To hear a sound, the pressure produced by sound waves must be converted into an electrical nerve signal. The cells inside the ear that perform this transformation are called hair cells, which are so named because they have hundreds of hair-like structures on their upper surface. Pressure from sound waves causes movements in the inner ear that bend these ‘hairs’. This causes the hair cells to release chemical signals to neighboring nerve cell terminals that ultimately transmit information about the sound to the brain.

The chemical signals are stored inside the hair cells in bubble-like compartments called vesicles. To release the chemicals from the cell, the vesicles merge with the membrane that surrounds the hair cell. Most cells that communicate in this way are limited in how long they can transmit such messages. However, hair cells can continuously fuse vesicles to the membrane even when a sound lasts for a long time. This suggests that the hair cells have a different way of producing vesicles and getting them to the membrane than other cell types.

Inside the hair cells, vesicles are stored in regions called active zones. Each active zone contains a “ribbon” (attached to which are hundreds of vesicles) and also ion channels that allow calcium ions to flow into the cell. (An increase in calcium ion concentration inside the cell is necessary for the vesicle to fuse with the cell membrane and so release its chemical content). Now, Vincent et al. show that in hair cells, a cage-like network made from a protein called actin surrounds each active zone. This network helps to position the calcium ion channels. Treating the hair cells with a compound that disorganized the actin networks speed up the process of vesicle movement, which suggests that the actin network also controls the rate at which vesicles reach the membrane.

Next, it will be important to identify how the actin network interacts with other molecules that help vesicles to release their contents; in particular a protein called otoferlin, which is thought to act as a calcium ion sensor.

Auditory hair cells convert tiny variations of sound pressure through the displacement of their apical hair bundles into analogous voltage waveforms. Neural encoding of these microphonic potentials occurs at the ribbon synapses of inner hair cells (IHCs) by mechanisms involving Cav1.3 channels (Platzer et al., 2000; Brandt et al., 2003; Brandt et al., 2005) and otoferlin-dependent exocytosis of synaptic vesicles (Roux et al., 2006; Beurg et al., 2010, Vincent et al., 2014). Unlike most neuronal central synapses, IHCs have the extraordinary property to sustain indefatigably high rates of exocytosis during continuous sound stimulation (Safieddine et al., 2012). The precise molecular mechanisms underlying this fast and massive recruitment of synaptic vesicles to the IHC ribbon active zones still remain elusive. The implication of an unconventional molecular regulation of synaptic vesicle fusion and replenishment of the releasable pool of vesicles has been proposed (Nouvian et al., 2011; Vogl et al., 2015). In other secretory cells such as neuroendocrine cells, a network of sub-membranous cortical F-actin is known to influence exocytosis greatly by tightly regulating plasma membrane tension and the access of the granules to the secretory sites (Apodaca, 2002; Torregrosa-Hetland et al., 2011; Gutierrez and Gil, 2011). In central neuronal synapses, F-actin is also involved in maintaining vesicle pools and regulating vesicle mobility (Cingolani and Goda, 2008). Electron-tomography evidence for F-actin and microtubules near the synaptic ribbons has been observed in bullfrog hair cells (Graydon et al 2011). Whether IHC synaptic exocytosis is modulated by F-actin remains unknown. Other factors affecting membrane tension and exocytosis in many cell types, such as mast cells, include hydrostatic pressure across the membrane (Solsona et al., 1998). This latter factor has also been shown to influence the spontaneous and evoked activity of vestibular hair cells of the dogfish by mechanisms that remain unknown (Fraser and Shelmerdine, 2002). In the present study, we investigated whether F-actin and intracellular hydrostatic pressure regulate synaptic exocytosis in mouse IHCs.

Direct labeling of F-actin with fluorescent phalloidin revealed the presence of a dense F-actin network that surrounded the IHC ribbon synaptic zones (Figure 1A). This network extended underneath the plasma membrane and formed intracellular dense cages beneath the synaptic ribbons. These cages displayed a mean size diameter of 0.8 ± 0.1 µm (n = 186 actives zones; Figure 1B). Overlapping with otoferlin, each F-actin cage was generally associated with one ribbon and one Cav1.3 co-immunoreactive patch. Similarly, Ca²⁺ channels and the secretory machinery have been shown to be associated with the borders of F-actin cytoskeletal cages in chromaffin cells (Torregrosa-Hetland et al., 2011).

