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Mechanical overstimulation causes acute injury and synapse loss followed by fast recovery in lateral-line neuromasts of larval zebrafish

  1. Melanie Holmgren
  2. Michael E Ravicz
  3. Kenneth E Hancock
  4. Olga Strelkova
  5. Dorina Kallogjeri
  6. Artur A Indzhykulian
  7. Mark E Warchol
  8. Lavinia Sheets  Is a corresponding author
  1. Department of Otolaryngology, Washington University School of Medicine, United States
  2. Eaton-Peabody Laboratory, Massachusetts Eye and Ear, United States
  3. Department of Otolaryngology–Head and Neck Surgery, Harvard Medical School, United States
  4. Department of Neuroscience, Washington University School of Medicine, United States
  5. Department of Developmental Biology, Washington University School of Medicine, United States
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Cite this article as: eLife 2021;10:e69264 doi: 10.7554/eLife.69264

Abstract

Excess noise damages sensory hair cells, resulting in loss of synaptic connections with auditory nerves and, in some cases, hair-cell death. The cellular mechanisms underlying mechanically induced hair-cell damage and subsequent repair are not completely understood. Hair cells in neuromasts of larval zebrafish are structurally and functionally comparable to mammalian hair cells but undergo robust regeneration following ototoxic damage. We therefore developed a model for mechanically induced hair-cell damage in this highly tractable system. Free swimming larvae exposed to strong water wave stimulus for 2 hr displayed mechanical injury to neuromasts, including afferent neurite retraction, damaged hair bundles, and reduced mechanotransduction. Synapse loss was observed in apparently intact exposed neuromasts, and this loss was exacerbated by inhibiting glutamate uptake. Mechanical damage also elicited an inflammatory response and macrophage recruitment. Remarkably, neuromast hair-cell morphology and mechanotransduction recovered within hours following exposure, suggesting severely damaged neuromasts undergo repair. Our results indicate functional changes and synapse loss in mechanically damaged lateral-line neuromasts that share key features of damage observed in noise-exposed mammalian ear. Yet, unlike the mammalian ear, mechanical damage to neuromasts is rapidly reversible.

Introduction

Hair cells are the sensory receptors of the inner ear and lateral-line organs that detect sound, orientation, and motion. They transduce these stimuli through deflection of stereocilia, which opens mechanically gated cation channels (LeMasurier and Gillespie, 2005; Qiu and Müller, 2018) and drives subsequent transmission of sensory information via excitatory glutamatergic synapses (Glowatzki and Fuchs, 2002). Excessive mechanical stimulation, such as loud noise, can damage hair cells and their synaptic connections to afferent nerves. The degree of damage depends on the intensity and duration of the stimulus, with higher levels of traumatic noise producing structural damage to sensory epithelia, including hair cells (Cho et al., 2013; Nordmann et al., 2000; Slepecky, 1986) and lower levels leading to various pathologies, including damage to the hair-cell mechanotransduction complex/machinery and stereocilia (Gao et al., 1992; Husbands et al., 1999), misshapen hair cells (Bullen et al., 2019), synaptic terminal damage, neurite retraction, and hair-cell synapse loss (Fernandez et al., 2020; Henry and Mulroy, 1995; Kujawa and Liberman, 2009; Puel et al., 1998). In addition to directly damaging hair cells, excess noise also initiates an inflammatory response (Hirose et al., 2005; Kaur et al., 2019). Such inflammation is mediated by macrophages, a class of leukocyte that responds to injury by clearing cellular debris and promoting tissue repair (Wynn and Vannella, 2016).

The variety of injury and range of severity suggests multiple mechanisms are involved in hair-cell organ damage associated with exposure to strong stimuli. Also unknown are the cellular processes mediating repair following such trauma. While hair cells show a partial capacity for repair of stereocilia and synaptic connections, some sub-lethal damage to hair cells is permanent. Numerous studies of the mammalian cochlea suggest that a subgroup of inner hair-cell synapses are permanently lost following noise exposure (Cho et al., 2013; Hickman et al., 2018; Kujawa and Liberman, 2009; Shi et al., 2013). Glutamate excitotoxicity is likely the pathological event that initiates noise-induced hair-cell synapse loss (Hu et al., 2020; Kim et al., 2019; Puel et al., 1998), but the downstream cellular mechanisms are still undefined. Further, the cellular mechanisms that promote hair-cell synapse recovery following damage-induced loss are also not understood.

