Wiring the Cochlea for Sound Perception
Abstract and Keywords
Sound information enters the auditory brainstem via spiral ganglion neurons (SGNs), which reliably encode and transmit information received from sensory hair cells in the cochlea. SGNs form both a structural and a functional bridge between the cochlea and the auditory brainstem beginning early in development, with long-term consequences for the maturation of central auditory circuits. This chapter summarizes the key events in SGN development, from their origin in the early otic vesicle and the emergence of the basic wiring pattern of the cochlea to the elaboration of specialized synapses and the acquisition of diverse firing properties that enable sound perception even within noisy environments. Key cellular events and molecular players are introduced, with emphasis on the impact of reciprocal interactions between the cochlea and the auditory brainstem throughout development.
Spiral ganglion neurons (SGNs) provide the primary conduit for sound information to enter the brainstem from the cochlea, allowing the brain to calculate what was heard and where it is located based on the spatiotemporal pattern of SGN activation. To fulfill this role, SGNs must develop several specialized properties that distinguish them from other primary sensory neurons. This includes formation of tonotopically organized connections in the cochlea and in the cochlear nucleus complex, elaboration of synapses that transmit with high fidelity, and acquisition of distinct firing properties that enable signaling across a wide dynamic range. Given their crucial position as the gateway between the periphery and the central nervous system (CNS), changes in the number, organization, or function of SGNs can have permanent consequences for how auditory information is subsequently represented in the brainstem and along the neuraxis. Efforts to understand how SGNs develop have established the basic sequence of events and begun to unravel the genetic networks that guide their specification and wiring.
Overview of SGN Development
SGNs develop from a pool of neurosensory progenitors that is progressively subdivided to generate all of the hair cells and neurons in the auditory and vestibular systems (Goodrich, 2016). In birds and rodents, neurosensory progenitors arise in the anteroventral quadrant of the otic cup (Adam et al., 1998; Cole et al., 2000; Morsli, Choo, Ryan, Johnson, & Wu, 1998), which is the embryonic anlage of the inner ear. As development proceeds, the otic cup invaginates and detaches from the hindbrain to form the otic vesicle (Anniko & Wikström, 1984; Hemond & Morest, 1991). Around the same time, neuroblasts begin to delaminate from the pro-neurosensory domain (PNSD) into the mesenchyme, where they aggregate to form a common cochlear-vestibular ganglion (CVG). The CVG is initially attached to the geniculate ganglion, which forms the seventh cranial nerve. In mice, the auditory and vestibular divisions gradually separate from each other (Carney & Silver, 1983; Sher, 1971), and the spiral ganglion grows and extends, due both to ongoing proliferation among the delaminated neuroblasts (Matei et al., 2005; Ruben, 1967) and convergent movements that lengthen the cochlea (J. Wang et al., 2005). Vestibular ganglion neuron (VGN) precursors exit the cell cycle first, followed shortly by an overlapping production of SGN precursors that differentiate along a gradient starting in the base of the cochlea and spiraling toward the apex (Ruben, 1967).
The earliest indication of SGN differentiation is the extension of peripheral and central processes, which eventually form topographically organized synapses in the cochlea and cochlear nucleus complex (CNC), respectively. Activity initiates in the embryonic cochlea and is propagated all the way along the auditory axis during postnatal stages, thus probably influencing the final maturation of auditory circuitry both in the cochlea and in the brainstem (H. C. Wang & Bergles, 2015). Indeed, removal of the cochlea during development has severe consequences for development of the CNC (Harris & Rubel, 2006). SGN development follows a similar progression across species, including humans (Bibas et al., 2006; Streeter, 1906), although the timing can differ. Much of our understanding derives from analysis of mice and other rodents, with additional important insights from chickens, which are more amenable to embryological manipulations.
Development of SGNs in the periphery overlaps temporally with development of olivocochlear neurons (OCNs) in the CNS. OCNs are a specialized population of efferent neurons that reside in the brainstem and provide reciprocal input to the cochlea (Bruce, Kingsley, Nichols, & Fritzsch, 1997; Fritzsch & Elliott, 2017). They comprise two broad subtypes: the lateral and medial olivocochlear neurons (LOCs and MOCs, respectively). LOCs are situated in the lateral superior olive (LSO) and extend largely ipsilateral projections that terminate close to the endings of Type I SGNs underneath the inner hair cells (IHCs). MOCs are located in the ventral nucleus of the trapezoid body (VNTB) and extend both ipsilateral and contralateral projections that directly innervate the outer hair cells (OHCs). Collectively, the olivocochlear efferent system may contribute to binaural processing and also protects the inner ear from noise damage (Darrow, Maison, & Liberman, 2006, 2007; Kujawa & Liberman, 1997). OCN axons arrive in the cochlea during embryonic stages and are closely associated with SGN axons as they innervate their peripheral targets. Hence, SGN development is temporally and spatially coordinated with similar developmental events in the brainstem.
Production of SGNs
As in other regions of the nervous system, SGNs are specified by the sequential action of familiar signaling pathways and networks of transcription factors. In mice, these events start around embryonic day 9 (E9), with an obvious CVG present rostral to the otocyst by E10.5 (Anniko & Wikström, 1984; Carney & Silver, 1983; Sher, 1971). VGNs are born around E8, followed by the SGNs from E9.5 until E13.5 (Matei et al., 2005; Ruben, 1967); staging in rats is similar but shifted by a day. Analogous events occur in fish (Haddon & Lewis, 1996; Riley, 2003) and chickens (Hemond & Morest, 1991). In all species examined, the first step is to set aside the PNSD, which occurs as part of the basic patterning of the early otic cup by signals from surrounding tissues, including the brainstem, notochord, mesenchyme, and overlying ectoderm (Bok, 2005; Bok et al., 2007; A. S. Brown & Epstein, 2011; Liang, Bok, & Wu, 2010; Riccomagno, Takada, & Epstein, 2005).
Within the PNSD, neuronal precursors are selected among a mixed pool that contains progenitors for both neurons and sensory cells, namely the hair cells (HCs) and support cells of the organ of Corti (Raft et al., 2007; Sapède, Dyballa, & Pujades, 2012; Satoh & Fekete, 2005). The decision to follow the neuronal fate depends on Notch pathway activity, which probably biases the outcome of cross-repressive interactions between the pro-neural Neurog1 transcription factor and the pro-sensory Atoh1 transcription factor (Bermingham et al., 1999; Kiernan, 2013; Ma, Anderson, & Fritzsch, 2000; Ma, Chen, del Barco Barrantes, de la Pompa, & Anderson, 1998). A complex and dynamic network of transcription factors, including Neurog1, Six1, Eya1, and Sox2, cooperate to induce neuronal fate and usher inner ear neuroblasts toward maturity (Evsen, Sugahara, Uchikawa, Kondoh, & Wu, 2013; Jahan, Pan, Elliott, & Fritzsch, 2015; E. Y. M. Wong, Ahmed, & Xu, 2013). Layered on top of this generic neurogenic network are additional transcription factors that determine which neuroblasts will populate the spiral versus the vestibular ganglion. This decision occurs early, prior to delamination (Bell et al., 2008; Koundakjian, Appler, & Goodrich, 2007; Sher, 1971). A major determinant of the auditory-vestibular fate decision is the transcription factor Lmx1a, which divides the PNSD into an Lmx1– vestibular division and an Lmx1a+ auditory division, where it inhibits the vestibular fate in nascent auditory neuroblasts (Koo et al., 2009; Nichols et al., 2008). SGNs are further subdivided into Type I and Type II subtypes, but no transcription factors have been directly implicated in this fate decision.
