, 70, 60 and 50 dB SPL. Unpaired Student’s
   t-test analyses were used to compare amplitudes for each group at each
   stimulus level tested (*p < 0.05). Error bars represent SEM.

   ABR wave I amplitude input/output function measurements allow the
   examination of suprathreshold activities of the auditory nerve. ABR
   wave I latencies give insight into functional integrity of the myelin
   in the auditory nerve. In DTX-treated mice, ABR wave I amplitudes
   measured at 8 kHz were decreased compared with controls when presented
   with stimuli at 90–50 dB SPL (Figure [206]10G). Wave I amplitudes give
   insight into the firing synchrony of the auditory nerve. Wave I
   amplitudes in response to 90 dB stimuli, measured at 4–40 kHz, were
   significantly lower in DTX-treated mice compared to vehicle-treated
   mice (Figure [207]10H). Additionally, increased wave I latencies were
   observed in DTX-treated mice at 8 kHz when 60–70 dB SPL stimuli were
   given (Figure [208]10I). No significant difference was seen in Wave I
   amplitudes or latencies measured at 8 kHz of 1 month DTX-treated and
   control mice when presented with stimuli at 90–40 dB (data not shown).
   These results demonstrate that transient macrophage-depletion impairs
   auditory nerve function. However, these impairments are temporary and
   hearing function between DTX-treated and control mice become
   comparable.

Discussion

   The refinement of auditory nerve fibers during development has been
   extensively studied. However, it remains unclear how glial cells, which
   ensheath SGNs and peripheral axons, are affected during the refinement
   period. In this study, we identify the temporal distribution patterns
   of cochlear glial cells during auditory nerve refinement. We
   demonstrate that cochlear macrophages eliminate supernumerary glial
   cells in the developing auditory nerve. We also show that macrophage
   depletion in the auditory nerve results in elevated glial cell numbers,
   abnormal myelin sheath formation, and diminished hearing function.
   These data reveal a critical role that cochlear macrophages play in
   glial cell regulation and auditory nerve maturation and hearing onset.

   Using immunohistochemical analyses, we found that postnatal glial cells
   undergo an increase in cell numbers between P3 and P7. Glial cell
   numbers significantly decrease after the first postnatal week, when
   mature auditory nerve circuitry is established. It is probable that
   extra glial cells, termed supernumerary glia in the context of this
   report, are temporarily produced to support processes important for
   proper nerve development. Glial cells, such as astrocytes and
   oligodendrocytes produce neurotrophic factors that are critical for the
   survival of neurons in the central nervous system (Purves et al.,
   [209]1988; Zhu et al., [210]2008). Glial cells from both the central
   and peripheral auditory nerve promote neuron survival and extension of
   SGN neurites in vitro (Hansen et al., [211]2001; Jeon et al.,
   [212]2011). It is possible that supernumerary glial cells are produced
   to ensure abundant production of trophic factors for the survival of
   developing neurons. Since glial cells have also been implicated as
   active participants in nerve refinement, it is also possible that
   supernumerary glial cells facilitate axonal and synaptic elimination
   during the auditory nerve refinement process. In mammalian
   neuromuscular junctions, glial cells facilitate synapse elimination and
   clear axonal debris during injury-induced nerve retraction (Bishop et
   al., [213]2004; Kang et al., [214]2014; Lee et al., [215]2016). Further
   studies are required to understand fully the function of supernumerary
   glial cells in auditory nerve refinement. It will be important to
   determine why glial cell numbers decrease following refinement and
   nerve maturation.