Zebrafish have proven to be a valuable model system for studying the molecular basis of hair-cell injury and repair. Zebrafish sensory hair cells are structurally and functionally homologous to mammalian hair cells (Coffin et al., 2004; Kindt and Sheets, 2018; Sebe et al., 2017). In contrast to other vertebrate model organisms, zebrafish hair cells are optically accessible in whole larvae within the lateral-line organs. These sensory organs, called neuromasts, contain clusters of ~14 hair cells each and are distributed throughout the external surface of the fish to detect local water movements. Zebrafish can repair and regenerate damaged tissues, including lateral-line organs (Kniss et al., 2016; Xiao et al., 2015). This capacity for organ repair combined with optical accessibility allows us to study cellular and synaptic damage and repair in vivo following mechanical trauma.

In order to model physical damage to hair-cell organs in zebrafish lateral line, we developed a protocol to mechanically stimulate the lateral line of free-swimming 7-day-old larvae. Using this protocol, we were able to induce mechanical injury to lateral-line organs that resembled the trauma observed in the mammalian cochlea following acoustic overstimulation. We observed synapse loss in a subset of neuromasts that appeared morphologically intact, as well as hair-cell loss and afferent neurite retraction in a subset of neuromasts that appeared morphologically disrupted. Hair-cell mechanotransduction, as measured by uptake of the cationic dye FM1-43, was significantly reduced after mechanical injury. We also observed an inflammatory response similar to that observed in the mammalian cochlea after noise trauma. Remarkably, mechanically induced lateral-line damage appeared to rapidly recover; hair-cell number and morphology returned to normal within 4 hr following exposure, concurrent with clearance of cellular debris by macrophages. Additionally, neuromasts showed partial recovery of afferent innervation within 2 hr following exposure and completely recovered hair-cell morphology and FM1-43 uptake within 4 hr. Cumulatively, these results support that mechanically injured neuromasts show similar features of damage observed in noise exposed ears, yet rapidly repair.

Results

Mechanical overstimulation of zebrafish lateral-line hair cells

To mechanically damage hair cells of lateral-line organs in free-swimming 7-day-old zebrafish, we developed a stimulation protocol using an electrodynamic shaker to create a strong water wave stimulus (Figure 1A). The frequency used for mechanical stimulation was selected and further verified (see Method Details) based on previous studies showing 60 Hz to be within the optimal upper frequency range of mechanical sensitivity of superficial posterior lateral-line neuromasts, which respond maximally between 10and 60 Hz, but a suboptimal frequency for hair cells of the anterior macula of the inner ear (Levi et al., 2015; Trapani and Nicolson, 2010; Weeg et al., 2002). Dorsal-ventral displacement of a six-well dish at 60 Hz and acceleration of 40.3 m/s2 (±0.5 m/s2) created water flow and disturbance of the water surface that was strong enough to trigger ‘fast start’ escape responses—a behavior mediated in part by zebrafish lateral-line organs to escape predation (Figure 1B inset) (McHenry et al., 2009; Nair et al., 2015).

Intense water wave produced by shaker apparatus stimulates lateral-line hair cells and evokes a relevant behavior response.

(A) The apparatus: a magnesium head expander holding a 6-well dish mounted on a vertically oriented electrodynamic shaker housed inside a sound-attenuation chamber. The stimulus consisted of a 60 Hz vertical displacement of the plate (hatched arrows) driven by an amplifier and controlled by a modified version of the Eaton-Peabody Laboratory Cochlear Function Test Suite. (B,D) Swimming behavior of 7-day-old larvae during exposure to the wave stimulus. Traces in (D) represent tracking of corresponding circled fish over 500 ms (1000 fps/ 500 frames). Asterisks indicate a ‘fast escape’ response (B; inset). (C,E) Swimming behavior of larvae whose lateral-line neuromasts were ablated with low-dose CuSO4. Arrows in (E) indicate where a larva was swept into the waves and could no longer be tracked.

‘Fast start’ escape responses can also be activated by stimulating hair cells of the lateral line and/or the posterior macula in the ear (Bhandiwad et al., 2013). To verify that the observed escape responses were mediated predominantly by flow sensed by lateral-line hair cells rather than hair cells of the macula, we exposed a group of larvae to low dose (3 µM) copper sulfate (CuSO4) for 1 hr to specifically ablate lateral-line hair cells, but leave hair cells of the ear intact (Olivari et al., 2008). Following a 2 hour recovery after CuSO4 treatment, we recorded fish behavior with a high-speed camera during the stimulus (1000 fps for 10 s) and compared the responses of fish with lesioned lateral-line organs with those of untreated control siblings (Figure 1B and C; Video 1 ). When subjected to intense water flow, we found that ‘fast start’ responses—defined as a c-bend of the body occurring within 15 ms followed by a counter-bend (Burgess and Granato, 2007; McHenry et al., 2009)—occurred significantly less frequently in larvae with ablated lateral line organs than in siblings with intact lateral line (Figure 1C and E; avg. ‘fast start’ responses:: 0.4 ( ± 0.1)/s in lateral-line ablated vs 1.5 ( ± 0.1)/s in control; three trials,10 s per trial; **p = 0.0043). Accordingly, some CuSO4-treated fish were unable to escape the waves and were swept out of view (Figure 1E; arrowheads; Video 2). These observations indicate that the strong water wave generated by our device is stimulating lateral-line hair cells and evoking a behaviorally relevant response.

