We first employed immunohistochemistry to study IHCs of 3-week-old RIBEYE knock-out mice (RBE^KO/KO), in which the unique A-domain exon of RIBEYE was deleted by Cre-mediated excision (described in Maxeiner et al., 2016). Next to the A-domain, RIBEYE contains a B-domain that is largely identical to the transcription factor CtBP2, which is spared by the genetic manipulation and used as a target in immunohistochemistry of ribbons and nuclei (Figure 1A,B; Khimich et al., 2005). Synaptic ribbons of IHC afferent synapses were identified as presynaptic RIBEYE/CtBP2-immunofluorescent spots in wild-type (Figure 1B, RBE^WT/WT) and heterozygous (Figure 1—figure supplement 1, RBE^WT/KO) mice. Their number per IHC did not change in the heterozygous condition (15.5 ± 0.7, S.D. = 1.58; n = 50 cells, N = 4 for RBE^WT/KO vs. 15.7 ± 1.1, S.D. = 2.19; n = 39 cells, N = 3 for RBE^WT/WT at P21), while their intensity was significantly reduced (in arbitrary units: 3.4 ± 0.7, S.D. = 1.78; n = 600 spots for 40 cells, N = 3 in RBE^WT/KO vs. 5.1 ± 1.1, S.D. = 2.23; n = 411 spots for 29 IHCs, N = 3 in RBE^WT/WT; p<0.0001, Mann-Whitney-Wilcoxon test; Figure 1—figure supplement 1A,B). RBE^KO/KO IHCs lacked synaptic RIBEYE/CtBP2 immunofluorescence spots (Figure 1B), while immunolabeling of nuclear CtBP2 remained present, corroborating previous findings in the retina (Maxeiner et al., 2016). The number of afferent synapses per IHC was determined by the count of postsynaptic densities (PSDs) identified as PSD-95 immunofluorescent spots (Figure 1C,D,E) and was unchanged when RIBEYE was removed (13.7 ± 0.8, S.D. = 2.04; n = 56 cells, N = 4 in RBE^KO/KO vs. 12.9 ± 0.6, S.D. = 2.13; n = 55 cells, N = 5 in RBE^WT/WT). Bassoon (Figure 1C) and RIM2 (Figure 1D), both presynaptic scaffold proteins (Khimich et al., 2005;Jung et al., 2015a), remained present at the ribbonless afferent synapses of RBE^KO/KO IHCs (marked by PSD-95). The scaffold protein piccolino, the short isoform of piccolo (Regus-Leidig et al., 2013) that is present in cochlear and retinal ribbons (Khimich et al., 2005; tom Dieck et al., 2005; Regus-Leidig et al., 2013), was absent from afferent synapses of RBE^KO/KO IHCs. However, piccolo immunofluorescence was present in the vicinity of afferent synapses likely marking the long form piccolo at the efferent presynaptic AZs (Figure 1E, see schematic in Figure 1A). The PSD areas were calculated by fitting a 2-dimensional Gaussian function to each PSD-95 immunofluorescent spot, revealing a significant increase in the RBE^KO/KO condition (2.82 ± 0.09 µm², S.D. = 1.25; n = 178 spots, N = 3 vs. 1.74 ± 0.05 µm², S.D. = 0.58; n = 163 spots, N = 3 in RBE^WT/WT IHCs; p<0.0001, Mann-Whitney-Wilcoxon; Figure 1F).

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In order to study the effects of RIBEYE deletion on the ultrastructure of afferent IHC synapses, we performed transmission electron microscopy on random sections and electron tomography. Random ultrathin (70–75 nm) sections prepared from P21 mice (two animals per genotype) after aldehyde fixation and conventional embedding procedures showed that IHCs from RBE^KO/KO mice completely lack synaptic ribbons, while RBE^WT/WT and heterozygous RBE^WT/KO typically display one ribbon per AZ (Figure 2A–C). Interestingly, ribbons of RBE^WT/KO IHCs were smaller in height, width and area compared to RBE^WT/WT IHC synaptic ribbons (Figure 2—figure supplement 1A–C; ribbon height: 118.32 ± 3.17 nm, S.D. = 31.84 nm; n = 101 ribbons, N = 2 for RBE^WT/KO vs. 197.09 ± 4.36 nm, S.D. = 44.93 nm; n = 106 ribbons, N = 2 for RBE^WT/WT; ribbon width: 119.80 ± 6.23 nm, S.D. = 62.27 nm for RBE^WT/KO vs. 168.34 ± 6.83 nm, S.D. = 70.27 nm for RBE^WT/WT; ribbon area: 11.5e3 ± 6.2e2 nm², S.D. = 6.3e3 nm² for RBE^WT/KO vs. 25.4e3 ±1.1e2 nm², S.D. = 1.1e3 nm² for RBE^WT/WT; p<0.0001, Mann-Whitney-Wilcoxon test for all) agreeing with the significantly reduced ribbon immunofluorescence intensity in the RBE^WT/KO condition (see above and Figure 1—figure supplement 1A–B).

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Random sections of synaptic contacts of RBE^KO/KO mice (Figure 2C) often showed more than one presynaptic density (PD), each associated with a cluster of synaptic vesicles (henceforth considered individual AZs). The multiple AZs typically faced one continuous PSD, which is different from the synapses of immature IHC synapses that show multiple appositions of pre- and postsynaptic densities (Sendin et al., 2007; Wong et al., 2014). Moreover, we found more than one PD per synaptic contact in IHCs of older RBE^KO/KO mice (Figure 2E,F; 6 weeks and 8 months, respectively), arguing against a delayed synaptic maturation to be the cause of the phenotype. Sections from tangential cuts of the synapse (Figure 2D), reconstructions from serial ultrathin sections (Figure 2G,G’) and quantifications of random sections (Figure 2H) corroborated the notion of multiple small ribbonless AZs at the synaptic contacts of RBE^KO/KO IHCs. Analysis based on serial 3D reconstructions of synaptic contacts of RBE^KO/KO IHCs from P21 animals showed on average 1.92 ± 0.34 PDs (S.D. = 1.16; n = 17 serial 3D reconstructions, N = 2) and 20.58 ± 2.98 total SVs per contact, S.D. = 10.34 (Figure 2I). The lateral extent of the individual PDs, determined in random sections, was comparable between RBE^KO/KO and RBE^WT/WT synapses (129.89 ± 2.53 nm, S.D. = 26.26 nm; n = 108 PDs, N = 2 for RBE^KO/KO vs. 129.35 ± 4.89 nm, S.D. = 50.86 nm; n = 108 PDs, N = 2 for RBE^WT/WT; p=0.92, NPMC test), while that of RBE^WT/KO was enlarged (Figure 2M; 157.64 ± 7.19 nm, S.D. = 72.24 nm; n = 101 PDs, N = 2; p=0.0004 for comparison to RBE^WT/WT, NPMC test). PSDs tended to be increased in length at RBE^KO/KO synapses compared to RBE^WT/WT PSDs and were significantly larger than RBE^WT/KO PSDs (Figure 2N; 623.77 ± 26.70 nm, S.D. = 264.33 nm; n = 98 PSDs, N = 2 for RBE^KO/KO vs. 555.91 ± 22.24 nm, S.D. = 236.42 nm; n = 113 PSDs, N = 2 for RBE^WT/WT vs. 521.34 ± 24.20 nm, S.D. = 242.03 nm; n = 100 PSDs, N = 2 for RBE^WT/KO; p=0.01 for RBE^KO/KO vs. RBE^WT/KO, Tukey’s test), which is consistent with the greater area of PSD-95 immunofluorescent spots in the knock-out condition (Figure 1E).

In the following, we characterized the populations of presynaptic SVs in random sections of vertically-cut IHC synapses. We counted membrane-proximal SVs (MP-SVs, ≤25 nm distance between SV membrane and plasma membrane, laterally within 80 nm of the PD, yellow in Figure 2J–L) as well as ribbon-associated SVs (RA-SVs, first layer of SVs around the ribbon within 80 nm, green in Figure 2J,K) or ‘PD-associated’ SVs (PDA-SVs, ribbonless AZs: SVs within 80 nm distance of the PD and not falling into the MP-SV pool (see above), green in Figure 2J,L). We found both MP-SVs (Figure 2O; 1.92 ± 0.09, S.D. = 0.93; n = 108 AZs, N = 2 for RBE^KO/KO vs. 2.99 ± 0.12, S.D. = 1.18; n = 101 AZs, N = 2 for RBE^WT/KO vs. 2.77 ± 0.12, S.D. = 1.18; n = 106 AZs, N = 2 for RBE^WT/WT; p<0.0001 for RBE^KO/KO vs. RBE^WT/WT, NPMC test) and PDA-SVs (Figure 2P; 4.12 ± 0.15, S.D. = 1.50; n = 108 AZs, N = 2 for RBE^KO/KO vs. 10.09 ± 0.27, S.D. = 2.75; n = 106 AZs, N = 2 for RBE^WT/WT; p<0.0001, Tukey’s test) of the individual ribbonless IHC AZs of RBE^KO/KO mice to be significantly fewer than the corresponding number of MP-SVs and RA-SVs counted at RBE^WT/WT AZs. The fraction of PDA-SVs relative to the total number of SVs at RBE^KO/KO AZs was less than that of RA-SVs at RBE^WT/WT AZs (Figure 2Q; 0.67 ± 0.02, S.D. = 0.16; n = 108 AZs, N = 2 for RBE^KO/KO vs. 0.78 ± 0.01, S.D. = 0.08; n = 106 AZs, N = 2 for RBE^WT/WT; p<0.0001, NPMC test). Consequently, we observed an increase in the fraction of MP-SVs at RBE^KO/KO AZs (Figure 2Q; 0.33 ± 0.02, S.D. = 0.16; n = 108 AZs, N = 2 for RBE^KO/KO vs. 0.22 ± 0.01, S.D. = 0.08; n = 106 AZs, N = 2 for RBE^WT/WT; p<0.0001, NPMC test). In line with the decreased ribbon size of RBE^WT/KO AZs, we found a reduced number of RA-SVs, indicating a hypomorphic phenotype upon the loss of one allele of the RIBEYE gene. The SV diameter was unchanged for all three genotypes when jointly considering SVs of all categories in random sections (Figure 2—figure supplement 1D; 39.59 ± 0.21 nm, S.D. = 5.37 nm; n = 108 AZs, N = 2 for RBE^KO/KO vs. 40.53 ± 0.14 nm, S.D. = 4.44 nm; n = 101 AZs, N = 2 for RBE^WT/KO vs. 41.80 ± 0.13 nm, S.D. = 4.79 nm; n = 106 AZs, N = 2 for RBE^WT/WT; p=0.30, NPMC test). However, we found a subtle but significant SV-diameter reduction in RBE^KO/KO and RBE^WT/KO for MP-SVs (Figure 2—figure supplement 1E; 39.29 ± 0.34 nm, S.D. = 4.82 nm; n = 108 AZs, N = 2 for RBE^KO/KO vs. 41.79 ± 0.26 nm, S.D. = 4.53 nm; n = 106 AZs, N = 2 for RBE^WT/WT; p<0.0001, NPMC test and 40.29 ± 0.25 nm, S.D. = 4.40 nm; n = 101 AZs, N = 2 for RBE^WT/KO vs. RBE^KO/KO; p=0.03, NPMC test) and for RA-/PDA-SVs (Figure 2—figure supplement 1F; 39.72 ± 0.27 nm, S.D. = 5.61 nm; n = 108 AZs, N = 2 for RBE^KO/KO vs. 41.81 ± 0.15 nm, S.D. = 4.86 nm; n = 106 AZs, N = 2 for RBE^WT/WT; p<0.0001, NPMC test and RBE^KO/KO vs. 40.63 ± 0.17 nm, S.D. = 4.45 nm; n = 101 AZs, N = 2 for RBE^WT/KO; p=0.003, NPMC test and RBE^WT/KO vs. RBE^WT/WT; p=0.02, NPMC test).