One of the key regulators of SGN development is the transcription factor Gata3. Gata3 is expressed from the earliest stages of inner ear development, initially throughout the otic placode (Lawoko-Kerali, Rivolta, & Holley, 2002; Lilleväli et al., 2006). Over time, expression becomes more limited to SGNs (Karis et al., 2001; Lawoko-Kerali et al., 2002; Lilleväli, Matilainen, Karis, & Salminen, 2004; Rivolta & Holley, 1998), although a few VGNs retain expression of Gata3 as well (Lu, Appler, Houseman, & Goodrich, 2011). Expression is maintained postnatally and into adulthood (Appler et al., 2013; Lu et al., 2011; Nishimura, Noda, & Dabdoub, 2017), raising the possibility that Gata3 influences multiple stages of SGN development and function. In support of this idea, different phenotypes were observed when Gata3 was removed from the entire otic vesicle or only from post-delaminated neuroblasts in mice. In Gata3 null mutants or after early conditional deletion, SGNs were absent altogether, but the VGN persisted (Duncan & Fritzsch, 2013; Duncan, Lim, Engel, & Fritzsch, 2011; Karis et al., 2001). After slightly later conditional deletions, SGNs were specified and delaminated, but were fewer in number (Duncan & Fritzsch, 2013). The mutant SGNs did not differentiate properly and underwent massive cell death beginning around E12.5 (Duncan & Fritzsch, 2013; Luo et al., 2013). Deletion using a neuronal-specific Cre driver circumvented these early phenotypes and revealed further roles for Gata3 in SGN differentiation (Appler et al., 2013), probably acting upstream of additional transcriptional effectors (Appler et al., 2013; Yu et al., 2013). For instance, upon later deletion, Gata3 mutant SGNs were present but differentiated prematurely and failed to form orderly radial bundles, although the peripheral axons still reached the organ of Corti. Subsequently, the mutant SGNs were lost, echoing the survival phenotypes observed after early deletion of Gata3. Thus, Gata3 is not only an early indicator of the SGN fate but also a major player in both the execution of this fate and subsequent survival.
Outgrowth and Targeting of SGN Peripheral Processes
In the cochlea, SGN peripheral processes are organized in a stereotypic manner that reflects the basic map of auditory information. Fibers are arranged within radial bundles that project like spokes of a wheel from the “modiolus” of the cochlea out toward the organ of Corti, responding to low sound frequencies in the apex and to high frequencies in the base. Type I SGN processes densely innervate the IHC region, with one neuron contacting one IHC (Figure 1). Type II SGNs instead turn toward the base and grow along rows of OHCs, forming synaptic contacts with multiple OHCs. In mice, the emergence of this striking pattern begins around E11.5, when SGNs first extend peripheral processes through the periotic mesenchyme and back toward the developing cochlear duct, arriving in the prosensory domain in the base of the cochlea around E14.5 (Carney & Silver, 1983; Koundakjian et al., 2007; Sandell, Butler Tjaden, Barlow, & Trainor, 2014). Differentiation progresses from the base to the apex, following the gradient of cell cycle exit. By E15.5, SGN peripheral processes lie in nascent radial bundles that line up beneath the IHCs along the length of the cochlea (Coate et al., 2012; Koundakjian et al., 2007).
Little is known about the cues that elicit peripheral process outgrowth during the early stages of cochlear wiring. Outgrowth does not require the presence of HCs (Elliott, Kersigo, Pan, Jahan, & Fritzsch, 2017; Fritzsch et al., 2005), indicating that a classic target-derived mechanism is unlikely to be at work. Instead, SGN peripheral processes may be steered locally by surrounding non-neuronal cells. During the earliest stages of axon outgrowth, young SGNs are surrounded by a “funnel” of mesoderm and glia (Carney & Silver, 1983). Moreover, the SGN peripheral processes are systematically interdigitated with neural crest-derived glia that could form a permissive corridor for outgrowth (Sandell et al., 2014), as occurs in other regions of the nervous system (Freter, Fleenor, Freter, Liu, & Begbie, 2013). Indeed, SGN neurites grow robustly along Schwann cells in vitro (Jeon, Xu, Xu, & Hansen, 2011; Whitlon, Tieu, Grover, Reilly, & Coulson, 2009). More strikingly, SGNs are misplaced and fail to form orderly radial bundles in mice with abnormal glial invasion (Mao, Reiprich, Wegner, & Fritzsch, 2014; Morris et al., 2006). Bundle formation also depends on Ephrin/Eph dependent interactions between SGNs and the surrounding mesenchymal cells (Coate et al., 2012). Neurotrophins are likely to contribute to these early stages (Yang, Kersigo, Jahan, Pan, & Fritzsch, 2011), but it can be difficult to distinguish effects on outgrowth from those on survival, since the final phenotype is reduced innervation in both cases. Overall, although many guidance cues are expressed in the developing inner ear and able to influence neurite outgrowth in vitro, specific roles remain to be defined in vivo (Bank et al., 2013; Battisti, Fantetti, Moyers, & Fekete, 2014; Battisti & Fekete, 2008; Bianchi & Gale, 1998; Bianchi & Gray, 2002; Fantetti & Fekete, 2012; Fekete & Campero, 2007).
During the next stage of cochlear wiring, SGN peripheral processes target appropriate synaptic partners: the IHCs for Type I SGNs and the OHCs for Type II SGNs. In both cases, synaptic communication occurs through glutamate-induced activation of AMPA receptors (AMPARs) in SGNs. In the absence of early molecular markers, studies have relied on morphological distinctions, namely radial versus spiraling fibers, to identify these two subtypes. Based on this criterion, the gross afferent innervation pattern at E16.5 already bears strong resemblance to that seen at maturity in mice, with most SGN processes innervating IHCs and a minority projecting to OHCs (Coate, Spita, Zhang, Isgrig, & Kelley, 2015; Koundakjian et al., 2007). That these patterns are seen before HCs are fully differentiated suggests that the Type I and II identities are established independent of interactions with their peripheral synaptic targets. Acquisition of SGN identities might be under the control of familiar transcriptional networks, as mature Type II SGNs express higher levels of nuclear-localized Gata3 and Mafb, complementing Prox1 expression in mature Type I SGNs (Nishimura et al., 2017). Enrichment of Gata3 in a subset of SGNs was detected as early as E17.5 in mice, suggesting that the Type I/II decision must occur then or earlier. Type I and Type II SGNs can eventually be identified by differential expression of EphA4 in Type I SGNs (Defourny et al., 2013) and Peripherin and Tyrosine hydroxylase in Type II SGNs (Hafidi, 1998; Hafidi, Després, & Romand, 1993; Vyas, Wu, Zimmerman, Fuchs, & Glowatzki, 2017), thus underscoring the likelihood that transcription factors such as Gata3, Mafb, and Prox1 may directly control aspects of Type I/II identity.