Video 1
Swimming behavior of control fish with intact lateral line organs.

Magenta circle indicates a fish prior to a ‘fast escape’ response.

Video 2
Swimming behavior of larvae whose lateral-line neuromasts were ablated with CuSO4.

Magenta circles indicate larvae that were swept into the waves and could no longer tracked.

A subset of mechanically injured neuromasts undergo physical displacement that is position dependent but does not require mechanotransduction

To induce mechanical damage to lateral line organs, larvae (10–15 per well) were exposed to an initial 20 minutes of strong water wave stimulus followed by 10 min of rest, then 2 hr of continuous stimulus (Figure 2—figure supplement 1 A). The 10-min break in exposure was introduced early on when establishing the stimulus duration because it appeared to enhance larval survival; it was therefore maintained throughout the study for consistency. Fish were euthanized and fixed immediately after exposure, then processed for immunohistofluorescent labeling of hair cells and neurons. Unexposed sibling fish served as controls. As posterior lateral-line (pLL) neuromasts have been shown to specifically initiate escape behavior in response to strong stimuli (Haehnel et al., 2012), analysis of the morphology of pLL neuromasts L3, L4, and L5 was conducted for exposed and control larvae (Figure 2A). Initially, we divided the observed neuromast morphology into two categories: ‘normal’, in which hair cells were radially organized with a relatively uniform shape and size, and ‘disrupted’, in which the hair cells were misshapen and displaced to one side, with the apical ends of the hair cells localized anteriorly (Figure 2B and C; see Methods Details for measurable criteria). Position of the neuromast along the tail was also associated with vulnerability to disruption; we observed a gradient of damage in the pLL from rostral to caudal that is L5 was more susceptible to disruption than L4, which was more susceptible to disruption than L3 (Figure 2F; Repeated measure One-way ANOVA *p = 0.0386, **p = 0.0049, *** = 0.0004).

Figure 2 with 2 supplements see all
Morphological changes in pLL neuromast hair cells exposed to strong water wave stimulus.

(A) Schematic of a larval zebrafish. Blue dots indicate neuromasts of the lateral-line organs; green lines indicate innervating afferent lateral-line nerves. pLL neuromasts L3, L4, and L5 were analyzed (dashed circles). (B–C) Maximum intensity dorsal top-down 2D projections of confocal images of control or stimulus-exposed neuromast hair cells (blue (B) or orange (C); Parvalbumin immunolabel). Exposed neuromast hair-cell morphology was categorized as ‘normal’ i.e. radial hair-cell organization indistinguishable from control or ‘disrupted’ i.e. asymmetric organization with the hair-cell apical ends oriented posteriorly. (D) Maximum intensity projections of supporting cells (SCs) expressing GFP (green), immunolabeled synaptic ribbons (magenta; Ribeye b) and all cell nuclei (blue; DAPI). Note that SCs underlying displaced hair cells also appear physically disrupted (indicated by white arrows). Scale bars: 5 µm (F) Average percentage of neuromasts with ‘disrupted’ morphology following mechanical stimulation. Each dot represents the percentage of disrupted neuromasts (NM) in a single experimental trial. Disrupted hair-cell morphology was place dependent, with neuromasts more frequently disrupted following sustained stimulus and when localized toward the posterior end of the tail (*p = 0.0386, **p = 0.0049, ***p = 0.0004) (G) Average percentage of exposed neuromasts (NM) with ‘disrupted’ morphology in lhfpl5b mutants, which lack mechanotransduction specifically in lateral-line hair cells, vs. heterozygous WT. lhfpl5b mutants show a similar gradient of neuromast disruption following mechanical injury as WT siblings. Error Bars = SEM.

Figure 2—source data 1

Summary of normal and disrupted neuromast counts following sustained and periodic stimulus exposures.

https://cdn.elifesciences.org/articles/69264/elife-69264-fig2-data1-v2.xlsx
Figure 2—source data 2

ummary of normal and disrupted neuromast counts in lhfpl5b mutants and wildtype siblings following sustained stimulus exposure.

https://cdn.elifesciences.org/articles/69264/elife-69264-fig2-data2-v2.xlsx

Additionally, we compared mechanical stress from our sustained exposure to an exposure protocol that delivered intermittent pulses of stimulus (‘periodic exposure’; Figure 2—figure supplement 1 A). We observed neuromast disruption less frequently with periodic exposures vs. sustained exposure of the same intensity (Figure 2—figure supplement 1 B; Unpaired t-test **p = 0.0034), supporting that displacement of neuromasts is a consequence of mechanical injury. Additionally, we examined hair-cell morphology in the ears of larvae exposed to sustained stimulus and observed no apparent damage (Figure 2—figure supplement 2), indicating our overstimulation protocol produces mechanical damage specifically to lateral-line organs.