Next, to capture the synapses in a near-to-native state and to evaluate vesicle tethering, we performed electron tomography on 250 nm thick sections that were prepared with high-pressure freezing and freeze-substitution (HPF/FS) of organs of Corti from P21 mice (Figure 3). Tomography confirmed the absence of synaptic ribbons and the presence of multiple AZs per contact, each with a clear PD (Figure 3B,D,F). However, we note that the 250 nm thick sections did typically not fully cover the synaptic contact, which leads to an underestimation for the total number of SVs particularly for the spatially extended RBE^KO/KO synapses. The PDs appeared roundish in the RBE^KO/KO with MP-SVs closely arranged around the PD as found at the more elongated RBE^WT/WT AZs (Figure 3C,D). For the tomograms, we followed the definition of MP-SV pool according to the 2D-random sections (Figure 2), but in addition we measured the MP-SVs also in a maximum distance of 50 nm from the AZ membrane and ≤100 nm from the PD (Figure 3—figure supplement 1 and supplementary file 1). This was motivated by the presence of long tethers connecting SV and AZ membrane and was previously introduced (Jung et al., 2015a). Further, we distinguished between tethered and non-tethered SVs (Figure 3G, Figure 3—figure supplement 1A). There, we focused our analysis on tethers to the ribbon/PD, plasma membrane and those interconnecting two adjacent SVs (Figure 3H,I,M,N). We found a significant reduction in the number of MP-SVs per AZ in RBE^KO/KO IHCs (Figure 3J; RBE^KO/KO = 6.30 ± 0.86, S.D. = 2.87 MP-SVs; n = 11 AZs, N = 3 vs. RBE^WT/WT = 8.70 ± 0.82, S.D. = 2.45 MP-SVs; n = 9 AZs, N = 3; p=0.04, Mann-Whitney-Wilcoxon test; significant also by the second analysis method: Figure 3—figure supplement 1B, supplementary file 1), while the fraction of tethered MP-SVs (No. of tethered MP-SVs/No. of all MP-SVs) was not significantly altered (Figure 3K; RBE^KO/KO = 0.75 ± 0.07, S.D. = 0.24; n = 11 AZs, N = 3 vs. RBE^WT/WT = 0.65 ± 0.06, S.D. = 0.18; n = 9 AZs, N = 3; p=0.30, t-test; Figure 3—figure supplement 1C, supplementary file 1). The majority of the MP-SVs were tethered via a single tether in both RBE^WT/WT and RBE^KO/KO IHCs. The fraction of MP-SVs with multiple (≥2) tethers was significantly larger in RBE^KO/KO IHCs (Figure 3K; single-tethered MP-SVs: RBE^KO/KO = 0.55 ± 0.06, S.D. = 0.19; n = 11 AZs, N = 3 vs. RBE^WT/WT = 0.61 ± 0.06, S.D. = 0.17; n = 9 AZs, N = 3; p=0.81; multiple-tethered MP-SVs: RBE^KO/KO = 0.20 ± 0.05, S.D. = 0.15; n = 11 AZs, N = 3 vs. RBE^WT/WT = 0.04 ± 0.02, S.D. = 0.05; n = 9 AZs, N = 3; p=0.01, Tukey’s test; Figure 3—figure supplement 1C, supplementary file 1). Further, and in line with analysis of random sections, the number of PDA-SVs per RBE^KO/KO AZ was smaller than that of RA-SVs at RBE^WT/WT AZs (Figure 3O; RBE^KO/KO: 9.30 ± 1.13, S.D. = 3.74 PDA-SVs; n = 11 AZs, N = 3 vs. RBE^WT/WT: 30.33 ± 3.00, S.D. = 9.01 RA-SVs; n = 9 AZs, N = 3; p<0.0001, Mann-Whitney-Wilcoxon test; Figure 3—figure supplement 1, supplementary file 1). However, the fraction of PDA-SVs tethered to the PD was not different from that of RA-SVs tethered to the ribbon (Figure 3P; RBE^KO/KO: 0.80 ± 0.06, S.D. = 0.19 tethered PDA-SV fraction; n = 11 AZs, N = 3 vs. RBE^WT/WT: 0.70 ± 0.06, S.D. = 0.17 tethered RA-SV fraction; n = 9 AZs, N = 3; p=0.12, t-test; Figure 3—figure supplement 1F, supplementary file 1).

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Finally, tomography indicated unchanged SV diameters at RBE^KO/KO AZs (Figure 3L,Q; MP-SV diameter: 50.17 ± 0.90 nm, S.D. = 2.95 nm; n = 11 PDs, N = 3 for RBE^KO/KO vs. 47.81 ± 0.60 nm, S.D. = 1.70; n = 9 ribbons, N = 3 for RBE^WT/WT; p=0.06, Mann-Whitney-Wilcoxon test, Figure 3—figure supplement 1D, supplementary file 1; RA/RA-SV diameter: 49.71 ± 0.83 nm, S.D. = 2.75; n = 11 PDs, N = 3 for RBE^KO/KO vs. 49.80 ± 0.78 nm, S.D. = 2.35; n = 9 ribbons, N = 3 for RBE^WT/WT; p=0.71, Mann-Whitney-Wilcoxon test; Figure 3—figure supplement 1G, supplementary file 1). We presume that differences in the comparison of RBE^KO/KO and RBE^WT/WT between the random section and electron tomography analysis primarily reflects the larger number of AZ analyzed by the former approach.

We then used confocal and stimulated emission depletion (STED) super-resolution immunofluorescence microscopy in order to study the abundance and spatial organization of presynaptic Ca_V1.3 Ca²⁺-channels (Neef et al., 2018), which contribute more than 90% of the voltage-gated Ca²⁺-influx into IHCs (Platzer et al., 2000; Brandt et al., 2003; Dou et al., 2004). Organs of Corti from 3-week-old RBE^KO/KO and RBE^WT/WT mice were processed in parallel for immunohistochemistry and imaging. Ca_V1.3 Ca²⁺-channels remained clustered at RBE^KO/KO AZs and were identified as Ca_V1.3 labeling juxtaposed to PSD-95 immunofluorescent spots (Figure 4A). In order to analyze the spatial organization of synaptic Ca²⁺-channels, we performed 3-color, 2D-STED immunofluorescence imaging for Ca_V1.3, bassoon (as a PD-marker), and PSD-95. While more than 80% of the RBE^WT/WT synapses showed the typical stripe-like co-alignment of Ca_V1.3 and bassoon immunofluorescence (Neef et al., 2018), imaging of RBE^KO/KO synapses indicated a high prevalence (over 70%) of smaller, rounder and often several Ca²⁺-channel clusters and PDs per synaptic contact (Figure 4B,C). We then quantified stripe-like clusters by measuring their long and short axis using 2D Gaussian fits and found no differences between RBE^KO/KO and RBE^WT/WT AZs (Figure 4D). Finally, we quantified the number of Ca_V1.3-immunofluorescent structures per contact (as indicated by PSD-95 immunofluorescence). While more than 80% of RBE^WT/WT synapses displayed a single cluster, over 60% of the RBE^KO/KO synapses contained two or more Ca²⁺-channel clusters (Figure 4E). Hence, the average number of Ca_V1.3-immunofluorescent structures was significantly higher at RBE^KO/KO synapses compared to RBE^WT/WT (2.06 ± 0.09, S.D. = 1.16; n = 178 spots, N = 3 vs. 1.16 ± 0.03, S.D. = 0.38; n = 183 spots, N = 2; p<0.0001, Mann-Whitney-Wilcoxon test) and we likely underestimated this difference due to the low resolution of 2D-STED in the z-axis. In summary, our results indicate that RIBEYE-disruption transforms the single ribbon-type AZ into a complex presynaptic organization with multiple conventional-like AZs facing the postsynaptic bouton.