Segregation of Type I and II afferents to distinct domains of the sensory epithelium is mediated by repulsive interactions that keep Type I fibers away from the OHC zone while permitting entry of Type II fibers (Druckenbrod & Goodrich, 2015). Such interactions are mediated in part by two molecular pathways. On the one hand, Ephrin A5 in the OHC zone repels Type I fibers, all of which express the receptor Epha4 (Defourny et al., 2013). On the other, the secreted chemorepellant Sema3a is expressed in the OHC zone and repels Type I SGNs, which express the cognate Neuropilin receptor Nrp2. How Type II SGNs, which also express Nrp2, manage to grow into the Sema3a zone remains unclear, but it may involve a co-receptor (Coate et al., 2015). Disruption of either cue results in increased invasion of the OHC area by Type I fibers. However, most Type I SGNs are still confined to the IHC zone in single mutants, suggesting that multiple mechanisms underlie selective targeting of SGN afferents. Indeed, EphB receptor signaling is required to keep SGN processes from growing even further, past the OHC region, probably via an EphrinB1 ligand (Zhou, Lee, Henkemeyer, & Lee, 2011). Studies so far have not identified the attractive guidance cues that may direct Type II axons to turn toward the base.
Like other nascent neural circuits, target innervation by SGN afferents is initially imprecise, with a fraction of Type I afferents invading the OHC area during late embryonic and early postnatal stages (Figure 1). Why ectopic branches are present and how this relates to synapse formation is not fully understood. Classic studies of Golgi-stained processes in the apex of the cochlea suggested that individual fibers branch exuberantly and contact both IHCs and OHCs, followed by a period of refinement (Echteler, 1992; D. D. Simmons, 1994). Likewise, putative Type I fibers, identified based on selective dye uptake, were observed among rows of OHCs (L.-C. Huang, Thorne, Housley, & Montgomery, 2007). Consistent with this interpretation, the putative Type I fibers found near OHCs during early postnatal stages exhibit synaptic AMPAR profiles that match those found at Type I-IHC synapses in mature animals (L.-C. Huang et al., 2012). However, live-imaging of SGNs ex vivo revealed that excess branches from Type I fibers are not positioned to form synapses with the OHCs, arguing against the notion that the IHC-like synapses on OHCs are made by Type I fibers that overextend into the OHC zone (Druckenbrod & Goodrich, 2015). In addition, both AMPAR-mediated current (by pharmacology) and the GluA2 AMPAR subunit (by immunohistochemistry) can be detected at OHC-Type II synapses (Martinez-Monedero et al., 2016), raising questions as to whether GluA2-containing OHC synapses observed at early postnatal stages (P0-P6) are formed by Type II or Type I fibers.
Nevertheless, there is broad agreement that Type I fibers that invade the OHC area undergo retraction, resulting ultimately in subtype-specific confinement to distinct spatial domains of the sensory epithelium. Reconstruction of individual radial and spiral fibers during postnatal stages confirmed a period of extensive branching followed by pruning, but these events appear to be confined to putative Type I SGNs (Druckenbrod & Goodrich, 2015). Among Type I fibers that terminate in the IHC zone, most are branched and innervate multiple IHCs or establish multiple contacts with the same IHC at neonatal stages, as well as extending rare branches to contact pillar cells and the first row of OHCs. All but one of these processes are pruned, resulting in strict mono-innervation of IHC by each SGN. Factors regulating this pruning process are currently unknown. Importantly, Type I SGNs seem to be able to form synapses with OHCs if given the opportunity, as occurs in EphA4 mutant mice (Defourny et al., 2013). Thus, cochlear wiring specificity may arise through guidance mechanisms rather than being dictated by a molecular recognition code, as occurs in other sensory systems such as the retina (Sanes & Yamagata, 2009).
Formation of Peripheral Synapses
Faithful encoding of acoustic information necessitates high fidelity of synaptic transfer at the IHC-SGN synapse. Reliable signal transmission is achieved in part via specializations such as the presynaptic ribbon, an electron-dense structure consisting of glutamate-filled vesicles arranged in a ring-like pattern and anchored at the active zone. Each ribbon is apposed by an SGN terminal ending, which forms a large post-synaptic density rich with AMPARs (Yu & Goodrich, 2014). The formation and maturation of synaptic connections in the cochlea occur over a prolonged period, starting at neonatal stages and lasting until about the fourth postnatal week in mice (Figure 2). The earliest events seem to initiate independently in HCs and SGNs. Subsequently, interactions between pre- and post-synaptic partners promote localization and maturation of the synapses, as well as the final synapse count. In the mature cochlea, the sizes of individual synapses on the modiolar and pillar sides of the IHC differ such that SGNs with different physiological properties receive acoustic information through morphologically distinct synaptic connections.
The onset of synapse development begins with the production of necessary presynaptic components in the IHCs, which produce immature ribbons even in the absence of SGN innervation (Sobkowicz, Rose, Scott, & Levenick, 1986). These events happen quite early, probably during late embryogenesis in rodents (Lenoir, Shnerson, & Pujol, 1980; Shnerson, Devigne, & Pujol, 1981). Synapse development is similar in OHCs, but slightly delayed (Sobkowicz et al., 1986). The ribbons are initially dispersed in the cytoplasm and become gradually localized to the basal pole of the IHC (Sendin, Bulankina, Riedel, & Moser, 2007; Shnerson et al., 1981; Sobkowicz et al., 1986), where they are anchored to the membrane by the scaffolding protein Bassoon (Jing et al., 2013), probably via interactions with the ribbon protein RIBEYE (Dieck et al., 2005). IHCs are capable of evoked exocytosis by late embryogenesis (Beutner & Moser, 2001; Johnson, Marcotti, & Kros, 2005). On the post-synaptic side, AMPARs are detectable in SGN endings at birth and then increase in expression during the first postnatal week (L.-C. Huang et al., 2012). Postsynaptic differentiation also lags behind morphologically, as a well-defined electron-dense PSD appears after the ribbons have already begun to be anchored to the active zone (Sobkowicz et al., 1986). Development of the PSD depends on the transcription factor Mafb, which is expressed in SGNs (but not in VGNs) beginning around E16.5 in mice (Yu et al., 2013). In Mafb mouse mutants, SGN peripheral processes reach the IHCs, but do not develop normal PSDs, evidenced by morphological changes and loss of glutamate receptor (GluR) immunoreactivity. Formation of contacts between IHCs and SGNs may also involve the Ig superfamily protein Neuroplastin, which is localized to the base of maturing IHCs and is required for cochlear synaptogenesis and hearing in mice (Carrott et al., 2016).