To determine if hair-cell activity plays a key role in the displacement of neuromasts, we exposed lhfpl5b mutants—fish that have intact hair-cell function in the ear, but no mechanotransduction in hair cells of the lateral line—to sustained stimulation (Erickson et al., 2019). We observed comparable morphological disruption of mutant neuromasts lacking mechanotransduction (Figure 2G), suggesting that displacement of lateral-line hair cells is due to physical damage from the stimulus. Further, we observed the adjacent supporting cells in neuromasts with disrupted hair-cell morphology appeared similarly displaced and elongated (Figure 2E ‘disrupted’; white arrows), indicating that mechanical injury disrupts the structural integrity of the entire neuromast organ.

Hair-cell loss and reduced afferent innervation correspond to neuromast disruption

Moderate noise exposures can cause damage or loss of cochlear hair-cell synapses, including swelling and retraction of afferent nerve fibers, while more extended and/or severe exposures lead to hair-cell loss. To address whether mechanical damage in the lateral line produced similar morphological changes, we surveyed the number of hair cells per neuromast and the percentage of neuromast hair cells lacking afferent innervation in fish immediately following exposure to sustained stimulation (Figure 3). We observed a reduction in the number of hair cells per neuromast immediately following exposure (Figure 3D and E; **Adj p = 0.0019; N = 13 trials) as well as a significant reduction in the percentage of hair cells per neuromast innervated by afferent neurons (Figure 3F and G; ****Adj p < 0.0001; N = 12 trials).

Hair-cell loss and de-innervation is specific to ‘disrupted’ neuromasts.

(A–C) Representative maximum intensity projection images of control (A) or exposed lateral line neuromasts with ‘normal’ (B) or ‘disrupted’ (C) morphology immediately following sustained strong wave exposure (0 hr). Synaptic ribbons (magenta; Ribeye b) and hair cells (blue; Parvalbumin) were immunolabled. Afferent neurons were expressing GFP. Scale bar: 5 µm (D) Hair-cell number per neuromast immediately post exposure. A significant reduction in hair-cell number was observed (**Adj p = 0.0019) and was specific to ‘disrupted’ neuromasts (Adj p = 0.3859 normal, ****Adj p < 0.0001 disrupted). Pink box plot (Exp) represents pooled exposed neuromasts, while gray (Norm) and red (Dis) plots represent neuromasts parsed into normal and disrupted groups. Numbers beneath each plot indicate the number of neuromasts per group. Whiskers = min to max (E) Differences of least squares means in hair-cell number per neuromast between groups. Bars represent 95 % confidence interval (CI). (F) Percentage of neuromast hair cells innervated by afferent nerves. Numbers within each bar indicate the number of neuromasts per group. A significant portion of neuromast hair cells lacked afferent innervation following exposure (****Adj p < 0.0001). Hair cells lacking afferent innervation were specifically observed in disrupted neuromasts (Adj p = 0.7503 normal, ****Adj p < 0.0001 disrupted). (G) Differences of least squares means in % hair cells innervated per neuromast between groups. Bars represent 95% CI.

Figure 3—source data 1

Raw data and statistical analysis of hair-cell counts and innervation immediately following sustained stimulus exposure.

https://cdn.elifesciences.org/articles/69264/elife-69264-fig3-data1-v2.xlsx

As described in Figure 2F, we found on average~ half of L3-L5 neuromasts examined showed ‘disrupted’ hair-cell morphology immediately following sustained stimulus exposure. To define the associations between overall neuromast morphology and specific structural changes in mechanically injured neuromasts, we examined the numbers of hair cells and the percent of hair cells contacted by afferent fibers in exposed neuromasts parsed into ‘normal’ and ‘disrupted’ morphologies. With hair-cell number, we observed significant loss specifically in ‘disrupted’ neuromasts, while ‘normal’ neuromast hair-cell number appeared comparable to control (Figure 3D and E; Adj p = 0.3859 normal, ****Adj p < 0.0001 disrupted). With hair-cell afferent innervation we observed a similar trend that is, a significant number of hair cells lacked afferent innervation in stimulus exposed neuromasts with ‘disrupted’ morphology, but not ‘normal’ neuromasts (Figure 3F and G; Adj p = 0.7503 normal, ****Adj p < 0.0001 disrupted).