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Next, we combined whole-cell patch-clamp with confocal Ca²⁺-imaging of IHCs to study Ca²⁺-influx at the whole IHC and single synapse levels using 5 mM [Ca²⁺]_e to augment the signal to noise. Using step-depolarizations in conditions that isolated the Ca²⁺-current (see Materials and methods), we probed the amplitude and voltage-dependence of IHC Ca²⁺-influx (Figure 5A). The amplitude of Ca²⁺-influx (Figure 5A_i; for Ca²⁺-current density, see Figure 5A_ii) was unaltered in RBE^KO/KO IHCs (-151 ± 12.9 pA, S.D. = 59 pA; n = 21 IHCs, N = 8 in RBE^KO/KO vs. -161 ± 15.4 pA, S.D. = 71 pA; n = 21 IHCs, N = 9 in RBE^WT/WT; p=0.62, t-test), in agreement with findings in retinal bipolar neurons (Maxeiner et al., 2016) but in contrast to our previous findings in ribbon-deficient IHCs of bassoon mutant mice (Khimich et al., 2005; Frank et al., 2010; Jing et al., 2013). Kinetics of Ca²⁺-channel activation were unchanged (Figure 5B), whereas inactivation kinetics were slightly faster in the RBE^KO/KO IHCs (smaller residual Ca²⁺-current at 200 ms of depolarization (normalized to the peak current): 0.82 ± 0.007, S.D. = 0.02; n = 10 IHCs, N = 5 for RBE^KO/KO vs. 0.85 ± 0.01, S.D. = 0.04; n = 11 IHCs, N = 7, in the RBE^WT/WT condition; p=0.017, Mann-Whitney-Wilcoxon test; Figure 5D). When analyzing the voltage-dependence of Ca²⁺-channel activation (Figure 5C), we found a small (2 mV) but significant depolarizing shift of the potential of half-maximal Ca²⁺-channel activation, V_h (Figure 5C_i, −22.96 ± 0.43 mV, S.D. = 2.39 mV; n = 21 IHCs, N = 8 in RBE^KO/KO vs. −25.04 ± 0.65 mV, S.D. = 2.98 mV; n = 21 IHCs, N = 9 in RBE^WT/WT; p=0.017, t-test). When analyzed in a smaller data set recorded in 2 mM [Ca²⁺]_e the depolarized V_h-shift did not reach statistical significance (data not shown). The average voltage-sensitivity of activation (slope factor k) was not altered (5 mM [Ca²⁺]_e: p=0.67, t-test, Figure 5C_ii). Together, this suggests a RIBEYE/ribbons-mediated regulation of IHC Ca²⁺-channels affecting their voltage-range of operation as well as their inactivation kinetics.

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We then used the low-affinity Ca²⁺-indicator dye Fluo-4FF (800 µM) to study Ca²⁺-influx at individual IHC AZs (Frank et al., 2009) using a spinning-disk confocal microscope that allows rapid registering and recording of the majority of the IHC synapses (Figure 6A, Ohn et al., 2016). We chose conditions in which the Ca²⁺-indicator fluorescence approximates synaptic Ca²⁺-influx (Frank et al., 2009; Ohn et al., 2016) and henceforth refer to synaptic Ca²⁺-influx when describing observations based on hotspots of Ca²⁺-indicator fluorescence at the basolateral IHC membrane. Prior to analysis of synaptic Ca²⁺-influx, we imaged fluorescently-conjugated CtBP2-binding peptide (Zenisek et al., 2004), which bound to the ribbon-occupied AZs in RBE^WT/WT IHCs while it only caused nuclear and diffuse cytosolic fluorescence in the ribbonless RBE^KO/KO IHCs (Figure 6B). We then employed ramp-depolarizations to assess amplitude and voltage-dependence of Ca²⁺-influx at the synapses located in the subnuclear, basal part of the IHCs (Figure 6A). We found comparable maximal amplitudes of the baseline-normalized fluorescence change (ΔF/F₀, 0.88 ± 0.08, S.D. = 0.66; n = 61 AZs in 15 IHCs, N = 7 for RBE^KO/KO vs. 0.85 ± 0.08, S.D. = 0.68; n = 78 AZs in 15 IHCs, N = 8 for RBE^WT/WT; p=0.20, Mann-Whitney-Wilcoxon test; Figure 6C). This is compatible with an unaltered number of synaptic Ca²⁺-channels at AZs of RBE^KO/KO IHCs and consistent with our observations of normal whole-cell Ca²⁺-current amplitudes. As previously reported (Frank et al., 2009; Ohn et al., 2016), there was a substantial variation of the maximal ΔF/F₀ among the AZs, which was also comparable between AZs of both genotypes (c.v. = 0.75 for RBE^KO/KO vs. c.v. = 0.80 for RBE^WT/WT).

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Next, we analyzed the voltage-dependence of activation for the synaptic Ca²⁺-influx as previously described (Ohn et al., 2016). Analysis of fractional activation revealed a depolarized shift in V_h by on average 5 mV in RBE^KO/KO IHCs (−22.76 ± 1.25 mV, S.D. = 9.26 mV; n = 55 AZs in 15 IHCs, N = 7 for RBE^KO/KO vs. −27.37 ± 0.90 mV, S.D. = 7.48 mV; n = 68 AZs in 15 IHCs, N = 8 for RBE ^WT/WT; p=0.0029, t-test; Figure 6D,Di), while the slope factor of voltage-dependent activation was unaltered (p=0.42, t-test, Figure 6D_ii). Such a shift in the operating range of synaptic Ca²⁺-influx is expected to alter spontaneous and sound-evoked transmitter release (see below and Ohn et al., 2016). Finally, we studied the spatial extent of the synaptic Ca²⁺-signals and estimated Full Width Half Maximum (FWHM) by fitting 2D Gaussian functions to the hotspots of Ca²⁺-indicator fluorescence and found a greater spread of Ca²⁺-signals at RBE^KO/KO AZs (Figure 6E,F; long axis (L.A.) = 1317 ± 49 nm, S.D. = 384 nm, short axis (S.A.) = 906 ± 36 nm, S.D. = 284 nm; n = 61 AZs in 15 IHCs, N = 7 vs. L.A. = 1083 ± 33 nm, S.D. = 283 nm; (p=0.00016, t-test), S.A. = 793 ± 27 nm, S.D. = 233 nm, (p=0.0029, t-test); n = 74 AZs in 15 IHCs, N = 8 for RBE^WT/WT). This larger spread of the presynaptic Ca²⁺-signals is in agreement with the presence of several Ca_V1.3-immunofluorescent clusters at RBE^KO/KO synapses. In order to exclude lower IHC Ca²⁺-buffering to contribute to the observed larger spread of presynaptic Ca²⁺-signals, we performed semi-quantitative immunofluorescence analysis for the three major cytosolic Ca²⁺-buffers, the EF-hand Ca²⁺-binding proteins parvalbumin-α, calretinin and calbindin-28k (Pangršič et al., 2015). We did not find any significant differences in their immunofluorescence intensity between IHCs of both genotypes (in arbitrary units, parvalbumin intensity: 2.24 ± 0.15, S.D. = 1.04 for RBE^KO/KO vs. 1.88 ± 0.15, S.D = 1.01 for RBE^WT/WT, p=0.08; calbindin intensity: 0.82 ± 0.06, S.D. = 0.43 for RBE^KO/KO vs.0.95 ± 0.07, S.D. = 0.49 for RBE^WT/WT, p=0.23; calretinin intensity: 0.91 ± 0.04, S.D. = 0.26 for RBE^KO/KO vs. 0.82 ± 0.04, S.D. = 0.28 for RBE^WT/WT, p=0.09; n = 49 cells and N = 4 for both conditions, Mann-Whitney-Wilcoxon test for all; Figure 6—figure supplement 1).

The ribbon has been proposed to play a crucial role in the exocytosis of SVs at the IHC AZ (Khimich et al., 2005). Therefore, we monitored stimulated exocytosis of SVs with perforated-patch whole-cell recordings of exocytic membrane capacitance changes (ΔC_m). Using IHCs from 2/3-week-old RBE^WT/WT and RBE^KO/KO mice, we found that ΔC_m in response to step-depolarizations to the potential that elicits maximal Ca²⁺-influx (−14 mV) were not different between IHCs with or without ribbons. Both, fast exocytosis elicited by depolarizations of up to 20 ms, attributed to the fusion of the readily releasable pool of SVs (RRP, Moser and Beutner, 2000), and longer stimuli, thought to reflect sustained exocytosis, ongoing SV replenishment and fusion, were unaltered in RBE^KO/KO IHCs (Figure 7A,B,C). On average, ΔC_m induced by 20 ms long maximal Ca²⁺-influx was 16.70 ± 1.67 fF (S.D. = 5.80 fF; n = 12 cells, N = 7) for RBE^KO/KO compared to 15.22 ± 0.98 fF (S.D. = 3.26 fF; n = 11 cells, N = 8) for RBE^WT/WT. Exocytic ΔC_m elicited by 200 ms long maximal Ca²⁺-influx (same IHCs as for 20 ms), on average, amounted to 62.09 ± 5.40 fF (S.D. = 18.70 fF) for RBE^KO/KO versus 63.28 ± 6.64 fF (S.D. = 22.04 fF) for RBE^WT/WT.

Figure 7

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Moreover, trains of 20 step-depolarizations to −17 mV of 20 ms pulse duration did not reveal impaired exocytosis in RBE^KO/KO IHCs, even when the inter-stimulus interval time was as short as 160 ms (Figure 7D; n = 11 cells, N = 5 for RBE^WT/WT and n = 13 cells, N = 8 for RBE^KO/KO). We further explored RRP recovery from partial depletion using a paired-pulse protocol (two strong 20 ms depolarizations to −14 mV separated by 50, 110, 260 and 510 ms inter-pulse intervals; Figure 7E,F). RRP recovery, estimated as the ΔC_m ratio of the second and the first pulse, was not altered in RBE^KO/KO IHCs at least when probing RRP exocytosis with maximal Ca²⁺-influx from a hyperpolarized resting potential (Figure 7F). These data are in strong contrast to our previous findings in IHCs of bassoon mutant mice, which we had equivalently analyzed. There, the loss of synaptic ribbons, combined with a loss of functional bassoon resulted in profound deficits in exocytosis (Khimich et al., 2005; Frank et al., 2010; Jing et al., 2013).