Over the first postnatal week, the pattern of synaptic connectivity changes significantly, both in terms of the morphology and number of IHC-SGN contacts. Immature synapses feature multiple ribbons and diffuse GluR patches, whereas each mature synapse contains a single elongated ribbon at the active zone apposed to an electron-dense PSD with a well-defined GluR patch (Figure 2; Sendin et al., 2007; Sobkowicz et al., 1986; A. B. Wong et al., 2013). Even at these early stages, HCs can signal to the SGNs (Beutner and Moser, 2001; Johnson et al., 2005; Tritsch et al., 2010), indicating that these synapses are functional. However, exocytosis is inefficient initially, probably reflecting the relative immaturity of the Ca2+ signaling system in the IHCs (Beutner & Moser, 2001; Johnson et al., 2005). During maturation, the number of ribbons per hair cell decreases (L.-C. Huang et al., 2012; Nemzou, Bulankina, Khimich, Giese, & Moser, 2006; Sendin et al., 2007; Sobkowicz, Rose, Scott, & Slapnick, 1982; Sobkowicz et al., 1986), with a corresponding increase in association between presynaptic ribbons and postsynaptic GluR patches, which is seen robustly starting around P10 in mice (L. D. Liberman & Liberman, 2016). At the same time, the ribbons fuse with each other and assume a more elongated morphology (Sendin et al., 2007; Sobkowicz et al., 1982). In addition, the overall number of synapses (i.e., ribbons co-localized with GluR patches) decreases by about 50% through development (L.-C. Huang et al., 2012; Nemzou et al., 2006; Sendin et al., 2007; Sobkowicz et al., 1982, 1986). These changes are particularly evident in the OHC region, with most mature OHCs retaining only one to three ribbons. Thus, the overall ribbon number can be influenced by differences in the initial formation of synapses, pruning of excess synapses, or loss of Type II SGNs during a wave of cell death postnatally (Barclay, Ryan, & Housley, 2011).
Not surprisingly, the formation of properly aligned and functional synaptic contacts involves coordination between HCs and SGNs (Sheets et al., 2011; Sobkowicz et al., 1986). For instance, disruption of post-synaptic differentiation upon loss of MafB from SGNs also results in dysmorphic presynaptic ribbons in mouse HCs (Yu et al., 2013). More broadly, the overall number of synapses decreases and the number of orphan ribbons is increased at P12. Conversely, premature overexpression of Mafb accelerated the localization of presynaptic ribbons to the basal pole of the IHCs. This Mafb-dependent coordination of IHC-SGN synapse development appears to proceed as part of a broader SGN-specific differentiation program orchestrated by the master regulator Gata3.
Following the formation of functional synaptic contacts, the cochlea undergoes an additional phase of maturation during which pre- and postsynaptic structures exhibit dynamic changes before acquiring heterogeneous morphologies that reflect Type I SGN identities. Type I SGNs in mammals are classified as low-SR, medium-SR, and high-SR subtypes based on their spontaneous firing rates (discussed further in the section on “Reciprocal interactions between the cochlea and the brainstem” and covered in the chapter by Fuchs, this volume). In the mature cochlea, these subtypes exhibit spatially ordered heterogeneity in synapse morphology along the basal surface of the IHCs (Figure 2). Because the cochlea spirals, the IHC axis is generally described based on the order of cells within the duct, with the “modiolar” side closer to the SGN cell bodies and the “pillar” side closer to the pillar cells (Figures 1 and 2). At each IHC, the SGN processes that terminate on the modiolar face (low- and medium-SR) have larger presynaptic ribbons and small postsynaptic GluR patches, while those on the pillar face (high-SR) have small ribbons and large GluR patches (L. D. Liberman, Wang, & Liberman, 2011). How these morphological features affect synaptic function, particularly in the context of known afferent physiological diversity, remains unknown. In stark contrast to the prolonged period over which SGN ion channel profiles mature postnatally, heterogeneity in ribbon size is already apparent by P3 in rats (Kalluri & Monges-Hernandez, 2017). However, ribbon morphologies appear to be dynamic throughout the first few weeks of development: the adult-like size gradient that is visible shortly after birth is lost by hearing onset, spatially reversed in the third postnatal week, and then reestablished sometime in the fourth week (Kalluri & Monges-Hernandez, 2017; L. D. Liberman & Liberman, 2016).
SGNs also undergo developmental changes in glutamate receptor composition that alter how they respond to glutamate released from HCs. Although all four AMPAR subunits are detected in SGNs, their expression patterns vary during development (Eybalin, Caicedo, Renard, Ruel, & Puel, 2004). GluA1 is expressed soon after birth but downregulates throughout the first week and is lost after hearing onset. On the other hand, GluA2 is undetectable early on but upregulates toward the end of the first postnatal week, with clear apposition of GluA2 puncta against presynaptic ribbons apparent before hearing onset (Eybalin et al., 2004; L. D. Liberman & Liberman, 2016). Both GluA3 and GluA4 are expressed at all stages, but GluA3 levels progressively decrease beyond the fourth postnatal week. The developmental switch from GluA1- to GluA2-mediated synaptic transmission around hearing onset may hold significance for Ca2+ permeability of AMPARs and plasticity (Cull-Candy, Kelly, & Farrant, 2006). Furthermore, it is unknown whether GluA subunits in SGNs exhibit a developmental switch in regulation at the mRNA and protein levels, for instance, via RNA editing, splicing, and phosphorylation, as occurs elsewhere in the CNS (Isaac, Ashby, & McBain, 2007). NMDARs, which shape glutamate-mediated SGN excitation in the neonatal cochlea and influence their spontaneous firing behavior (Zhang-Hooks, Agarwal, Mishina, & Bergles, 2016), also exhibit a developmental switch: GluN1 and GluN2A are expressed in the prehearing cochlea, but GluN1 is downregulated and GluN2A is lost while GluN2B, GluN2C, and GluN2D are upregulated with maturation (Knipper et al., 1997; Sanchez, Ghelani, & Otto-Meyer, 2015). Notably, this transition from a predominantly GluN2A- to GluN2B-containing NMDA receptor profile among SGNs is opposite of the developmental switch that occurs in the CNS (Yashiro & Philpot, 2008), including the auditory brainstem (Sanchez, Wang, Rubel, & Barria, 2010).