   Myelin production and integrity are essential for the proper function
   and survival of neurons. Complete myelination of SGNs and nerve fibers
   by glial cells is important for proper auditory signal conduction from
   the peripheral auditory nerve to central auditory processes (Kim et
   al., [216]2013). Mouse models with abnormal myelination in the inner
   ear, including Connexin29-deficient mice and Saposin B-deficient mice,
   have been previously described (Tang et al., [217]2006; Akil et al.,
   [218]2015). In the inner ear, the gap junction protein Connexin29 is
   highly expressed in myelinating glial cells that ensheath the somas and
   processes of SGNs. Connexin29-deficient mice demonstrate abnormal
   myelination of SGN somas, elevated ABR thresholds, and significantly
   delayed ABR wave I latencies (Tang et al., [219]2006). Saposin B is a
   glycoprotein responsible for lipid transfer in myelin. Satellite cells
   in the auditory nerve of Saposin B-deficient mice display abnormal
   inclusion bodies in their myelin sheaths, leading to SGN degeneration
   and progressive nerve fiber loss (Akil et al., [220]2015). These
   pathological alterations result in increased ABR thresholds and delayed
   wave I latencies (Akil et al., [221]2015).

   Our transcriptomic analysis detected increased expression of immune
   response genes during the period of nerve refinement, including H2-K1,
   H2-D1, C1q and C3aR. These results are consistent with a previous study
   (Lu et al., [222]2011), suggesting that immune signals play a role in
   auditory nerve refinement. Previous studies also have identified a role
   for complement and MHC class I signaling molecules in regulating
   microglia-mediated nerve pruning in the developing mammalian
   hippocampus, cerebellum, motor cortex, and optic nerve (Stevens et al.,
   [223]2007; Paolicelli et al., [224]2011; Berg et al., [225]2012;
   Schafer et al., [226]2012; Bialas and Stevens, [227]2013; Dixon-Salazar
   et al., [228]2014). Interestingly, C1q deficient and H2-K/H2-D1 double
   knockout mice do not exhibit altered auditory nerve fiber or synapse
   numbers (Calton et al., [229]2014). However, it remains to be
   determined whether these signaling molecules are involved in other
   components of auditory nerve development, such as glial cell
   maturation. It is possible that these molecules could play a role in
   glial cell maintenance. H2-K/H2-D1 double knockout mice demonstrate
   progressive hearing impairment, shown by elevated ABR thresholds,
   suggesting that these mice may possess morphologic abnormalities in the
   auditory nerve that remain to be identified (Calton et al., [230]2014).
   Further exploration of the differentially expressed genes identified in
   our study may provide insight into molecular cues or gene networks that
   regulate auditory nerve maturation.

   We found that macrophages engulf nerve fiber fragments during auditory
   nerve refinement. At the same time, macrophages seem to engulf glial
   cells. Similarly, several studies have shown that microglia or
   macrophages in the CNS regulate the number of synaptic outputs during
   development, as well as in adulthood, by engulfing synaptic elements
   that are misconnected or redundant (Paolicelli et al., [231]2011;
   Schafer et al., [232]2012; Zusso et al., [233]2012; Chen et al.,
   [234]2014; Shigemoto-Mogami et al., [235]2014). Our loss-of-function
   experiments found that activated macrophages are needed to eliminate
   supernumerary glial cells during refinement. Quantitative assessment of
   glial cells remaining in macrophage-depleted auditory nerves revealed a
   significant increase in glial cell numbers, when compared to controls.

   It remains to be determined why supernumerary glial cells are targeted
   by macrophages for elimination. Our analysis of the ultrastructure in
   P7 CBA mice revealed that macrophages engulf both myelinating and
   non-myelinating glial cells as they enwrap degenerating axons. However,
   it is unknown how targeted glial cells are marked for macrophage
   phagocytosis. Recent studies have identified several signals that act
   as “find me” and “eat me” signals for macrophage phagocytosis of
   apoptotic cells (Truman et al., [236]2008; Elliott et al., [237]2009).
   Identification of glial-related “eat me” signals will be important for
   further understanding the glia-macrophage relationship we have
   identified in the developing auditory nerve.