Given the finding of a small depolarized shift in the operating range of Ca²⁺-channels in RBE^KO/KO IHCs (Figure 6D), we also probed the voltage-dependence of ΔC_m elicited by 100 ms step-depolarizations (Figure 7G,H). In agreement with the results obtained at maximal Ca²⁺-influx, we did not find significant differences in ΔC_m for stronger depolarizations (e.g. pulses to −39 mV elicited an average ΔC_m of 20.67 ± 7.46 fF, S.D. = 23.58 fF, n_min = 10 IHCs, N = 9 for RBE^KO/KO vs. 24.12 ± 4.04 fF, S.D. = 13.98 fF, n_min = 10 IHCs, N = 9 for RBE^WT/WT; p=0.20; Mann-Whitney-Wilcoxon test). However, for weaker depolarizations in the range of physiological receptor potentials (Russell and Sellick, 1983), we observed a subtle but significant reduction in exocytosis for RBE^KO/KO IHCs (Figure 7H, p=0.0115, p=0.0295 and p=0.1321 for −45, –43 and −41 mV; without definitive outliers as determined by Graphpad Prism: p=0.0017, p=0.0042 and p=0.0489, respectively; Mann-Whitney-Wilcoxon test for all). For instance, depolarization to −45 mV elicited a ΔC_m of 4.79 ± 2.26 fF for RBE^KO/KO (S.D. = 7.14 fF; n_min = 10 cells, N = 9) compared to 9.85 ± 1.60 fF for RBE^WT/WT (S.D. = 5.05 fF; n_min = 10 cells, N = 8). The Ca²⁺-current integral (Ca²⁺-charge, Q_Ca), as well, tended to be reduced for RBE^KO/KO IHC at these mild depolarizations, which, however, did not reach statistical significance (e.g. Q_Ca for −45 mV: 3.90 ± 0.49 pC, S.D. = 1.54 pC for RBE^KO/KO vs. 5.15 ± 0.54 pC, S.D. = 1.72 pC for RBE^WT/WT; p=0.1053; t-test). In summary, we found exocytosis to be unaltered for strong depolarizations but mildly decreased for more physiological stimuli in RBE^KO/KO IHCs, which is in line with the findings of the companion paper by Becker et al..

Next, we studied sound encoding in RBE^KO/KO mice in vivo. First, we recorded auditory brainstem responses (ABR) and found a significant reduction in the amplitude of wave I that reflects the SGN compound action potential (1.14 ± 0.13 µV, S.D. = 0.38 µV, N = 10 for RBE^KO/KO vs. 3.30 ± 0.51 µV, S.D. = 1.54 µV, N = 10 for RBE^WT/WT, p=0.0007, NPMC test). This indicates less synchronous SGN activation in the absence of synaptic ribbons (Figure 8A,B). The subsequent ABR waves (Figure 8—figure supplement 1) were normal in amplitude (waves II, IV and V, while wave III was reduced) indicating a degree of central auditory compensation for the sound encoding deficit, for example via coincidence detection of converging SGN input in the cochlear nucleus (Joris et al., 1994; Strenzke et al., 2009). We found a non-significant trend of ABR threshold to be increased across all frequencies in RBE^KO/KO mice (approximately 10 dB across all frequencies, Figure 8C; refer to the companion paper Becker et al. showing significantly increased ABR-thresholds based on a larger sample, N = 28 RBE^KO/KO mice vs. 22 RBE^WT/WT mice). Cochlear amplification, probed by recordings of distortion product otoacoustic emissions (DPOAE, Figure 8D), was intact in RBE^KO/KO mice. Additionally, RBE^WT/KO mice showed no significant changes in ABR wave I amplitudes and ABR thresholds (Figure 8), suggesting that the subtle morphological differences observed for afferent synapses of RBE^WT/KO IHCs by electron and confocal-immunofluorescence microscopy did not turn into a deficit of sound coding measurable by ABR recordings. The wave I amplitude reduction and ABR threshold elevation were much less pronounced than in bassoon mutant mice (Khimich et al., 2005; Buran et al., 2010; Jing et al., 2013).

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We then turned to in vivo extracellular recordings from single auditory neurons by targeting glass microelectrodes to where the auditory nerve enters the anteroventral cochlear nucleus (AVCN) in the brainstem (Taberner and Liberman, 2005; Jing et al., 2013). ‘Putative’ SGNs (hereafter dubbed SGN for simplicity) were identified based on the depth of electrode position and their firing response to pure-tone stimulation (primary-like peristimulus time histogram and latency, Figure 9) and analyzed in separation from ‘putative’ cochlear nucleus neurons (Figure 10). Since all firing of the individual SGN is thought to be driven by transmitter release from a single IHC AZ (Heil et al., 2007; Liberman, 1978; Robertson and Paki, 2002), these recordings provide insight into single AZ function. We first assessed the spontaneous firing activity and found an increased abundance of SGNs with low spontaneous firing rates in RBE^KO/KO mice (72% with rates < 10 Hz, n = 43 SGNs, N = 9 vs. 50% in RBE^WT/WT, n = 40 SGNs, N = 8; p=0.0267, Kolmogorov-Smirnov test; Figure 9A). Frequency tuning was intact in RBE^KO/KO SGNs (Figure 9B): the sharpness of tuning expressed by the Q_10dB (width of tuning curve 10 dB above threshold at the characteristic frequency (C_f) normalized by C_f) was comparable (mean: 9.28 ± 1.01, S.D. = 6.32 and median: 7.41 for RBE^KO/KO SGNs, n = 39 SGNs, N = 9 vs. mean: 12.50 ± 1.98, S.D. = 11.91 and median: 8.36 for RBE^WT/WT SGNs, n = 36 SGNs, N = 9; p=0.28, Mann-Whitney-Wilcoxon test). However, the sound threshold at C_f was significantly elevated by almost 20 dB in RBE^KO/KO mice (35.60 ± 3.45 dB SPL, S.D. = 22.66 dB SPL for RBE^KO/KO SGNs, n = 43 SGNs, N = 9 vs. 16.05 ± 2.47 dB SPL, S.D. = 15.42 dB SPL for RBE^WT/WT SGNs, n = 39 SGNs, N = 9, p<0.0001, Mann-Whitney-Wilcoxon test; Figure 9C). Given the normal frequency tuning and DPOAE, this threshold increase seems unlikely to result from a putative functional cochlear deficit upstream of the IHCs.

Figure 9

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Figure 10

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Next, we studied the firing response of SGNs to 50 ms tone bursts (at C_f and 30 dB above sound threshold, 200 ms inter-stimulus interval), which is governed by the presynaptic glutamate release and postsynaptic spike generation. The peak firing rate at sound onset is thought to reflect the initial rate of release from the SV-occupied release sites of the RRP (‘standing RRP’, [Oesch and Diamond, 2011; Pangršič et al., 2012]). Refractoriness and the decline of release rate due to partial depletion of the standing RRP likely dominate the subsequent spike rate adaptation. Finally, the adapted firing rate reports SV replenishment and subsequent fusion (reviewed in Pangršič et al., 2012; Rutherford and Moser, 2016). We observed reduced spike rates of SGNs from RBE^KO/KO mice (Figure 9D,E) both at sound onset (p=0.0001, n = 39 SGNs, N = 8 in RBE^KO/KO and n = 38 SGNs, N = 9 in RBE^WT/WT, t-test) and after short-term adaptation (p=0.0023, Mann-Whitney-Wilcoxon test). Both, peak and adapted rates were similarly affected by the RIBEYE-disruption, indicated by the scatter plot of peak vs. adapted rates (Figure 9E). A significant peak rate reduction was also observed at higher stimulation frequencies (10 Hz, Figure 9G–H). The spike rates were better preserved in RBE^KO/KO SGNs than in SGNs of bassoon mutant mice (Bsn^Δex4/5 data of Jing et al. (2013), purple data in Figure 9G–I). We approximated adaptation within the 50 ms response by single-exponential fitting since double exponential fitting did not regularly report two temporally discernible components in RBE^KO/KO SGNs. The mean apparent adaptation time constant reported by single-exponential fitting were significantly slowed in RBE^KO/KO SGNs (9.83 ± 0.50 ms, S.D. = 2.85 ms, median: 10.46 ms, n = 32 SGNs, N = 8) as compared to RBE^WT/WT SGNs (8.71 ± 0.50 ms, S.D. = 3.05 ms, median: 8.73 ms, n = 37 SGNs, N = 9, p=0.033, Mann-Whitney-Wilcoxon test). The results of double-exponential fitting of RBE^WT/WT and RBE^KO/KO SGNs support the slowed adaptation kinetics and are presented in Table 1. As expected for the reduced peak firing rate, we found prolonged first spike latency which also showed greater temporal jitter (Figure 9F). The reduced peak firing rate together with increased first spike latency jitter likely explain the reduction in ABR wave I amplitude. The firing of putative AVCN neurons was better preserved: putative bushy cells showed normal sound driven rates and chopper cells only a mild reduction in peak rate (Figure 10).

Next, we explored the encoding of sound intensity by estimating the mean firing rate during 50 ms tone bursts at different sound pressure levels. These ‘rate-level functions’ (Figure 11A) indicated that the spike rate increase with the sound pressure level (p=0.068, n = 24 SGNs, N = 8 in RBE^KO/KO and n = 19 SGNs, N = 7 in RBE^WT/WT, Mann-Whitney-Wilcoxon test, Figure 11—figure supplement 1A) and the dynamic range of sound coding (sound pressure level for which the spike rate changes from 10–90%, Figure 11—figure supplement 1B, p=0.3044, t-test) were not significantly altered. We then used transposed tones (C_f at 500 Hz modulation frequency) in order to probe for the temporal fidelity and reliability of firing in RBE^KO/KO SGNs in the steady state (Figure 11D). These experiments corroborated the reduced maximal firing rate of RBE^KO/KO SGNs (n = 22 SGNs, N = 7 in RBE^KO/KO and n = 15 SGNs, N = 6 in RBE^WT/WT, p<0.0001, t-test) and indicated that the temporal precision of sound coding is impaired also in the steady state (reduced Synchronization Index: p=0.0043, t-test).