Mechanically overstimulated neuromasts with ‘normal’ morphology lose a greater number of hair-cell synapses when glutamate clearance is inhibited

Hair cells contain electron-dense presynaptic specializations—called dense bodies or synaptic ribbons—apposing afferent PSDs which constitute afferent synaptic contacts (Davies et al., 2001; Sheets et al., 2011). In the larval zebrafish lateral line, afferent nerve fibers innervate multiple hair cells per neuromast forming ~3–4 synaptic contacts per hair cell. To determine whether strong water wave stimulus exposure generated lateral-line hair-cell synapse loss, we counted the number of intact synapses (ribbons juxtaposed to PSDs; Figure 4A–C) in control and exposed larvae. We observed significant reduction in the number of intact synapses per hair cell following sustained exposure (Figure 4D and F; **Adj p = 0.0078). When we compared ‘normal’ and ‘disrupted’ neuromasts following exposure, we observed a loss of intact synapses per hair cell in all exposed neuromasts, with significantly fewer synapses in ‘normal’ exposed neuromasts relative to control (Figure 4E–F; **Adj p = 0.0043 normal, Adj p = 0.1207 disrupted). Additionally, significant synapse loss was observed in the neuromasts of fish exposed to the less mechanically damaging ‘periodic’ stimulus (Figure 2—figure supplement 1 F’).

Significant hair-cell synapse loss is observed in ‘normal’ neuromasts following mechanical overstimulation and exacerbated by blocking glutamate uptake.

(A–C) Representative maximum intensity projection images of unexposed (A), or stimulus exposed lateral-line neuromast with ‘normal’ (B) or ‘disrupted’ (C) morphology. Synaptic ribbons (magenta; Ribeye b), PSDs (green; MAGUK) and hair cells (blue; Parvalbumin) were immunolabled. Scale bar: 5 µm (D–E) Intact synapses per neuromast hair cell. Pink box plot in D (Exp) represents pooled exposed neuromasts while, in E, gray (Norm) and red (Dis) plots represent neuromasts parsed into normal and disrupted groups. Whiskers = min to max. The average number of intact synapses per hair cell was significantly reduced in exposed neuromasts (D; **Adj p = 0.0078); when parsed, this reduction was significant in the ‘normal’ exposure group relative to control (E; **Adj p = 0.0043 normal, Adj p = 0.1207 disrupted). (F) Differences of least squares means in number of intact synapses per hair cell between groups. Bars represent 95% CI. (G) The number of intact synapses per hair cell in larvae co-treated with TBOA, to block glutamate clearance, or drug carrier alone during exposure. Synapse loss was significantly greater in ‘normal’ neuromasts co-exposed to TBOA compared to fish co-exposed to the drug carrier alone (Two-way ANOVA. *p < 0.0187).

Figure 4—source data 1

Raw data and statistical analysis of synapse counts immediately following sustained stimulus exposure.

https://cdn.elifesciences.org/articles/69264/elife-69264-fig4-data1-v2.xlsx

Previous studies indicate that excess glutamate signaling may be a key factor driving inner hair-cell synapse loss following exposure to damaging noise (Chen et al., 2010; Kim et al., 2019). We therefore inhibited glutamate clearance from neuromast hair-cell synapses by pharmacologically blocking uptake with the glutamate transporter antagonist Threo-beta-benzyloxyaspartate (TBOA) during sustained stimulus exposure. We observed a significantly greater degree of hair-cell synapse loss in stimulated neuromasts with ‘normal’ morphology co-treated with TBOA than in stimulated neuromasts co-treated with drug carrier alone (Figure 4E; Two-way ANOVA. *p < 0.0187), suggesting glutamate excitotoxicity contributes to hair-cell synapse loss observed in mechanically overstimulated neuromasts with intact morphology.

To further characterize synapse loss in relation to afferent neurite retraction, we examined fish in which we immunolabeled synaptic ribbons, PSDs, afferent nerve fibers, and hair cells (Figure 5A and B). We then quantified instances where synapses, that is juxtaposed pre- and postsynaptic components associated with hair cells, were no longer adjacent to an afferent nerve terminal. As these synapses appeared detached and suspended from their associated neurons, making them no longer functional, we refer to them as synaptic debris. While we rarely observed synaptic debris in unexposed neuromasts, we observed a greater relative frequency of synaptic debris in neuromasts exposed to strong water wave stimulus (Figure 5C; One sample Wilcoxon test, p = 0.1250 (control), *p = 0.0313(normal), **p = 0.0039(disrupted)). Taken together, we observed two distinct types of morphological damage to afferent synapses in mechanically overstimulated neuromasts: loss of synapses within neuromasts that appear intact that is exacerbated when glutamate uptake is blocked and a higher incidence of synaptic debris that appear detached from afferent neurites in all exposed neuromasts.