   Here, CD11b^DTR/EGFP mice treated with DTX at P4/5 demonstrated myelin
   abnormalities and functional impairments similar to the Connexin29 and
   Saposin B mouse models mentioned above. DTX-treated CD11b^DTR/EGFP mice
   presented with abnormally high glial cell numbers, the presence of
   myelin blebbing, abnormal myelination at the paranodes of the nodes of
   Ranvier, and supernumerary glial cell clustering around SGN axons.
   However, DTX-treated mice did not demonstrate alterations in neuron
   numbers or HC innervation and afferent synapse patterns. Our
   ultrastructural investigation of auditory nerves following
   macrophage-depletion confirmed the presence of supernumerary glial
   cells not directly enclosing SGNs or axons. In addition, pathological
   alterations of myelin sheaths in the auditory nerve suggest that
   macrophage depletion leads to abnormal glial cell function. Moreover,
   cochleae of DTX-treated CD11b^DTR/EGFP mice exhibited impaired
   suprathreshold function of the auditory nerve, indicating that proper
   macrophage activities are critical for auditory nerve maturation and
   normal hearing function. Further studies will be necessary to determine
   how elevation of glial cell numbers following macrophage depletion
   contributes to the pathophysiological changes in the auditory nerve.

   DTX-treated mice demonstrated auditory impairment at P15, after hearing
   function begins. DTX-treated mice continued to produce worsened ABR
   responses than control mice at P21. Additionally, DTX-treated mice
   demonstrated dampened wave I amplitudes and increased wave I latencies
   while DPOAE threshold responses remained comparable between DTX-treated
   mice and controls. By 1 month of age, control and DTX-treated animals
   presented similar ABR thresholds, with significant differences only
   seen in the higher frequencies tested, suggesting that DTX-treated mice
   eventually employed compensatory mechanisms for auditory nerve
   maturation.

   The stria vascularis maintains the homeostasis of the endolymph in the
   inner ear. Maintenance of the stria vascularis is essential for the
   production of the endocochlear potential, which is required for hearing
   function (Salt et al., [238]1987; Takeuchi et al., [239]2000; Juhn et
   al., [240]2001). Within the stria vascularis are resident macrophages
   referred to as perivascular-resident macrophage-like melanocytes. These
   cells are positive for macrophage-specific markers including F4/80,
   CD68 and CD11b (Zhang et al., [241]2012). Previous studies demonstrated
   that these macrophage-like cells contribute to the integrity of the
   stria vascularis in adult mice (Zhang et al., [242]2012). Additionally,
   disruption of CD11b^+ cells in the cochlear LW causes leakage from
   capillaries of the stria compartment, leading to a significant
   reduction in endocochlear potential and auditory function decline
   (Zhang et al., [243]2012, [244]2013). In the current study, strial
   edema was seen in the LWs of P11 DTX-treated mice, suggesting that the
   perivascular-resident macrophage-like melanocytes may be affected by
   DTX treatment. However, edema was not found in the stria compartment of
   P16 DTX-treated mice, suggesting that the LW is capable of recovering
   from macrophage depletion. Since proper generation of endocochlear
   potential is critical for maintenance of auditory function, it is
   possible that the changes of LW function also contribute to the
   temporary hearing impairments seen in the DTX-treated mice. It will be
   important for future studies to investigate the role of macrophages in
   LW development and contributions to endocochlear potential of postnatal
   mice.

   Recently, our studies demonstrated that glial cell numbers are
   increased after auditory nerve injury and that auditory glial cells
   have the potential to differentiate into neural progenitor cells
   following acute nerve injury (Lang et al., [245]2011, [246]2015).
   Interestingly, macrophage numbers are increased following auditory
   injury and are required for SGN protection and the maintenance of SGN
   numbers after cochlear injury in the adult cochlea (Hirose et al.,
   [247]2005; Lang et al., [248]2006, [249]2016; Okano et al., [250]2008;
   Sato et al., [251]2010; Kaur et al., [252]2015). Investigation of glial
   cell refinement and glia-macrophage relationships during development
   will lead to better understanding of the auditory nerve
   microenvironment. This will be vital for identifying novel strategies
   that may promote auditory nerve repair or regeneration.

Author Contributions

   LNB, YX, JLB and HL designed the research; LNB, YX, KVN, JZ, NMS, and
   JLB performed the research; LNB, YX, KVN, JLB, CHP and MCB analyzed the
   data; LNB, JLB and HL wrote the article.

Conflict of Interest Statement

   The authors declare that the research was conducted in the absence of
   any commercial or financial relationships that could be construed as a
   potential conflict of interest.

Acknowledgments