In summary, SGNs begin to receive synaptic signals from HCs perinatally while both the cochlea and brainstem are still maturing. The nature of synaptic transmission between the IHCs and SGNs changes as the system develops and matures. Since SGNs can evoke action potentials in post-synaptic targets in the brainstem as early as E15 in mice (Marrs & Spirou, 2012), developmental changes in the pattern of activity in the cochlea could influence the development of circuits in the brainstem.
Development of Central Axons
Acoustic information encoded by SGNs is relayed to a diverse array of neurons in the auditory brainstem that subserve circuits underlying detection of distinct sound features. All SGNs exit the cochlea via the eighth cranial nerve and then bifurcate after they enter the CNC, sending an ascending projection to the anterior ventral cochlear nucleus (aVCN) and a descending projection to the posterior VCN (pVCN) that ultimately terminates in the dorsal cochlear nucleus (DCN; Figure 3A). In mice, SGN axons reach the brainstem by E11.5 and bifurcations are obvious one day later (Lu et al., 2011). Thus, the cochlea is connected to the developing brainstem even before all of the SGNs have become post-mitotic. Likewise, neurons in the CNC are produced between E10.5 and E15.5 in mice (Martin & Rickets, 1981), overlapping with eighth nerve development. This has important implications for circuit assembly, since the arrival of SGN axons marks the first time that tonotopic order in the cochlea is communicated to the CNS.
The nascent CNC spans rhombomeres 2 to 5 (R2 to R5) in the brainstem (Cramer, Fraser, & Rubel, 2000; Di Bonito & Studer, 2017; Farago, Awatramani, & Dymecki, 2006), so the first step for SGN axons is to target the correct rostro-caudal position. This might be aided by the presence of a stream of neural crest cells emanating from R4 (Sandell et al., 2014). In support of this idea, the eighth nerve is malformed after ablation of neural crest cells in chick embryos. There is also a band of GFAP-positive cells that is well-positioned to demarcate where 8th nerve fibers should enter (Siddiqui & Cramer, 2005). As expected based on their birth dates, the vestibular axons arrive slightly before the auditory axons, by E10.5 in mice, at about the same time that facial motor neurons have extended axons out through the 7th nerve, which develops in close proximity (Fritzsch & Nichols, 1993). Transplant studies in Xenopus suggest that cranial nerve development might be somewhat promiscuous. For instance, motor neurons are able to innervate an ectopic inner ear in the trunk. Conversely, VGNs send projections back to the spinal cord, apparently growing along whichever nerves are available (Elliott & Fritzsch, 2010; Elliott, Houston, & Fritzsch, 2013). Similarly, in mice, SGNs still send central axons into the vicinity of the CNC even when most CNC neurons are missing (Maricich et al., 2009). Thus, it is possible that SGN central axons simply follow the path available to them rather than responding to specific cues, with some contributions from permissive cues at the nerve entry site.
The eighth nerve contains a variety of fibers that are organized in a systematic fashion. For instance, the auditory and vestibular branches of the nerve are segregated from each other, and auditory fibers are further organized tonotopically (Sando, 1965). Axon-axon interactions within and across these different branches may contribute both to the guidance of central axons to the brainstem and the organization of the axons within the nerve. In other regions of the nervous system, such orderly arrangement within nerves and tracts is mediated by selective fasciculation (Sitko and Mason, 2015). Chicken auditory and vestibular axons express different complements of Eph receptors and ligands, but no role in segregation has been confirmed (Siddiqui & Cramer, 2005). However, a disorganized nerve forms when Pax2-cre is used to abolish Neurod1 expression, with intermingling of auditory and vestibular axons that project to inappropriate targets in the hindbrain (Jahan, Kersigo, Pan, & Fritzsch, 2010). Whether this is a primary effect on the guidance behavior of the neurons remains to be determined.
In the auditory system, sound frequency is represented as a place code in the cochlea and relayed via topographically organized projections to higher auditory centers. Tonotopy starts in the cochlea, with HCs and SGNs that respond to high sound frequencies located in the base and those for low frequencies in the apex. The same order is preserved in the 8th nerve, with SGN axons from high frequency regions segregated from those for low frequency regions (Muniak et al., 2013; Sando, 1965). The axons go on to terminate in a cochleotopic manner in all three divisions of the CNC, a pattern that is reiterated along the auditory axis (Kandler, Clause, & Noh, 2009). Topographic maps exist throughout the nervous system and form through a combination of hard-wired and activity-dependent mechanisms, with crucial roles for Ephrin/Eph signaling during the first phase and acetylcholine-driven waves of spontaneous activity in the second (Cang & Feldheim, 2013). In the auditory system, SGN axons are topographically organized before they form synaptic connections either in the periphery or in the CNS (Koundakjian et al., 2007; Molea & Rubel, 2003). For instance, as early as E15 in mice, SGN axons from the apex innervate a restricted region of the developing CNC, with no signs of overshooting that are typical in other sensory systems (Koundakjian et al., 2007). Likewise, SGN axons show a coarse topographic organization in the CNC of newborn kittens (Leake, Hradek, Chair, & Snyder, 2006). It is unclear whether Eph receptors influence topography within the auditory nerve as they do in other regions of the nervous system, but some family members are expressed in gradients within the nerve (Siddiqui & Cramer, 2005) and seem to play a role in the distribution of 8th nerve fibers that enter the hindbrain in chickens (Allen-Sharpley, Tjia, & Cramer, 2013). In addition, reduction of EphrinB2, which is expressed uniformly among SGN axons and in the CNC, leads to slightly blurred tonotopy in DCN (Miko, Nakamura, Henkemeyer, & Cramer, 2007).
Although initial development of SGN-CNC connections may be hard-wired, the maps become more precise with age, possibly due to activity-dependent mechanisms. For instance, the broad maps seen in the newborn kitten are more adult-like by P6 (Snyder & Leake, 1997), but remain coarse if the animals are deafened at birth (Leake et al., 2006). This period of refinement corresponds to a phase during which rhythmic bursts of spontaneous activity occur among SGNs in neonatal cats (Jones, Leake, Snyder, Stakhovskaya, & Bonham, 2007). However, it is unclear whether spontaneous activity was actually disrupted by deafening, which was achieved by neomycin-induced HC death. Additionally, complete removal of the cochlea during early postnatal life reduces the number of neurons in the CNC (Harris & Rubel, 2006): many neurons die in the aVCN, causing reduction in size of the nucleus (Figure 3B; Mostafapour, Cochran, Del Puerto, & Rubel, 2000), and there is a loss of Wnt-sensitive progenitors that would otherwise produce neurons during this time frame (Volkenstein et al., 2013). Mass ablation of HCs from neonatal (but not mature) mice had the same effect (Tong et al., 2015). Thus, it is unclear which changes drive the observed degradation of tonotopic organization seen in deafened cats.