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In order to further scrutinize the potential role of the synaptic ribbon in vesicle replenishment, we studied the response to prolonged tone-stimulation (Figure 11B, 500 ms at C_f and 30 dB above threshold, 2 s inter-stimulus interval). The peak rate was better preserved in RBE^KO/KO SGNs than seen with shorter inter-stimulus interval (e.g. 200 ms, Figure 9C), likely reflecting more complete SV-replenishment (i.e. larger standing RRP) owing to the longer recovery interval (2 s vs. 200 ms). However, the adapted spike rate of RBE^KO/KO SGNs was even more reduced than found with 50 ms tone bursts (to about half of that for RBE^WT/WT SGNs, Figure 11B,C) highlighting the impaired SV replenishment during prolonged stimulation. Finally, we evaluated presynaptic vesicle pool dynamics by recording and modeling responses to forward masking protocols (Harris and Dallos, 1979), that are thought to reflect depletion and recovery of the RRP (Figure 11E,F; Figure 11—figure supplement 1C). We approximated the recovery from forward masking by single exponential fitting (Figure 11—figure supplement 1C) to provide an estimate of the kinetics of vesicle pool replenishment. The time constant of recovery was prolonged in RBE^KO/KO SGNs (90.80 ± 8.66 ms, S.D. = 45.00 ms, n = 27 SGNs, N = 8 vs. 33.53 ± 5.74 ms, S.D. = 28.11 ms, in RBE^WT/WT SGNs, n = 24 SGNs, N = 7, p<0.0001, Mann-Whitney-Wilcoxon test) indicating slowed RRP replenishment in the absence of the ribbon. We noted that SGNs showed considerably lower spontaneous and evoked rates during the forward masking paradigm compared to other stimulus protocols (compare Figure 11B E), likely due to enhanced RRP depletion with the more extended stimulation in this protocol.

Amplitude and waveform of the forward masking responses were consistent with a two-fold reduction of the number of contributing vesicular release sites of the RRP (N’_slot: contributing release sites during forward masking, N_slot: the contributing release sites for the same AZs during tone bursts at 5 Hz stimulation). The ratio N_slots/N’_slots was estimated to be 2.2 for RBE^KO/KO and 2.3 for RBE^WT/WT from the drop in spontaneous and evoked SGN spiking rates. We used a previously developed biophysical model of RRP dynamics and spike generation (Frank et al., 2010; Jung et al., 2015b) to extract information on fusion and replenishment rate constants as well as the N_slot by fitting the responses to tone bursts, that is PSTHs with 100, 200 and 2000 ms inter-stimulus interval, and also the forward masking spiking data across all recovery intervals (Figure 11E). Importantly, only the first 50 ms of the 500 ms stimulus response during the PSTH 2000 were included in the fit; any later adaptation processes were disregarded, as they were not accounted by the model’s equations.

The results of model fitting suggested that during the forward masking only about half of all release sites (N_slots) were engaged in the response (N’_slots). Throughout, RBE^KO/KO SGNs showed a lower fusion rate than the RBE^WT/WT SGNs, reflecting the reduced onset response in RBE^KO/KO SGNs. When more recovery time was allowed, that is in the 0.5 Hz tone burst and the forward masking, where recovery times from around 250 to 500 ms occured between the probe and subsequent masker, the onset response improved in RBE^KO/KO. Consequently, the estimated fusion rate almost reached the level of RBE^WT/WT in the forward masking fits. With the scaling factors of approximately 2, the estimates for the number of release sites were consistent between tone bursts and forward masking data, and in both cases only slightly smaller for the RBE^KO/KO (Table 2). Taken together the fits from forward masking and tone bursts suggest that the total number of release sites (RRP) was only slightly reduced at ribbonless synapses of RBE^KO/KO IHCs. However, a strong firing response at sound onset, that is release of a large standing RRP, required longer recovery indicating more efficient SV replenishment in the presence of ribbons, which is reflected in the larger refilling rate constants estimated by the model of RBE^WT/WT synapses.

Our work confirms the central role of RIBEYE for forming synaptic ribbons (Schmitz et al., 2000; Magupalli et al., 2008; Maxeiner et al., 2016). We did not observe structures reminiscent of ‘ghost ribbons’ reported for ribeye mutants in zebrafish neuromast hair cells (Lv et al., 2016) in IHCs of RBE^KO/KO mice. These ghost ribbons were characterized as a halo of synaptic vesicles around a non-electron-dense area that resembled in size, though smaller, and shape to a synaptic ribbon. In zebrafish, two gene copies of ribeye (ribeye a and b) exist, making it harder to achieve a complete knock-out (Lv et al., 2016; Van Epps et al., 2004). In keeping with this notion, Lv et al. found residual immunofluorescence of ribeye a in the double mutants. Hence, we speculate that residual RIBEYE, possibly together with other scaffold proteins such as piccolo, might have formed the observed electron-translucent SV-framed structures (Lv et al., 2016). In contrast, immunofluorescence, as well as electron microscopy, revealed the complete absence of RIBEYE and ribbons in IHCs of RBE^KO/KO mice in our work and the companion study (Becker et al.), which is in agreement with findings in the RBE^KO/KO mouse retina (Maxeiner et al., 2016).

IHC synapses normally employ a single ribbon-type AZ. But in the absence of RIBEYE, there were typically two or more ribbonless AZs, akin to multiple conventional AZs (Figure 2). These ribbonless ‘conventional’ AZs at RBE^KO/KO IHC synapses consist mostly of roundish PDs, each with a cluster of SVs, of which approximately one third were directly adjacent to the plasma membrane (membrane-proximal: MP-SVs). Using electron tomography we found that about two-thirds of the MP-SVs were tethered to the AZ membrane, which was comparable to RBE^WT/WT AZs (Figure 3). We speculate that SVs associated with the PD, but not facing the membrane (PDA-SVs), serve to replenish the release sites once tethered MP-SVs fused, and that the ribbonless PD more likely acts in long-range SV tethering to the AZ in analogy to what is considered for conventional AZs (Cole et al., 2016; Fernández-Busnadiego et al., 2013; Siksou et al., 2007). We assume that absence of RIBEYE does not alter SV size since electron tomography, which provides the most reliable estimation of SV size, did not reveal differences in SV diameter between RBE^KO/KO and RBE^WT/WT AZs, at least when considering all SVs. The RBE^KO/KO PDs, like in RBE^WT/WT, contained bassoon, Ca_V1.3, and RIM2, but lacked piccolino which is likely part of the ribbon in RBE^WT/WT (Figure 1) (Dick et al., 2001; Khimich et al., 2005; Limbach et al., 2011; Regus-Leidig et al., 2013).

To some extent, the multi-AZ morphology is reminiscent of IHC synapses prior to synaptic maturation (Huang et al., 2012; Sendin et al., 2007; Sobkowicz et al., 1982; Wong et al., 2014). While we cannot rule out some sort of developmental delay of RIBEYE-deficient IHCs, we suspect that the morphological transformation into a multi-AZ morphology reflects a compensatory effort. Reasons for our interpretation include (i) the same morphological phenotype of RBE^KO/KO IHCs at 8 months of age (Figure 2), (ii) the finding of highly regular PDs at RBE^KO/KO IHCs synapses, which differs from less well-defined PDs at immature AZs (Wong et al., 2014), (iii) the typical continuous and large PSD of RBE^KO/KO IHCs synapses (Figure 2, see also the accompanying paper by Becker et al.) as a characteristic of a mature synapse, rather than the several smaller PSD patches at developing IHC synapses (Wong et al., 2014), and (iv) the synaptically confined Ca_V1.3 Ca²⁺-channel clusters, normal amplitude of IHC I_Ca and mature amplitude of synaptic Ca²⁺-signals, rather than massive extrasynaptic Ca_V1.3 abundance and larger whole-cell I_Ca but smaller synaptic Ca²⁺-signals in immature IHCs (Wong et al., 2014; Zampini et al., 2010). The multi-AZ morphology of the RBE^KO/KO IHC synapses was also corroborated by high- and super-resolution microscopy of bassoon and Ca_V1.3 immunofluorescence (Figure 4). The organization in several smaller Ca²⁺-channel clusters likely explains the broader spread of the presynaptic Ca²⁺-signal at RBE^KO/KO synapses (Figure 6). In contrast to bassoon mutant mice (Frank et al., 2010; Jing et al., 2013), the number of synaptic Ca²⁺-channels was not reduced in RBE^KO/KO mice as shown by normal amplitudes of whole-cell I_Ca and synaptic Ca²⁺-signals. Therefore, the loss of synaptic Ca²⁺-channels from the bassoon-deficient ribbonless IHC synapses, indicates a role of bassoon in promoting Ca²⁺-channel tethering at the AZ likely via interaction with RIM-binding protein (Davydova et al., 2014), which was previously shown to interact with Ca_V1.3 Ca²⁺-channels (Hibino et al., 2002) and is required for establishing a normal Ca²⁺-channel complement of the IHC AZ (Krinner et al., 2017).

Interestingly, we observed changes in Ca²⁺-channel function in RBE^KO/KO IHCs: the voltage-dependence of Ca²⁺-channel activation was slightly, but significantly, shifted to more depolarized potentials both at the levels of whole-cell Ca²⁺-current (V_h +2 mV) and synaptic Ca²⁺-influx at individual synapses (V_h +5 mV) (at 5 mM [Ca²⁺]_e, Figure 5 and 6). Similar as in this study, an enhanced inactivation (Figure 5) of I_Ca was also found in bassoon-deficient IHCs, while their V_h was actually mildly shifted in the opposite direction (−3 mV for imaging of synaptic Ca²⁺) and unaltered at the level of the whole-cell l_Ca (Frank et al., 2010). One potential reason for why the depolarized V_h-shift of the synaptic Ca²⁺-influx was greater than that of the whole-cell Ca²⁺-influx is the contribution of extrasynaptic Ca²⁺-channels to the whole-cell Ca²⁺-influx. They are thought to contribute approximately 30% of the Ca²⁺-influx (Brandt et al., 2005) and are not regulated by RIBEYE/ribbon. In order to test whether the depolarized V_h-shift of synaptic Ca²⁺ translates into changes in transmitter release, we recorded exocytic ΔC_m for different depolarization potentials. A small, but significant reduction of exocytosis for weak depolarizations in RBE^KO/KO IHCs (Figure 7, seen also in the accompanying paper by Becker et al.) suggests that the V_h shift is relevant for hair cell transmission (see also below). How enhanced I_Ca inactivation might affect sound encoding is addressed by work on Ca²⁺-binding proteins (CaBPs) that are thought to antagonize calmodulin’s role in mediating I_Ca inactivation (Lee et al., 1999; Peterson et al., 1999). Among the several CaBPs expressed in IHCs (Cui et al., 2007; Picher et al., 2017; Schrauwen et al., 2012; Yang et al., 2006), CaBP2 is defective in human genetic hearing loss DFNB93 (Picher et al., 2017; Schrauwen et al., 2012) and required for hearing likely via inhibition of IHC I_Ca inactivation (Picher et al., 2017). However, deletion of CaBP4 in mice caused only a very mild increase of I_Ca inactivation similar to the one found here and did not alter auditory brainstem responses (Cui et al., 2007). Future studies need to address how RIBEYE/ribbons mechanistically regulate the function and spatial organization of Ca²⁺-channels.