Figure 1

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A 45-min treatment of the organ of Corti in vitro with 1 µM extracellular latrunculin-A completely disorganized the synaptic F-actin cages and the Cav1.3 immunoreactive-patches at the IHC ribbons (Figure 1C, D). The mean distance between the Cav1.3 immunoreactive patches and the ribbons increased from 218 ± 11 nm (n = 71) in controls to 260 ± 12 nm with latrunculin-A treatment (n = 102; p<0.05). In the latter conditions, surprisingly, whole-cell patch clamp recordings revealed a largely facilitated exocytosis as compared to controls, while voltage-gated Ca²⁺ currents were unchanged (Figure 2A,B). After 100 ms depolarization, from -80 to -10 mV, the exocytotic response in control IHCs reached a maximum amplitude of 22.0 ± 2.9 fF (n = 11). Considering that a 40 nm diameter synaptic vesicle corresponds to 37 aF (Lenzi et al., 1999), we estimated a RRP size of about 590 vesicles, i.e 33 vesicles per ribbons if we assume a number of 18 ribbons per IHCs. This number of RRP vesicles fit well with previous findings (Johnson et al., 2008; Vincent et al., 2014). In latrunculin-treated IHCs, the exocytotic responses reached, at 100 ms, 32.4 ± 3.3 fF (n = 18), a value significantly larger as compared to control IHCs (p<0.05; Figure 2B). Remarkably, while the exocytotic response saturated at 90–100 ms in controls, the response did not show saturation in latrunculin-treated IHCs. Since the exocytotic response was unchanged for short impulses below 30 ms (Figure 2B), these results suggested that the disruption of F-actin did not affect the steps of vesicle docking and priming but facilitated the replenishment of the RRP. In bassoon mutants with abnormal number of anchored ribbons and reduced Ca²⁺ currents both short (20 ms) and long (100 ms) impulse activated-exocytosis were affected (Jing et al., 2013).

Figure 2

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The exocytotic facilitation in latrunculin-treated IHCs was greatly reduced to 21.8 ± 2.5 fF at 100 ms (n = 7) and comparable to controls (p=0.9) when intracellular Ca²⁺ buffering was increased with 5 mM EGTA (Figure 2B). This sensitivity to EGTA suggested a spatial disorganization of the Ca²⁺ channel clusters in regards to the release sites in latrunculin-treated IHCs, in good agreement with confocal imaging (Figure 1C,D). Rising the EGTA concentration could also affect the extent to which neighboring calcium sources interact and summate to produce a global effect on free calcium.

The intriguing question now is: why is the disruption of the synaptic F-actin with latrunculin-A facilitating exocytosis in IHCs? One explanation is that the synaptic F-actin network, in addition to organizing Ca²⁺ microdomains, also acts as a diffusion barrier for synaptic vesicles limiting the access to the site of release, as suggested in some central synapses (Cingolani and Goda, 2008). Its disruption with depolymerising agents would therefore facilitate vesicle replenishment of the release sites, as shown in a large variety of secretory cells (Malacombe et al., 2006), by increasing the number of available vesicles for docking and priming. Alternatively, a disrupted F-actin could facilitate the diffusion of Ca²⁺ from its sites of entry and stimulate replenishment.

To directly test these hypothesis, we studied the effect of F-actin depolimerization on exocytosis triggered by intracellular Ca²⁺ uncaging, i.e. independently of the activation and the organization of the Ca²⁺ channels. In these experiments, a large exocytotic facilitation was also observed in latrunculin-treated IHCs (Figure 2C,D). The peak exocytotic rate, obtained by measuring the first derivative function of the curves in Figure 2C, was nearly two fold larger in latrunculin-treated IHCs as compared to controls (Figure 2D; 25.9 ± 2.7 pF/s (n = 14) and 13.7 ± 1.7 pF/s (n = 16), respectively; p<0.05). The levels of the peak intracellular Ca²⁺ concentration ([Ca²⁺]_free) reached upon UV-flash Ca²⁺ uncaging were verified to be similar in control (n = 7) and latrunculin-treated IHCs (n = 5), respectively 59 ± 7 µM and 57 ± 5 µM (p=0.9; Figure 2C-inset). These Ca²⁺ uncaging experiments again suggested that a synaptic F-actin network controls the diffusion rate of the synaptic vesicles to the sites of release in IHCs.