Analysis of a mouse mutant with an unusually specific effect on sensory axon development (Lu et al., 2014) hints that SGNs can in fact influence the formation of tonotopically ordered circuits within the CNC. SGN axons exhibit a stereotyped bifurcation that enables distribution of auditory information to all three divisions of the CNC. Bifurcation differs from axon branching in that the primary axon gives rise to two equal axons that go on to navigate independently (Schmidt & Rathjen, 2010). SGN bifurcation (Lu et al., 2014) relies on the same pathway used by sensory neurons in the dorsal root ganglion (DRG), which bifurcate in a similar fashion upon reaching the spinal cord (Schmidt et al., 2002, 2007). Like DRG neurons, SGNs express the natriuretic peptide receptor 2 (Npr2) during the time of bifurcation, which fails to occur in Npr2 mutant mice (Lu et al., 2014). The Npr2 ligand, c-type natriuretic peptide (CNP), is detectable in mice by E9.5 exactly where the SGN axons enter (Ter-Avetisyan, Rathjen, & Schmidt, 2014), pointing to a model where the localized bifurcation of SGN axons is controlled by CNP-Npr2 interactions. In Npr2 mutants, peripheral wiring and myelination appear normal, as is the tonotopic organization of SGN axons within the eighth nerve (Lu et al., 2014). However, the unbifurcated axons preferentially turn caudally and tonotopic organization within the CNC becomes blurred (Figure 3C). Whether these guidance defects are mediated by Npr2 or are instead secondary to the failure of bifurcation is unclear, as Npr2 mutant axons also elaborate collateral branches that can innervate multiple divisions of the CNC and hence further degrade tonotopy. Notably, the dramatic change in the pattern of SGN innervation is accompanied by changes in the organization of circuitry within the CNC. Tuberculoventral (TV) cells normally form tonotopically organized connections that link DCN to VCN, and in Npr2 mutants, this organization is less precise, mirroring effects on the SGN afferents (Lu et al., 2014). Although it remains to be proven that Npr2 acts solely in SGNs, this phenotype underscores the influential role that SGN afferents may play in the organization of central auditory circuits, starting at the very first site of processing.
Targeting and Synaptogenesis in the Cochlear Nucleus Complex
When SGN axons reach each target region within the CNC, they face the challenge of forming a variety of synapses with multiple post-synaptic partners. A particularly important step is the formation of unusually large calyceal synapses, which are crucial for the sound localization pathway. At a broad level, proper targeting and synaptogenesis of SGN afferents in the CNC are critical events that enable parcellation of neural signal onto distinct sub-circuits and extraction of diverse acoustic features.
A fundamental difference in the central innervation patterns of Type I and Type II neurons is that they primarily innervate cells in the magnocellular core and the granule cell domain (GCD), respectively (Figure 3A; Berglund & Brown, 1994; M. C. Brown, Berglund, Kiang, & Ryugo, 1988). Within the GCD, Type II fibers establish synaptic contacts mainly with small cells, and to some extent with other principal cell types, particularly the multipolar cells. Notably, the GCD also receives descending input from the auditory cortex (Weedman & Ryugo, 1996), inferior colliculus (Caicedo & Herbert, 1993), and superior olivary nucleus (Ryan, Schwartz, Helfert, Keithley, & Wang, 1987), as well as non-auditory sources (Burian & Gstoettner, 1988; Kevetter & Perachio, 1989; Ohlrogge, Doucet, & Ryugo, 2001; Zhan, Pongstaporn, & Ryugo, 2006; Zhan & Ryugo, 2007) such as the vestibular and somatosensory pathways (Haenggeli, Pongstaporn, Doucet, & Ryugo, 2005; Itoh et al., 1987; Weinberg & Rustioni, 1987; D. D. Wright & Ryugo, 1996). Thus, there is likely multimodal and top-down influence on the neurons targeted by Type II fibers.
Type I fibers exhibit systematic heterogeneity in their central innervation at multiple levels. Most Type I fibers terminate in the magnocellular core of the aVCN and pVCN, and the deep layer of the DCN (Fekete, Rouiller, Liberman, & Ryugo, 1984; M. C. Liberman, 1991, 1993; Ryugo & May, 1993). However, the pattern of innervation differs among the three physiologically defined SR groups, both in terms of the CN subdivisions they innervate and the cell types they contact. Low- and medium-SR fibers provide the sole source of input to the small cell cap (SCC), a thin sheet of small cells at the peripheral margin of the VCN (Ryugo, 2008). Within the SCC, low-SR collaterals ramify extensively, mostly within the neuropil, and likely contact multiple cells. Since cells in the SCC project to MOCs (Ye, Machado, & Kim, 2000), the selective innervation of SCC by low-SR SGNs, which have high thresholds, is thought to be part of a feedback circuit triggered by high sound intensities (Ryugo, 2008). On the other hand, all three SR subtypes project to the aVCN and pVCN, where they synapse with both common and distinct postsynaptic cell types (M. C. Liberman, 1991, 1993). Stellate cells in the aVCN receive input from all SR groups, but multipolar cells are contacted by only low- and medium-SR fibers (M. C. Liberman, 1993). In the pVCN, granule cells are innervated predominantly by high-SR fibers and multipolar cells are innervated by all SR groups. In the DCN, high-SR inputs generally form somatic contacts onto small cells in the deep layer, whereas medium-/low-SR inputs are non-somatic and occur closer to the fusiform cell layer (M. C. Liberman, 1993). That SR subtypes form connections that vary both by the identity of the postsynaptic cell and the subcellular location of the synapse suggests that the Type I SGNs collectively relay acoustic information to distinct circuit motifs within the lower auditory pathway. Although the number and nature of the synapses that link SGNs to the CNS determine how acoustic information is initially encoded in the brain, nothing is known about how the correct pattern is established in development.
Across all SR groups, Type I SGNs are tasked with faithfully conveying the complex temporal properties of sound up the auditory pathway. Fidelity is achieved in part via huge and complex synaptic structures, called the endbulbs of Held (Figure 3D), which are found at Type I SGN terminal endings on to globular and spherical bushy cells (SBCs) in the aVCN and are critical for sound localization. They occur as both axodendritic and axosomatic synapses, feature hundreds of presynaptic active zones (Ryugo, Wu, & Pongstaporn, 1996), and are powerful drivers of postsynaptic firing, especially considering that up to three endbulbs contact a single SBC (Brawer & Morest, 1975; Cant & Morest, 1979; Ryugo & Sento, 1991). The endbulbs release glutamate that is detected by the SBCs via both AMPA and NMDA-type receptors (Pliss, Yang, & Xu-Friedman, 2009; Y. X. Wang, Wenthold, Ottersen, & Petralia, 1998). Endbulb morphologies vary extensively between low- and high-SR SGNs with systematic differences in the complexity of the presynaptic terminal, which are accompanied by variations in PSD size and shape (Ryugo & Sento, 1991; Ryugo et al., 1996).