Mechanically overstimulated neuromasts showed retracted neurites and detached synaptic debris.

(A–B) Representative images of control (A) and exposed (B) neuromasts. Synaptic ribbons (magenta; Ribeye b) and PSDs (green; MAGUK) were immunolabeled; hair cells were also immunolabeled, but not shown for clarity. Afferent neurons (white) were labeled with GFP. Insets: Arrows indicate intact synapses adjacent to afferent neurons; arrowheads (B) indicate synaptic debris. Scale bars: 5 µm (main panels), 1 µm (insets). (C) Frequency histogram of observed synaptic debris per neuromast (NM). While control neuromasts occasionally had one detached synapse, exposed neuromasts were observed that had up to five detached synapses.

Mechanically overstimulated lateral line neuromasts show signs of hair-cell injury and macrophage recruitment

The inner ears of birds and mammals possess resident populations of macrophages, and additional macrophages are recruited after acoustic trauma or ototoxic injury (Warchol, 2019). A similar macrophage response occurs at lateral line neuromasts of larval zebrafish after neomycin ototoxicity (Warchol et al., 2020). Analysis of fixed specimens, as well as time-lapse imaging of living fish (e.g. Hirose et al., 2017), has demonstrated that macrophages migrate into neomycin-injured neuromasts and actively phagocytose the debris of dying hair cells. To determine whether a similar inflammatory response also occurs after mechanical injury to the lateral line, we characterized macrophage behavior after sustained stimulation. These studies employed Tg(mpeg1:yfp) transgenic fish, which express YFP in all macrophages and microglia. Fish were fixed immediately after exposure, or allowed to recover for 2, 4, or 8 hr. Control fish consisted of siblings that received identical treatment but were not exposed to mechanical stimulation. Data were obtained from the two terminal neuromasts from the pLL of each fish (Figure 6A). In agreement with data shown in Figure 3D, we observed a modest but significant decline in hair-cell number in specimens that were examined immediately and at 2 hr after sustained exposure (Figure 6B; **p < 0.003). Consistent with earlier studies (Hirose et al., 2017), 1–2 macrophages were typically present within a 25 µm radius of each neuromast (Figure 6C). In uninjured (control) fish, those macrophages remained outside the sensory region of the neuromast and rarely contacted hair cells. However, at 2, 4, and 8 hr after sustained stimulus, we observed increased macrophage-hair cell contacts (Figure 6D; *p = 0.024), as well as the presence of immunolabeled hair-cell debris within macrophage cytoplasm (suggestive of phagocytosis, Figures 6A and 4 h., arrow). Macrophage-hair cell contact and phagocytosis peaked at 2 hr after exposure (Figure 6E; **p = 0.0013 (2 h)). Notably, the numbers of macrophages within a 25 µm radius of each neuromast remained unchanged at all time points after exposure, suggesting that the inflammatory response was mediated by local macrophages and that mechanical injury did not recruit macrophages from distant locations (Figure 6C). This pattern of injury-evoked macrophage behavior is qualitatively similar to the macrophage response observed in the mouse cochlea after acoustic trauma (Hirose et al., 2005; Kaur et al., 2019; Kaur et al., 2015).

Macrophage response to mechanical overstimulation of lateral line hair cells.

Experiments used Tg(mpeg1:yfp) fish that express YFP under regulation of the macrophage-specific mpeg1 promoter. All images and data were collected from the two distal-most neuromasts of the posterior lateral line (Figure 2A; term). (A) Macrophages (green) responded to mechanical injury by entering neuromasts, contacting hair cells and internalizing Otoferlin-immunolabeled debris (arrows, magenta). Images show examples of macrophage behavior at different time points after noise trauma. (B) Quantification of hair-cell number in the terminal neuromasts. Hair-cell number was significantly reduced at 0–2 hr after noise exposure (Mixed-effects analysis: **p < 0.003). (C) Quantification of macrophages within a 25 µm radius of the neuromasts at 0–8 hr after noise injury. Most neuromasts possessed 1–2 nearby macrophages and this number was not changed by noise exposure. (D) Quantification of direct contacts between macrophages and hair cells. The number of macrophage-hair cell contacts was counted at each survival time after noise exposure and normalized to the total number of sampled neuromasts. Increased levels of contact were observed at 2 and 4 hr after noise (*p = 0.0243). (E) Quantification of phagocytosis as a function of post-noise survival time. The numbers of macrophages that had internalized otoferlin-labeled material were counted at each time point and normalized to the total number of sampled neuromasts. The percentage of macrophages that contained such debris was significantly increased at 0–2 hr after strong water wave stimulus (*p = 0.0465; **p = 0.0013). Data were obtained from 26 to 50 neuromasts/time point. Error Bars = SD.