Over some decades, research on retinal photoreceptors and bipolar cells, on hair cells of the inner ear and the lateral lines, on electroreceptors as well as pineal cells, has aimed to elucidate the function(s) of the synaptic ribbon. Current hypotheses state that the ribbon functions in (i) replenishing release sites ([Bunt, 1971; Frank et al., 2010; von Gersdorff et al., 1996; Lenzi et al., 2002; Maxeiner et al., 2016; Snellman et al., 2011; Vaithianathan and Matthews, 2014] for a deviating view see {Jackman et al., 2009]), potentially by facilitated diffusion of SVs on the ribbon surface towards the site of consumption (Graydon et al., 2014) and SV priming (Grabner and Zenisek, 2013; Snellman et al., 2011), (ii) establishing a large complement of vesicular release sites and Ca²⁺-channels at the active zone (Frank et al., 2010; Khimich et al., 2005), which remained hard to disentangle from potential function of bassoon (Frank et al., 2010; Jing et al., 2013), (iii) ensuring close spatial coupling of Ca²⁺-channels and vesicular release sites (Maxeiner et al., 2016) or enhancing presynaptic Ca²⁺-signals by limiting diffusional Ca²⁺-spread (Graydon et al., 2011), (iv) contributing to multivesicular release (Graydon et al., 2011; Jing et al., 2013; Mehta et al., 2013), and (v) contributing to SV reformation from endocytosed membranes (Jung et al., 2015b; Khimich et al., 2005; Schwarz et al., 2011). Clearly, SV-replenishment was impaired at the ribbon-less IHC synapses of RBE^KO/KO mice. This is shown by slowed recovery from forward-masking and the use-dependent reduction of peak and adapted firing rates, which we further scrutinized by modeling. Therefore, our study supports a role of the ribbon in vesicle replenishment, which is also found in the accompanying paper by Becker et al.. Why RRP-recovery was not significantly altered when probed with pairs 20 ms long maximal Ca²⁺-influx by membrane capacitance measurements in IHCs (Figure 7) will need to be addressed in future studies, ideally using paired pre-and postsynaptic recordings of synaptic transmission with depolarizations of varying strength.

Each of the ribbon-manipulations employed to analyze its role has strengths, but also weaknesses, such as changes in other AZ proteins and long-term compensatory processes (e.g. bassoon deletion), complex manipulations (e.g. diurnal changes or hibernation: [Hull et al., 2006; Mehta et al., 2013; Spiwoks-Becker et al., 2004]) and photoablation (Mehta et al., 2013; Snellman et al., 2011). Genetic RIBEYE manipulations (Lv et al., 2016; Maxeiner et al., 2016; Sheets et al., 2011; Van Epps et al., 2004) have the greatest molecular specificity, but in some cases, were incomplete, and to some extent masked by compensation. In fact, our study of IHCs, unlike the situation for bipolar cell retinal ribbons (Maxeiner et al., 2016), suggests that some features of the IHC ribbon-type AZ can be very well replaced by a ribbonless multi-AZ morphology: the synaptic complement of Ca²⁺-channels and SVs, as well as exocytic ΔC_m elicited by strong depolarizations, were similar. Therefore, we likely underestimated the role of the ribbon in sound encoding in our present study.

For sound encoding at the afferent synapses between IHCs and SGNs, we observed some commonalities and differences with the bassoon mutants and RBE^KO/KO mice, whereby the stronger phenotype of bassoon mutants suggests additive effects of bassoon and ribbon loss. Recordings from single SGNs indicate reduced peak and adapted release rates at the IHC synapses of ribbonless synapses, as well as impaired temporal precision of coding. High temporal precision is a hallmark of synaptic sound encoding (e.g. (Köppl, 1997). Reduced release rates or smaller EPSC sizes would increase the temporal jitter (Buran et al., 2010; Li et al., 2014; Rutherford et al., 2012; Wittig and Parsons, 2008). Reduced spike rates and increased jitter of release likely explain the reduced ABR wave I amplitude in both mutants. A striking difference from bassoon mutants, however, is that sound encoding in RBE^KO/KO mice was impaired substantially, despite unaltered exocytic ΔC_m upon strong stimulation. We propose two mechanisms with likely additive effects to explain this surprising finding: i) the small depolarized shift of synaptic Ca²⁺-channels might contribute the lower spontaneous and evoked firing rates as well as higher sound thresholds of SGNs and ii) the reduced SV-replenishment might not suffice to balance the rate of consumption leading to a smaller standing RRP in vivo, while the arrest of exocytosis in the voltage-clamped IHCs for tens of seconds likely enables complete filling of the release sites (max. standing RRP). The changes in Ca²⁺-channel gating observed in IHCs of RBE^KO/KO mice were unexpected, as so far, a direct or indirect interaction of RIBEYE and Ca²⁺-channels have not been described. Clearly, future studies, including studies on the potential regulation of Ca_V1.3 Ca²⁺-channels by RIBEYE and piccolino in heterologous expression systems, paired pre- and postsynaptic recordings, as well as further computational modeling, are required.

Constitutive RIBEYE knockout mice (RBE^KO/KO derived from Ctbp2^tm1.2Sud by Cre-recombination) were generated by Maxeiner and colleagues (Maxeiner et al., 2016) and were back-crossed to C57BL/6 for five generations (corresponding to a C57BL/6 background contribution of >95%). All experiments complied with national animal care guidelines and were approved by the University of Göttingen Board for Animal Welfare and the Animal Welfare Office of the State of Lower Saxony (permit number: 14–1391). The colony was maintained by mating heterozygous mice. Whenever possible, experiments were performed in parallel on mutant mice and their wildtype littermates. However, the experimental schedule did not always permit this and we occasionally used individual mice from the same colony but without littermate controls. Moreover, for some experiments giving rise to Figure 6—figure supplement 1 and Figure 7E,F,G,H we also used C57Bl/6 wild-type mice and combined their results with those of wildtype littermate controls.

The apical 2/3 turns of organs of Corti from P14 to P28 aged mice were freshly dissected in HEPES Hank's solution containing (in mM): 5.36 KCl, 141.7 NaCl, 10 HEPES, 0.5 MgSO_4-7H₂O, 1 MgCl₂-6H₂O, 5.6 D-glucose, and 3.4 L-glutamine (pH 7.2, ~300 mOsm). The basolateral membranes of the IHCs were exposed by carefully removing the surrounding cells with a suction pipette. All experiments were conducted at room temperature (20–25°C).

Perforated-patch-clamp recordings were performed as described previously (Moser and Beutner, 2000). The pipette solution contained (in mM): 130 Cs-gluconate, 10 tetraethylammonium (TEA)-Cl, 10 4-AP, 10 HEPES, 1 MgCl₂, as well as 300 μg/ml amphotericin B (pH 7.2, ~280 mOsm). The extracellular solution contained the following (in mM): 112 NaCl, 35 TEA-Cl, 2.8 KCl, 1 MgCl₂, 1 CsCl, 10 HEPES, 2 CaCl₂, and 11.1 D-glucose (pH 7.2, ~305 mOsm). External solution change was done by bath exchange through a perfusion system.

For live-imaging, the patch pipette solution contained (in mM): 111 Cs-glutamate, 1 MgCl₂, 1 CaCl₂, 10 EGTA, 13 TEA-Cl, 20 HEPES, 4 Mg-ATP, 0.3 Na-GTP and 1 L-Glutathione (pH 7.3, ~290 mOsm). To visualize the Ca²⁺-hotspots and the ribbons, the Ca²⁺-indicator Fluo-4FF penta-K⁺ salt (0.8 mM, Life Technologies, Germany) and the TAMRA-conjugated CtBP2/RIBEYE-binding dimer peptide (10 µM, Biosynthan, Germany) were added to the intracellular solution. The extracellular solution contained the following (in mM): 2.8 KCl, 102 NaCl, 10 HEPES, 1 CsCl₂, 1 MgCl₂, 5 CaCl₂, 35 TEA-Cl, and 11.1 D-Glucose (pH 7.2, ~300 mOsm).

EPC-10 amplifiers controlled by Patchmaster or Pulse software (HEKA Elektronik, Germany) were used for the measurements. IHCs were held at −87 mV or −69 mV. All voltages were corrected for liquid junction potential offline (17 mV or 14 mV, depending on intra- and extracellular solutions used) and voltage-drops across the series resistance (R_s). Currents were leak corrected using a p/10 protocol in exocytosis experiments. Recordings were discarded when the leak current exceeded −55 pA, R_s exceeded 30 MΩ (for perforated-patch) or 15 MΩ within 4 min after break-in (for ruptured-patch), or Ca²⁺-current rundown exceeded 25%. All passive electrical properties of the patch-clamp recording experiments are detailed in Supplementary file 2.

Exocytosis was studied by measuring the membrane capacitance increments (ΔC_m) using the Lindau-Neher technique (Lindau and Neher, 1988). Cells were stimulated by step depolarizations of different durations to −17 or −14 mV, or by 100 ms pulses to voltages ranging from −53 to −37 mV. A resting interval of 10–100 s between the stimuli was used. Each protocol was applied two to three times and only IHCs with reproducible exocytosis during the rounds were included. For display, traces were subjected to 1, 5 or 10 pass Binomial Smoothing using Igor Pro. Current-voltage relationships (‘IVs’) displayed in Figure 5A (ruptured-patch, 3-week-old mice) were obtained by applying 20 ms depolarizing step pulses of increasing voltage from −82 to 63 mV in 5 mV steps.