Synapse formation begins during the first postnatal week, when SGN axons contact SBCs and form relatively simple terminals. Over the next few days, these terminals grow much larger and begin to extend thin branches locally. Growth continues for many more weeks, as the terminal gradually envelops most of the SBC cell body and becomes ever more elaborate (Ryugo, Montey, Wright, Bennett, & Pongstaporn, 2006). Why SGN axons form endbulbs only upon contacting SBCs is not known. SGN axons are able to form functionally normal synaptic connections in Npr2 mutant mice despite the lack of tonotopic order, indicating that synaptic maturation may be controlled by local cell-cell interactions (Lu et al., 2014). The characteristic growth of the endbulbs, on the other hand, is influenced by activity. The role of activity is evident both in the natural variation in endbulb complexity that occurs among SGNs with different spontaneous firing rates (Ryugo & May, 1993), and in the loss of this complexity in congenitally deaf animals (Figure 3D; Baker, Montey, Pongstaporn, & Ryugo, 2010). Instead of well-defined and dome-shaped PSDs, deaf cats have significantly enlarged and flattened PSDs. Some aspects of this phenotype, such as enlargement of PSDs and increase in presynaptic vesicle density, may indicate homeostatic plasticity compensating for reduced activity. Consistent with this idea, transmitter release probability is elevated at the endbulb of Held-SBC synapse in deafness (dn/dn) mutant mice (Oleskevich & Walmsley, 2002), and reintroduction of activity via cochlear implants at three months of age corrects the synaptic morphology defect (O’Neil, Limb, Baker, & Ryugo, 2010). However, rescue does not occur when activity is restored at six months of age, which suggests a critical period during which synaptic morphology remains plastic with respect to activity in the auditory pathway. Little is known about how SGNs select the correct complement of post-synaptic partners and develop the morphologically appropriate type of synapse.
Reciprocal Interactions between the Cochlea and the Brainstem
SGNs provide a direct link between the periphery and the central auditory system that serves as both a physical and an electrical substrate for communication. Physically, the SGN axons offer a roadway for OCN axons into the cochlea. The OCNs, in turn, have important effects on the maturation of HCs and connectivity within the cochlea. Electrically, SGNs convey patterns of activity from the cochlea to the central nervous system, likely further refining tonotopic maps.
OCNs develop together with more canonical motor neurons in the ventral hindbrain (Fritzsch & Elliott, 2017). Like other motor neurons, they are primarily cholinergic, although a subset produce other transmitters and peptides such as GABA and CGRP (Eybalin, 1993; Sewell, 2011). OCNs share a particularly close relationship with the facial branchial motor neurons (FMBNs), which send axons into the 7th nerve. Both populations arise mostly in the fourth rhombomere and then undergo extensive migrations, leaving their axons behind them (Fritzsch & Elliott, 2017; Han, Gupta, & Novitch, 2018). As a result, OCN efferent axons follow along the genu of the 7th nerve. Upon exiting the brainstem, the OCN efferent axons diverge from the FBMN axons and reach the developing otocyst by E11.5 in mice (Bruce et al., 1997). At this stage, the earliest inner ear neuron axons, mostly vestibular, have already reached the brainstem, raising the possibility that the OCN efferents grow along the pre-existing inner ear afferents to reach the ear. There is broad agreement that SGN peripheral processes provide a scaffold for OC efferent growth inside the cochlea, but the idea has not been directly tested experimentally, nor is the molecular mechanism known (Fritzsch, 1996; Simmons, 2002). Nonetheless, in a wide variety of mouse mutants with disorganized SGN peripheral processes, OCN efferents are likewise disorganized (Duncan and Fritzsch, 2013; Duncan et al., 2011; Huang et al., 2001; Ma et al., 2000). Thus, the early extension of SGN peripheral and central processes may enable the subsequent guidance of OCN efferents into the inner ear.
The arrival of OCN efferents initiates reciprocal interactions that influence cochlear maturation and function. The most obvious effect comes from the MOC population. Upon first arriving in the cochlea, the MOC efferents elaborate transient branches that innervate the IHCs, despite the fact that in the mature cochlea, MOC efferents only innervate the OHCs (Glowatzki & Fuchs, 2000; Goutman, Fuchs, & Glowatzki, 2005). This contact may influence IHC maturation (Johnson et al., 2013). Less is known about the impact of LOC efferents, which are harder to study because the axons are thinner and unmyelinated. However, there are indications that input from the LOC neurons is poised to contribute to the different spontaneous firing rates among Type I SGNs (M. C. Liberman, 1980a, 1980b). In support of this idea, severing of the olivocochlear bundle prevents the maintenance of the mature pattern of synaptic heterogeneity associated with SR subtypes (Yin, Liberman, Maison, & Liberman, 2014). With no way to access LOC neurons selectively prior to birth, it is unclear whether they also affect the initial establishment of this pattern (M. C. Liberman, O’Grady, Dodds, Mcgee, & Walsh, 2000). However, analysis of a mouse mutant with early disruption of the efferent system prevented normal emergence of the synaptic gradient during early postnatal life (Hickman, Liberman, & Jacob, 2015). Overall, the olivocochlear system is well-positioned to influence the pattern of activity that emanates from the cochlea during development, either through effects on the HCs or on the SGNs (Johnson et al., 2011, 2013). Indeed, when cholinergic signaling is disrupted during development, the temporal pattern of activity is altered, with long-term consequences for the refinement of tonotopic projections in the lateral superior olive (Clause et al., 2014).
As well as providing a structural bridge between the PNS and CNS, SGNs directly inform the hindbrain about levels of activity in the cochlea, information that could influence assembly of circuits all the way up to the cortex. This activity begins quite early in auditory circuit development, as SGNs are spontaneously active by E14 in mice (Marrs & Spirou, 2012). By E15, stimulation of the auditory nerve can generate responses in the developing VCN, although these responses are slow and small. Thus, activity in the SGNs could influence the CNC even before synapses have formed in the periphery. In the mouse cochlea, IHCs first generate action potentials around E17 (Johnson et al., 2005; Marcotti, Johnson, Holley, & Kros, 2003), about the same time that mechanosensitivity can first be detected (Géléoc & Holt, 2003). The IHCs are functionally connected to the SGNs by birth, although the synapses are still immature (Shnerson et al., 1981; Tritsch et al., 2010). During these early stages, IHCs are activated by waves of ATP and potassium ions released from inner supporting cells (ISC) in Kolliker’s organ, a transient structure adjacent to the developing organ of Corti (Tritsch, Yi, Gale, Glowatzki, & Bergles, 2007; H. C. Wang et al., 2015). Bursts of action potentials in IHCs induce similar bursts of activity in SGNs and indeed along the auditory pathway (Tritsch et al., 2010). Spontaneous activity peaks during a time of significant change and maturation in auditory circuits, from the pruning of Type I SGN peripheral contacts with IHCs to the emergence of precise tonotopy and the remarkable growth of endbulbs in the CNC (Tritsch & Bergles, 2010; H. C. Wang & Bergles, 2015). Although cochlear ablation studies confirm that this spontaneous activity originates in the cochlea (Tritsch et al., 2010), it has been challenging to pinpoint the in vivo consequences for cochlear wiring and function.