Furthermore, since intracellular hydrostatic pressure has been suggested to influence membrane tension and exocytosis through the F-actin network in many cell types such as mast cells (Solsona et al., 1998), we probed Ca²⁺-evoked exocytosis under various intracellular osmotic pressures in auditory IHCs. We first found that increasing intracellular osmotic pressure from 310 to 390 mOsm with sucrose produced a significant increase in the resting membrane capacitance of IHCs. The resting size of IHCs, voltage-clamped at -80 mV for a period of 2 min after break-in, was 9.93 ± 0.27 pF (n = 11, 310 mOsm) and 11.10 ± 0.34 pF (n = 10, 390 mOsm; p<0.05), respectively. This augmentation of the IHC resting membrane capacitance was about 50 times larger than the size of the RRP evoked by membrane depolarization (Figure 2B; RRP = 22 fF). Where does this large addition of membrane come from? One possible explanation was that high intracellular hydrostatic pressure triggers the fusion of a large amount of extrasynaptic vesicles to the plasma membrane, as previously suggested for Ca²⁺ uncaging (Vincent et al., 2014).

Remarkably, in these latter conditions of intracellular hyperosmotic stress at 390 mOsm, voltage-dependent Ca²⁺ currents displayed larger amplitude (Figure 3A,C) and accelerated activation kinetics as compared to control conditions at 310 mOsm (Figure 3E). A shift in their voltage-dependence toward more negative potentials was also observed (Figure 3A,D). The hydrostatic pressure effects on Ca²⁺ currents were no longer visible when IHCs were pre-treated with latrunculin-A (Figure 2A, pink line) and were greatly reduced in IHCs lacking otoferlin (Otof^-/-; Figure 3B,C). This indicated that the Cav1.3 channels of IHCs are mechanosensitive, like the Cav1.2 channels in smooth muscle cells (Lyford et al., 2002). We cannot exclude the addition of Ca²⁺ channels to the plasma membrane during the massive vesicular fusion caused by increased hydrostatic pressure. However, this mechanism appears unlikely in regards to results obtained in non-secretory cells such as smooth muscle and HEK cells where similar effect on Ca²⁺ channels were obtained (Lyford et al., 2002). In our study, the sensitivity to membrane tension in IHCs required an intact synaptic F-actin network and otoferlin, a proposed synaptic Ca²⁺ sensor thought to interact physically with the Cav1.3 II-III loop (Ramakrishnan et al., 2009). Notably, an increase in Ca²⁺ current amplitude in low bath hydrostatic pressure (equivalent to increasing intracellular pressure) was also reported in dissociated guinea pig vestibular hair cells (Duong Dinh et al., 2009; Haasler et al., 2009). In these latter studies, the change in Ca²⁺ currents could be interpreted as due to a pressure activation of K⁺ currents leading to less pronounced depolarization. Although pressure effects on K⁺ currents have also been shown in guinea-pig IHCs (Kimitsuki, 2013), we don't think that these currents would influence our Ca²⁺ current measurements since we were working in conditions where most K⁺ currents are blocked. Overall, our results suggested here that the mechanosensitivity of the Cav1.3 channels is mediated through an intact synaptic F-actin network.

Figure 3

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Exocytosis triggered by voltage-activation of Ca²⁺ channels from -80 to -10 mV (a voltage at which Ca²⁺ current amplitude is maximum, Figure 3A) also showed maximum facilitation when intracellular pressure was increased from 310 to 390 mOsm (Figure 3F,G). Notably, on the contrary to what observed when disrupting the synaptic F-actin, exocytotis increased for short impulses (10–20 ms) at 390 mOsm (Figure 3F,G), suggesting that the last steps of vesicle  exocytosis  (docking, priming and/or fusion) were here accelerated. Exocytosis for longer steps up to 90 ms also increased, from 1.40 ± 0.23 fF/pC (n = 7, 310 mOsm) to 2.58 ± 0.49 fF/pC (n = 5, 390 mOsm; p<0.05) at 50 ms (Figure 3G). These latter results suggested that the replenishment rate of the RRP was also accelerated.

This facilitation of exocytosis, under intracellular hyperosmotic stress, appeared unrelated to the effects on Ca²⁺ channels since it was also observed when directly uncaging intracellular Ca²⁺ (Figure 4). The levels of the peak [Ca²⁺]_free reached upon UV-flash Ca²⁺ uncaging were verified to be similar in 310 mOsm (n = 7) and 390 mOsm (n = 5) conditions, 59 ± 7 µM and 64 ± 6 µM (p=0.6; Figure 4A-inset). In these latter conditions of intracellular hyperosmotic stress, the strength of exocytotic facilitation was maximal at 390 mOsm and abruptly decreased at 410 mOsm, a high pressure at which the intrinsic organization of the IHC exocytotic machinery may have been damaged. At 390 mOsm, the maximum amplitude of the exocytotic response was 1.5 ± 0.15 pF (n = 9) as compared to 1.0 ± 0.1 pF (n = 16) in controls at 310 mOsm (p<0.05; Figure 4A,C). The peak exocytotic rate was largely increased at 390 mOsm as compared to 310 mOsm (25.0 ± 4.5 pF/s (n = 9) and 13.7 ± 1.7 pF/s (n = 16), respectively; p<0.05, Figure 4D). Interestingly, the facilitation of exocytosis by high intracellular hydrostatic pressure was not observed in IHCs treated with latrunculin-A (Figure 2D, pink bars) and in Otof^-/- IHCs (Figure 3H and Figure 4B–D). The peak [Ca²⁺]_free reached upon UV-flash Ca²⁺ uncaging was unchanged in Otof^-/- IHCs as compared to control IHCs (Vincent et al., 2014).