Ca²⁺-imaging was performed with a spinning disk confocal scanner (CSU22, Yokogawa, Germany) mounted on an upright microscope (Axio Examiner, Zeiss, Germany) with 63x, 1.0 NA objective (W Plan-Apochromat, Zeiss). Images were acquired by a scientific CMOS camera (Neo, Andor, Germany). Ca²⁺-indicator F4FF and TAMRA-conjugated peptide were excited by diode-pumped solid-state lasers with 491 nm and 561 nm wavelength, respectively (Cobolt AB). The spinning disk was set to 2000 rpm to synchronize with the 10 ms acquisition time of the camera.

Using a piezo positioner for the objective (Piezosystem, Germany), a scan of the entire cell was performed 4 min after breaking into the cell, taking sections each 0.5 µm at an exposure time of 0.5 s in the red (TAMRA-peptide) channel from the bottom to the top of the cell. In order to study the voltage-dependence of Ca²⁺-indicator fluorescence increments at the synapses, the confocal scans were acquired every 0.5 µm from the bottom to the top ribbon in the RBE^WT/WT mice. For the RBE^KO/KO mice, the scanning was done from the bottom of the cell to +12 µm, which on average corresponds to the bottom of the nucleus. Ca²⁺-currents were evoked by applying a voltage ramp stimulus from −87 to +63 mV during 150 ms (1 mV/ms) in each focal plane. Simultaneously, fluorescence measurements were made in the green channel (Fluo-4FF) with a frame rate of 100 Hz. In order to overcome the limitations of the frame rate and increase the voltage resolution of the fluorescent signal acquired, the voltage ramp protocol was applied twice, once shifted by 5 ms such that for any given frame during the second ramp the voltage was shifted by 5 mV compared to the first stimulus. Alternating planes were acquired to avoid photobleaching encountered with the consecutive plane acquisition.

Apical turns of organs of Corti from 3-week-old mice were prepared for ‘whole-mount imaging’ as described in (Ohn et al., 2016). In brief, the samples were fixed either in formaldehyde (4%, 10 min on ice), or methanol (20 min at −20°C). Afterwards, the following primary antibodies were used: mouse anti-CtBP2 (1:200, BD Biosciences, Germany 612044), mouse anti-PSD-95 (1:200, Sigma Aldrich, Germany P246-100ul), mouse anti-bassoon SAP7f407 (1:200, Abcam, Germany, ab82958), guinea pig anti-bassoon (1:500, Synaptic Systems, Germany, 141 004), rabbit anti-RIM2 (1:100, Synaptic Systems 140 103), rabbit anti-Ca_v1.3 (1:75 or 1:100, Alomone Labs, Germany, ACC 005), rabbit anti-piccolino (1:500, kind gift of JH Brandstätter; see Regus-Leidig et al., 2013), guinea pig anti-parvalbumin α (1:1000, Synaptic Systems, 195 004), mouse anti-calbindin 28 k (1:500, Swant, Germany, 07(F)), and rabbit anti-calretinin (1:1000, Swant 1893–0114). Secondary antibodies used were Alexa Fluor 488 conjugated anti-rabbit, Alexa Fluor 488 conjugated anti-guinea-pig, Alexa Fluor 568 conjugated anti-mouse, and Alexa Fluor 647 conjugated anti-rabbit (1:200, Invitrogen, Germany, A11008, A11004, A11073, and A31573 respectively). For high resolution STED microscopy, STAR580 and STAR635p conjugated anti-rabbit and anti-mouse (1:200, Abberior, Germany, 2-0002-005-1, 2-0012-005-8, 2-0002-007-5, and 2-0012-007-2) have been used as secondary antibodies. Images were acquired using either a Leica SP5 with a 1.4 NA 63x oil immersion objective or an Abberior Instruments Expert Line STED microscope, with excitation lasers at 488, 561, and 633 nm and STED lasers at 595 nm, 1 W, and 775 nm, 1.2 W, using a 1.4 NA 100x oil immersion objective, either in confocal or in 2D-STED mode. Images were adjusted for brightness and contrast using ImageJ.

ABR, DPOAE and extracellular recordings from single SGNs were performed essentially as described before (Jing et al., 2013; Strenzke et al., 2016). ABR and DPOAE recordings were performed on 6-week-old mice. For extracellular recordings from individual SGNs, 6 to 10 week-old mice were anesthetized by i.p. injection of urethane (1.32 mg/kg), xylazine (5 mg/kg) and buprenorphine (0.1 mg/kg), a tracheostomy was performed and the mice were then placed in a stereotactic system. After partial removal of the occipital bone and cerebellum to expose the anteroventral cochlear nucleus (AVCN), a glass microelectrode was advanced through the posterior AVCN portion to reach the auditory nerve. Acoustic stimulation was provided by an open field Avisoft ScanSpeak Ultrasonic Speaker (Avisoft Bioacoustics, Germany), and ‘putative’ SGNs (auditory nerve fibers formed by the central SGN axons) were identified and distinguished from cochlear nucleus neurons based on their stereotactic position (>1.1 mm from the surface of the cochlear nucleus), spontaneous and noise-burst induced firing, peristimulus time histogram (PSTH), regularity of firing, and first spike latency. Recordings were performed using TDT system III hardware and an ELC-03XS amplifier (NPI Electronics, Germany), offline analysis using waveform-based spike detection using custom-written MATLAB software (Source code 1).

High-pressure freezing, freeze-substitution followed by electron tomography were essentially performed as described previously (Vogl et al., 2015; Jung et al., 2015a). After freeze-substitution and embedding in epoxy resin (Agar 100 kit, Plano, Germany), 250 nm semithin sections for electron tomography were obtained on an Ultracut E ultramicrotome (Leica Microsystems, Germany) with a 35° diamond knife (Diatome, Switzerland). Sections were placed on 1% formvar-coated (w/v in water-free chloroform) copper 100 mesh grids (ATHENE, Plano, Germany, 3.05 mm Ø) and post-stained with UAR-EMS (Science Services, Germany) and Reynold’s lead citrate.

For electron tomography, 10 nm gold particles (British Bio Cell/Plano, Germany) were applied to both sides of the stained grids. Single tilt series at 12,000-x magnification, mainly from −60 to +60° (if only fewer angles were possible, the tomograms were only accepted for quantification if the quality was sufficient) were acquired with an 1° increment at a JEM2100 (JEOL, Germany)) transmission electron microscope at 200 kV using the Serial-EM software (Mastronarde, 2005). The tomograms were generated using the IMOD package etomo and models were generated using 3dmod (Kremer et al., 1996).

Live-imaging and IHC-patch-clamp data were analyzed using custom programs in Igor Pro 6.3 (Wavemetrics, Portland, OR, USA; Source Code 2). For analysis of IV-curves, the evoked Ca²⁺-current was averaged from 5 to 10 ms after the start of the depolarization. The total Ca²⁺-charge was estimated by taking the integral of the leak-subtracted current during the depolarization step. For most protocols, ΔC_m was estimated as the difference between the mean of C_m 400 ms after and before the depolarization (the initial 60 ms after the end of depolarization were skipped). For paired pulse experiments, the calculation of the mean of C_m before and after the depolarization was limited to the time remaining in the inter pulse interval after skipping (the initial 30 ms after the end of depolarization were skipped).

ΔF images were generated by subtracting the fluorescence intensity inside the cell at the resting state (F₀, an average of 10 frames) from the one at the depolarized state (an average of 6 frames during voltage ramp protocol). ΔF was calculated as the average of a 3 × 3 pixel square placed in the region showing the greatest intensity increase within the fluorescence hotspot. Maximal ΔF (ΔF_max) was the average of 5 ΔF values obtained between −17 and +8 mV during the voltage ramp (around the peak Ca²⁺-influx). Only AZs presenting a ΔF_max greater than the mean of the fluorescence intensity plus two standard deviations at rest were considered for further analysis. For analysis of the voltage dependence of synaptic Ca²⁺-signals, raw traces were fitted to the following

where $f_v$ is the fluorescence-voltage-relationship ΔF/ΔV obtained by linear fitting to the FV-curve in the range of 3 to 23 mV, $V_r$ the reversal potential of 65.6 mV, and $V$ the command voltage, in order to obtain $V_h$, the voltage of half-maximal activation, and $k$, the slope factor. The spatial extent of the synaptic Ca²⁺-signals was estimated by fitting of a 2D Gaussian function to the fluorescent hotspot using a genetic fit algorithm (Sanchez del Rio and Pareschi, 2001) to obtain the full width at half maximum in the long and short axis. For each spot, the calculations were made at those confocal sections where the intensity of the spot was strongest.

Activation time constants of Ca²⁺-currents at differing potentials were obtained by fitting to the first 5 ms of the current traces the following equation:

Confocal and STED immunofluorescence images were analyzed and z-projected with Fiji software and further analyzed using Igor Pro. For synapse counting, co-localized pre- and postsynaptic immunofluorescent spots were counted manually. The spatial extent of the line-shaped Ca²⁺-channel clusters was estimated by fitting a 2D Gaussian function to the individual clusters in 2D STED images to obtain the full width at half maximum in the long and short axis. The areas of the PSD were calculated by the following formula: area = π x (Long Axis/2) (Short Axis/2). The semi-quantitative immunofluorescence analysis of the proteinaceous Ca²⁺-buffers was performed by calculating the mean immunofluorescence intensity of a volume (40 (X) x 40 (Y) x 4 (Z) voxels or 2.8 × 2.8 × 2 µm) below the nucleus and above the synapses. This and the count and intensity of the CtBP2 immunofluorescent spots have been analyzed in Imaris 7.6.5 with custom Matlab routines (Source Code 4).