Mechanically injured neuromasts rapidly repair following exposure

To determine if damage to mechanically injured neuromasts was progressive, persistent, or reversible, we compared neuromast morphology, hair-cell number, and innervation at 0, 2, and 48 hr following sustained exposure to strong water wave stimulus. We observed a decrease in the percentage of neuromasts showing ‘disrupted’ morphology 2 hr following exposure, relative to fish fixed immediately following exposure (Figure 7A; 54 % disrupted (0 h) vs. 32 % disrupted (2 hr); N = 4 trials; ), suggesting that physical disruption of neuromast morphology following mechanical injury is rapidly reversible. Consistent with this observation, the average hair-cell number per neuromast at 2 hr post-exposure appeared to recover (Figure 7C; *Adj p = 0.0321 (0h disrupted), Adj p = 0.1875 (2h disrupted); N = 6 trials; ). Recovery of hair-cell number occurred within 2–4 hours (Figure 6A and B) and corresponded with macrophages infiltrating neuromasts and phagocytosing hair-cell debris (Figure 6A and E). We also observed, compared to immediately following exposure, a lesser degree of afferent fiber retraction (Figure 7E; ****Adj p < 0.0001 (0h disrupted), **Adj p = 0.0016 (2h disrupted); N = 6 trials) indicating partial recovery of innervation.

Mechanically overstimulated neuromasts recover hair-cell morphology, hair-cell number, and innervation.

(A,B) Average percentage of exposed neuromasts with ‘normal’ vs. ‘disrupted’ morphology following exposure. Each dot represents the percentage of disrupted neuromasts (L3–L5) in a single experimental trial; lines connect data points from the same cohort of exposed fish following 2 hr (A) or 48 hr (B) recovery. (C,D) Multilevel analysis of hair-cell number per neuromast immediately (0 hr) post-exposure or after 2 or 48 hr recovery. Numbers beneath each plot indicate the number of neuromasts per group. Whiskers = min to max. Morphologically ‘disrupted’ neuromasts have significantly fewer hair cells at 0 hr but not 2 hr following exposure C; *Adj p = 0.0321 (0h disrupted), Adj p = 0.1875 (2h disrupted). Most exposed neuromasts were morphologically ‘normal’ following 48 hr recovery and had a comparable number of hair cells relative to control (D; Adj p = 0.4443). (E,F) The percentage of ‘disrupted’ neuromast hair cells lacking afferent innervation was significant following 0 hr and 2 hr recovery (E; ****Adj p < 0.0001 (0h disrupted), **Adj p = 0.0016 (2h disrupted)). All hair cells were fully innervated following 48 hr recovery, including the few neuromasts with ‘disrupted’ morphology (F; aligned red dots).

Figure 7—source data 1

Summary of normal and disrupted neuromast counts following sustained exposure with 0, 2, or 48 hr recovery.

https://cdn.elifesciences.org/articles/69264/elife-69264-fig7-data1-v2.xlsx
Figure 7—source data 2

Raw data and statistical analysis of hair-cell counts and innervation following sustained stimulus exposure with 0 and 2 hr recovery.

https://cdn.elifesciences.org/articles/69264/elife-69264-fig7-data2-v2.xlsx
Figure 7—source data 3

Raw data and statistical analysis of hair-cell counts and innervation following sustained stimulus exposure with 0 and 48 hr recovery.

https://cdn.elifesciences.org/articles/69264/elife-69264-fig7-data3-v2.xlsx

A recent study characterized zebrafish lateral-line hair-cell damage induced by exposure to ultrasonic waves and reported a delayed hair-cell death and synapse loss 48–72 hr following exposure (Uribe et al., 2018). To determine if lateral line neuromasts exposed to the strong wave stimulus generated by our apparatus underwent delayed hair-cell loss, we examined hair-cell morphology, number, and innervation 48 hr following sustained stimulus exposure. Most exposed neuromasts examined showed ‘normal’ HC morphology Figure 7B; with no significant difference in hair-cell number (Figure 7D; Adj p = 0.4443; N = 4 trials). Hair-cell afferent innervation after 48 hr was comparable to control fish; even the few neuromasts that remained morphologically ‘disrupted’ were fully innervated (Figure 7F).