Figure 4

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Overall, the facilitation of exocytosis by high intracellular hydrostatic pressure could be explained by an increased membrane tension that impacts on membrane fusion (Kozlov and Chernomordik, 2015) but also by an increased vesicular mobility, possibly as the result of reduced molecular crowding and loosened vesicle interactions, accelerating the replenishment rate. In the same way, the mobility of vesicles in pancreatic cells and primary hepatocytes was shown to be affected by hydrostatic pressure, a process related to molecular crowding and microfilaments polymerization (Nunes et al., 2015).

Do large changes in hydrostatic pressure occur in the cochlea during physiological or pathological conditions? In the cochlea, large intercompartmental osmotic gradients from 289 to 322 mOsmol/kg H₂0 are present between the perilymphatic and endolymphatic compartments, respectively (Sterkers et al., 1984). These osmotic gradients are likely regulated by aquaporins present in outer sulcus cells (Eckhard et al., 2015) and genetic deletion of aquaporin-4 in mice leads to impaired hearing (Li and Verkman, 2001). Osmolarity changes in these inner fluid compartments have long been suspected to be contributing factors to inner ear disorders such as tinnitus and fluctuating hearing loss, inclusive of Menière's disease (Angelborg et al., 1982). The outer hair cells, producing the electro-mechanical amplification of sound in the cochlea, have been shown to be mechanically sensitive to extracellular osmotic variation (Dulon, et al., 1987), a factor that greatly influence hearing (Choi and Oghalai, 2008). Our study, showing that IHC exocytosis is sensitive to osmotic forces, suggests that a defect in synaptic transmission at the auditory ribbon synapses may also underlie some hearing disorders associated with Menière's disease.

In conclusion, we propose that a synaptic F-actin network tightly controls the flow of synaptic vesicles during exocytosis at the IHC ribbons. This synaptic F-actin network also influences the sensitivity of exocytosis to hydrostatic pressure changes, presumably through a regulation of membrane tension at the active zones. Such actin-based regulation of intracellular hydrostatic pressure and membrane tension, recently shown in a variety of physiological processes such as in the secretory properties of neuroendocrine cells (de Wit, 2010; Gutiérrez and Gil, 2011), the mitotic cell rounding during cell division (Stewart et al., 2011), the photomechanical responses of photoreceptors (Hardie and Franze, 2012) to name a few, is here described for the first time in auditory hair cells.

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Author details

1. Philippe FY Vincent

   Bordeaux Neurocampus, Equipe Neurophysiologie de la Synapse Auditive, Université de Bordeaux, Bordeaux, France

   Contribution

   PFYV, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   philippe.vincent@inserm.fr

   Competing interests

   The authors declare that no competing interests exist.

2. Yohan Bouleau

   Bordeaux Neurocampus, Equipe Neurophysiologie de la Synapse Auditive, Université de Bordeaux, Bordeaux, France

   Contribution

   YB, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

3. Christine Petit

   1. Unité de Génétique et Physiologie de l’Audition, Institut Pasteur, Paris, France
   2. UMRS 1120, Institut National de la Santé et de la Recherche Médicale (INSERM), Paris, France
   3. Sorbonne Universités, UPMC Université Paris, Paris, France
   4. Syndrome de Usher et Autres Atteintes Rétino-Cochléaires, Institut de la Vision, Paris, France
   5. Collège de France, Paris, France

   Contribution

   CP, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

4. Didier Dulon

   1. Bordeaux Neurocampus, Equipe Neurophysiologie de la Synapse Auditive, Université de Bordeaux, Bordeaux, France
   2. UMRS 1120, Institut National de la Santé et de la Recherche Médicale (INSERM), Paris, France

   Contribution

   DD, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   didier.dulon@inserm.fr

   Competing interests

   The authors declare that no competing interests exist.

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