For extracellular SGN recordings, PSTHs were calculated as average firing rates across 200 presentations of 50 ms or 500 ms tone bursts presented at 0.1 s/0.2 s or 2 s intervals, resp. (PSTH at 10/5 Hz and 0.5 Hz) at C_f, 30 dB above the threshold and binned at a width of 2 ms. Peak rate was determined as the largest bin of the PSTH in a time window 3–11 ms after stimulus onset. Adapted rate was averaged in a window spanning 35–45 ms or 405–415 ms after stimulus onset (for PSTH at 5 Hz and 0.5 Hz, respectively). Rate level functions were acquired using 50 ms tone bursts presented at C_f and at 5 Hz. 25 repetitions for each stimulus intensity (5 dB steps) were recorded. Maximal steepness was calculated as the maximal increase in spike rate between two consecutive 5 dB increment steps. Dynamic range was calculated by using sigmoidal fits in the rate level functions as described in and measuring the range of sound pressure between 10% and 90% of maximal firing rate. For amplitude modulation analysis, synchronization index was calculated as described by (Goldberg and Brown, 1969). Synchronization index estimation was only considered valid when at least 15 spikes occurred in a 3 s time window and the Rayleigh statistic was below 13.8.

For analysis of forward masking experiments, spike counts in a 10 ms interval starting from responses of both masker and probe onset were determined and presented as the ratio of probe and masker responses for at least 25 repetitions for every masker-probe interval from each unit. Exponential fitting to the plots of each individual SGN approximated the recovery kinetics.

Quantitative analysis of electron microscopy data was performed with ImageJ for conventional embedded samples and with IMOD for HPF/FS tomograms. According to the presence of ribbon-occupied and ribbonless synapses, we considered the following analysis criteria:

For ribbon-occupied synapses, membrane-proximal synaptic vesicles (MP-SVs, within a distance of ≤25 nm from the AZ membrane and ≤80 nm from the presynaptic density) and ribbon-associated synaptic vesicles (RA-SVs, first layer around the ribbon with a maximum distance of 80 nm from the vesicle membrane to the ribbon) were counted (Figure 2J, random sections analysis according to Strenzke et al., 2016 and Figure 3G for tomograms according to 2D-random section analysis criteria). The tomogram analysis parameters were further modified, as used in Jung et al., 2015a. Here, the MP-SVs were defined as vesicles with ≤50 nm from the AZ membrane and with the shortest distance from the vesicle membrane to the presynaptic density of ≤100 nm, excluding RA-SVs (Figure 3—figure supplement 1A). For random sections, SV diameters were calculated by the averaged measurements of the horizontal and vertical axis. The ribbon size was measured in height and width, taking the longest axis of the ribbon excluding the PD, and the edges of the synaptic ribbon were traced manually using ImageJ. The length of the PD was measured along the AZ membrane (Figure 2J).

For ribbonless synapses, a presynaptic density-associated synaptic vesicle (PDA-SVs) pool was defined considering all clustered vesicles ≤80 nm around the PD that did not fulfill the criteria of a MP-SV (see above, also Figure 2J for random sections). The MP-SV pool, as well as the SV diameter and PD length, were analyzed as for the ribbon-occupied synapses. For tomograms, the PDA-SV pool was defined as the SVs in the first layer ≤ 80 nm to the PD, excluding the MP-SVs. The MP-SV pool criteria are the same as described in the previous paragraph (Figure 3G, according to 2D-random section and Figure 3—figure supplement 1A, according to Jung et al., 2015a). For tomograms, the according pools were further distinguished into tethered and non-tethered vesicles (Figure 3G and Figure 3—figure supplement 1A). All vesicles were annotated using a spherical ‘scattered object’ at its maximum projection in the tomogram, encompassing the outer leaflet of the vesicles. The vesicle radii were determined automatically (Helmprobst et al., 2015) with the program ‘imodinfo option -p’ of the IMOD software package (Kremer et al., 1996).

The coding of sound onset differs among the various SGNs in time due to different durations of the traveling wave, synaptic delays and conduction times. To obtain an average PSTH for modeling that is not smeared out due to such differences between units, the individual PSTHs were aligned beforehand by shifting their timing relative to each other. Onset detection was based on a change in spike statistics. For spontaneous activity, the 99.5 percentile of spike counts was determined. Next, the time at which response rises to twice this percentile was found. This is certainly a point within the sound response. Finally going back from this point, a drop back baseline activity, that is below the percentile was detected and used as onset time. Aligned PSTH from all units were averaged. This averaged PSTH from the forward masking data were fit with a model waveform using a genetic fit algorithm implemented in IGOR Pro (Wavemetrics, Lake Oswego, OR, USA). The purpose of the model is to give insight into the dynamics of SV cycling at the average IHC AZ. More specifically, the notion of Ca²⁺-nanodomain-like control of RRP exocytosis (Brandt et al., 2005; Graydon et al., 2011; Pangršič et al., 2015; Wong et al., 2014), as well as the limited MP-SVs at the AZ (see Figures 2 and 3) motivates the notion of a limited, quasi-fixed number of available vesicular release sites or slots, N_slot,(Frank et al., 2010; Wong et al., 2014) that constitute the RRP. Each of these sites can be either empty or occupied by a release ready SV (whereby all filled slots constitute the ‘standing’ RRP) and at each time point, a release ready SV will fuse with a certain probability described by the fusion rate constant k_fus. Its value depends on the sound pressure level in a relation we assume to be linear within the dynamic range of the synapse/fiber. While the sound pressure level rises from silence to saturation k_fus increases from k_{fus, spont} to k_fus,stim. The refilling of empty sites is described by a refill rate constant k_refill, which also depends on the sound intensity (k_{refill, spont} to k_refill,stim).

The state of the release site was described by:

Although this equation is formulated for SV fusion rates, a scaling factor f can be used to account for the fraction of fusion events that cannot successfully trigger an action potential (AP) despite sufficient neural excitability for example because of the too small size of the elicited excitatory post synaptic current. This factor effectively operates as if the number of release sites was scaled down. The scaled equation then gives a rate R of potentially supra-threshold EPSCs as the product of the number of occupied release sites, the fusion rate constant and the scaling factor f:

The stationary solutions of Equation 3 together with Equation 4 determine steady state occupancy and steady state event rates:

In this equation, ‘condition’ is either silence or saturating sound pressure level.

In order to connect the postsynaptic event rate of potentially supra-threshold EPSCs to the actual AP rate, refractoriness is considered as a combination of an absolute refractory period t_abs, during which the probability of an EPSC to trigger an AP is zero, with a relative refractory period during which this trigger probability returns to one with an exponential time course characterized by τ_rel (Berry and Meister, 1998). This description of refractoriness can be applied to the ‘driving’ EPSC rate R by means of a delayed differential equation. The equation is motivated by the concept of three possible states of the SGN: ‘absolute refractory’, ‘relative refractory’ or ‘available’ (fully excitable). At any point, the probability that the SGN turns from ‘available’ to ‘refractory’ is proportional to the rate R(t). The return back to ‘available’ happens ‘delayed’ by t_abs and with a probability that is proportional to 1/τ_rel.

Together with Equation 6 the stationary solution of this description of refractoriness connects the observable steady state rates during silence and stimulation to the rate constants k_refill und k_fusion:

To go beyond the description of steady state event rates and to use the model for a parameterized description of the actual time course of experimentally observed PSTHs acquired during forward masking (Figure 11E), it is necessary to define the relation between the applied stimulus and the fusion and refill rate constants. For the experimental data presented here, the stimulus level was increased from silence to 30 dB above fiber threshold within a 4 ms ramp having a quarter of a sin² shape. It was assumed that k_fusion and k_refill follow the stimulus increase simultaneously. The ordinary differential and delayed differential equations above were combined into a fit function. PSTHs (one masker followed by one probe) were averaged per genotype for each masker probe interval (4, 16, 64 and 256 ms) and were fitted in parallel with one parameter set. During experiments, trials were acquired in immediate succession without gaps. Therefore, the model implements cyclic boundary conditions for the occupancy of the slots. This model only captures the short term processes, assuming that a set of experiments, for example forward masking trials quickly lead to a steady state. Slow adapting processes were not explicitly modeled. The observed drop of the apparent number of available slots in the forward masking experiments was described here as a change in the number of slots from N_slots to a reduced capacity N’_slots and for a given spiral ganglion neuron that was tested with tone bursts and forward masking, the ratio N_slots/N’_slots could be estimated from the change in rates (see Results).

The data were analyzed using Matlab (Mathworks), Excel, Igor Pro 6 (Wavemetrics), Origin 9.0 (Microcal Software), and GraphPad Prism (GraphPad Software). Averages were expressed as mean ± standard error of the mean (S.E.M.). For every dataset, the standard deviation (S.D.), number of replicates (n) and animals (N) were indicated. For Figure 7, n_min corresponds to the minimum number of cells included in the analysis of each depolarization potential given that the number of cells for each potential differs. In order to compare two samples, data sets were tested for normal distribution (Jarque-Bera test, D’Agostino and Pearson omnibus normality test or the Shapiro-Wilk test) and equality of variances (F-test), followed by two-tailed unpaired Student’s t-test, or, when data were not normally distributed and/or variance was unequal between samples, the unpaired two-tailed Mann-Whitney-Wilcoxon test was used. Cumulative distributions in Figure 9A were statistically compared using the Kolmogorov-Smirnov test. The ROUT method (Q = 0.1%) from GraphPad Prism was used to identify definitive outliers for Figure 7H.

For multiple comparisons, statistical significance was calculated by using one-way ANOVA test (two-way ANOVA in the case of ABR thresholds) followed by Tukey’s test for normally distributed data or Kruskal-Wallis (K-W) test followed by non-parametric multiple comparisons test (NPMC) for non-normally distributed data.

For SV diameter quantifications in random sections, a custom-written routine using Java Statistical Classes library (JSC) (Bertie, 2002) was utilized for statistical analysis (Source code 5). Due to the tied ranks of SV diameter measurements obtained for random sections, their S.E.M. was used as a tolerance value for the usage of Kruskal-Wallis test as suggested by Bertie et al. in JSC library (Bertie, 2002), where two values were treated as equal if their difference was ≤ S.E.M.. The non-significant difference between samples is reported as n.s., significant differences are reported as *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank N Dietrich, T Goldak, S Gerke, C Senger-Freitag and I Herfort for expert technical assistance, Gerhard Hoch for software and analysis development, and Frank Schmitz for providing RBE^KO/WT mice.

Animal experimentation: All experiments complied with national animal care guidelines and were approved by the University of Göttingen Board for Animal Welfare and the Animal Welfare Office of the State of Lower Saxony (permit number: 14-1391).

© 2018, Jean et al.

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