Electrophysiological Maturation of SGNs
Long after their central and peripheral connections are established, SGNs continue to undergo molecular changes that produce adult electrophysiological signatures. These changes occur in concert with maturation of presynaptic inputs from the HCs and the olivocochlear efferent system, which further sculpt SGN firing behaviors. Intrinsically, developing SGNs must fine-tune the elements that generate and propagate action potentials, including expression and proper localization of a large cohort of voltage- and ligand-gated ion channels (Reijntjes & Pyott, 2016). Although the precise onset or dynamics of expression is unknown for most ion channels, adult profiles likely emerge after gene-specific variation in expression through development. For example, Kv1.1 protein level ramps up between P3 and P8 in mice and may contribute to a change in SGN excitability (Crozier & Davis, 2014). Proper localization of ion channels must go hand-in-hand with such expression changes. Indeed, in rats, the potassium channel Kv3.1b is initially diffuse but co-localizes with the scaffold protein AnkG in nascent nodes by P17 and stabilizes thereafter. Confinement of the sodium channel Nav1.6 to the AnkG domain, however, progresses until P30 (Kim & Rutherford, 2016). Hence, assembly of nodal components, which begins with the onset of myelination toward the end of the first postnatal week (Kim & Rutherford, 2016; Romand & Romand, 1985), continues at least until P30. Although nodes are almost fully assembled around hearing onset, the heminode, which is thought to be the axon initial segment, approaches mature configuration only after hearing begins.
Spatiotemporal changes in SGN ion channel profiles are contemporaneous with multifaceted changes in SGN physiology in the first few postnatal weeks. SGNs exhibit IHC-driven spontaneous activity at birth but undergo a switch from random to burst firing a few days later (Tritsch & Bergles, 2010). Burst firing disappears by the end of the second postnatal week and spontaneous SGN firing patterns instead exhibit temporal features suggestive of a Poisson process (A. B. Wong et al., 2013). It is unknown if such early changes in the temporal structure of SGN firing behavior are influenced by fluxes in SGN molecular profiles. However, cell-intrinsic changes likely occur, since spontaneous action potentials in immature SGNs occur both in response to input from IHCs and via IHC-independent mechanisms. With maturation, intrinsically generated spikes contribute less to SGN spontaneous activity, disappearing sometime in the third postnatal week (Wu, Young, & Glowatzki, 2016). Overall, the increase in the average rate and total range of spontaneous firing with the emergence of progressively higher SR is one key aspect of post-hearing electrophysiological maturation of SGNs (Kim & Rutherford, 2016; Wu et al., 2016). These changes in spontaneous activity are accompanied by maturation of responses to sound-evoked responses, including a decrease in threshold and latency, and an increase in the synchrony of auditory nerve fiber discharge (Geal-Dor, Freeman, Li, & Sohmer, 1993; Uziel, Romand, & Marot, 1981; A. B. Wong et al., 2013).
Whole-cell recordings of cultured SGNs have offered additional insights into intrinsic differences in SGN electrical properties. For example, Type I SGNs from neonatal mice exhibit tonotopic differences in membrane kinetics and degrees of accommodation (Crozier & Davis, 2014), which may be driven by differences in expression of voltage-gated potassium and calcium channels (Chen et al., 2011; Q. Liu, Lee, & Davis, 2014). Furthermore, a decrease in excitability occurs together with an alteration of accommodation properties within the first two postnatal weeks (Crozier & Davis, 2014), thus indicating that SGN maturation includes changes in cell-intrinsic electrical properties. However, the extent to which this accounts for changes observed in vivo is unknown. Likewise, although SGN firing behavior is subject to modulation by neurotrophins in vitro (Crozier & Davis, 2014; T. Wright, Gillespie, O’Leary, & Needham, 2016), the relevance for functional maturation of SGNs in vivo is unclear.
Several presynaptic changes that occur over the same period may further contribute to the emergence of mature SGN firing behaviors. These include transition of IHCs from producing action potentials to graded receptor potentials, maturation of presynaptic ribbon structures, and stronger coupling of Cav1.3 channels and vesicle release sites in IHCs (A. B. Wong et al., 2014), as well as establishment of synaptic contacts by LOC efferent terminals on Type I afferent endings. Indeed, heterogeneities in LOC innervation of SGNs (Yin et al., 2014) and functional diversity across IHC active zones (Meyer et al., 2009) are thought to contribute to the wide spontaneous firing rate diversity among SGNs in the mature cochlea.
How SGN physiological diversity is linked to observed molecular differences is not well understood. It is unclear, for example, which of the many genes that are differentially expressed among Type I SGNs underlie their in vivo physiological properties. For instance, in cultured neonatal SGNs, relative levels of two calcium-binding proteins, calbindin and calretinin, are variable and correlate with a temporal parameter of SGN firing (W. Liu & Davis, 2014). This observation suggests that SGN functional signatures may be shaped by calcium-binding proteins, in addition to the expected variation in ion channel components. More work is needed to establish a causal relation between such candidate molecules and distinct physiological features.
In summary, emerging evidence indicates that SGN molecular profiles undergo spatiotemporal refinement that may underlie the striking shifts in electrophysiological features observed across development. Although the biggest changes coincide with hearing onset around P11, subtler changes continue to shape SGN activity in subsequent weeks, driven both by shifts in the intrinsic properties of SGNs and the extrinsic influence of their presynaptic inputs, which mature over the same time frame.
SGNs are the main conduits through which auditory signals captured by the inner ear reach the brain. During development, SGNs make precise, specialized synaptic connections and acquire unique electrophysiological properties that involve both cell-intrinsic and activity-dependent mechanisms, starting well before animals are able to respond to airborne sounds. SGN processes are in place long before their target cells have matured and therefore play a significant role in auditory circuit assembly and maturation. Indeed, the arrival of SGN processes in the cochlear nucleus anlage marks the first time that cochlear tonotopy coordinates are relayed to the developing auditory brainstem, thus suggesting a crucial role for SGNs in setting up topographic order centrally. Indeed, the cochlea has a strong influence on the development of auditory circuits at multiple levels, from regulating cell survival locally to tonotopic order deeper in the brainstem. Given that SGNs connect the PNS and the CNS both developmentally and functionally, it will be important for the field as a whole to gain a better understanding of the primary effects of congenital mutations on SGNs and their impact on the organization of central auditory circuits.
We thank Lorna Wu for figure illustrations, and Michelle Frank, Dr. Austen Sitko, Dr. Kirupa Suthakar, Dr. Paul Fuchs, and Dr. David Ryugo for feedback. Our work is supported by funding from the NIH, the Lefler Family Foundation, and the Harvard Brain Initiative. BRS is a recipient of a F32 Postdoctoral Individual National Research Service Award and a Goldenson Postdoctoral Fellowship.
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