PSDs are enlarged in all neuromasts following mechanical overstimulation

Previous studies in mice and guinea pigs indicate moderate noise exposures modulate the size of synaptic components (Kim et al., 2019; Song et al., 2016). To determine if pre- and postsynaptic components were also affected in our model, we compared the relative volumes of neuromast hair-cell presynaptic ribbons and their corresponding PSDs in control and stimulus exposed larvae. We observed a moderate reduction in synaptic-ribbon size following exposure; ribbon volumes were significantly reduced relative to controls following 2 hr recovery (Figure 8B; Kruskal-Wallis test *p = 0.0195; N = 3 trials), and this reduction was specific to ‘disrupted’ neuromasts (Figure 8C). While the changes in ribbon volume we observed were modest and delayed in onset, we saw dramatic enlargement of PSDs immediately and 2 hr following exposure (Figure 8D; Kruskal-Wallis test ****p < 0.0001; N = 3 trials). In contrast to the observed reduction in ribbon size, relative PSD volumes were significantly enlarged in all exposed neuromasts regardless of whether neuromast morphology was ‘normal’ or ‘disrupted’ (Figure 8E). These data reveal enlarged PSDs as the predominant structural change in mechanically overstimulated neuromast hair-cell synapses.

Changes in synaptic ribbon and PSD sizes following sustained mechanical overstimulation.

(A-A’’) Representative images of control (A) and exposed (A’, A”) neuromasts. Synaptic ribbons (magenta; Ribeye b), PSDs (green; MAGUK), and hair cells (blue, Parvalbumin) were immunolabeled. Scale bars: 5 µm (main panels), 1 µm (insets). (B–E) Box and whisker plots of relative synapse volumes normalized to 0 hr control. Whiskers indicate the min. and max. values; ‘+’ indicates the mean value, horizontal lines indicate the relative median value of the control. (B) Ribbon volume appeared comparable to control immediately following exposure but was reduced 2 hr after exposure (*p = 0.0195). (C) Significant reduction in ribbon size relative to control was specific to disrupted neuromasts (Kruskal-Wallis test: ***p = 0.0004 (2h)). (D) Significantly larger PSDs were observed both immediately and 2 hr following exposure (****p < 0.0001). (E) Enlarged PSDs were present in both ‘normal’ and ‘disrupted’ exposed neuromasts, with a greater enlargement observed 0 hr post-exposure (Kruskal-Wallis test: ****p < 0.0001 (0h); ***p = 0.0001, **p = 0.0024 (2h)).

Mechanically injured neuromasts have damaged kinocilia, disrupted hair-bundle morphology, and reduced FM1-43 uptake immediately following exposure

An additional consequence of excess noise exposure in the cochlea is damage to mechanosensitive hair bundles at the apical end of hair cells and, correspondingly, disruption of mechanotransduction (Wagner and Shin, 2019). Larval zebrafish lateral-line hair cells each have a hair bundle consisting of a single kinocilium flanked by multiple rows of actin-rich stereocilia (Kindt et al., 2012). To determine if our exposure protocol damaged apical hair-cell structures, we used confocal imaging and scanning electron microscopy (SEM) to assess hair bundle morphology in both unexposed control larvae and larvae fixed immediately following sustained exposure. All neuromasts throughout the fish were evaluated, but to remain consistent with our fluorescence imaging results, we closely assessed the appearance of the caudal pLL neuromasts. We found the caudal neuromasts to be more damaged than the ones positioned more rostrally: the frequency of neuromasts with apparently disrupted appearance increased the closer its position to the tail (Figure 9—figure supplement 1). This is consistent with our fluorescence observations (Figure 2F; Figure 9—figure supplement 2 B) in which L5 neuromasts were more likely to be disrupted than more anteriorly positioned L3.

A closer examination of neuromast morphology revealed a difference of the kinocilia length and bundling. The neuromasts of the control fish carry a bundle of long (10–15 µm), uniformly shaped kinocilia (Figure 9A–E). In contrast, the neuromasts of the fish fixed immediately after sustained exposure often appear to carry much shorter kinocilia (Figure 9F–H, yellow arrows), which lack bundling and, in some cases, pointing to different directions (Figure 9G and H). The apparent kinocilia length difference between control and overstimulated neuromasts suggests at least some kinocilia may undergo a catastrophic damage event at the time of stimulation, as their distal parts break off the hair cells. This is further supported by some examples of kinocilia with thicker, ‘swollen’ proximal shafts closer to the cuticular plate of the cell, some of which extend into a thinner distal part while others appear to lack the distal part completely (Figure 9I–K, yellow arrowheads). Accordingly, the average diameter of kinocilia at the level of the hair bundle (L2-5) was significantly larger than control, with a few kinocilia showing dramatically thicker widths ~ 2 x greater than the thickest control (Figure 9L; ***p = 0.0007). When measured ~3–5 µm above the bundle, the exposed neuromast kinocilia have a somewhat larger average diameter relative to control, but not as dramatic as observed at the base (Figure 9M; *p = 0.0243). Stereocilia bundles from both groups of animals carried tip links, but we were unable to systematically evaluate and quantify their abundance. However, we observed signs of damaged bundle morphology following overstimulation, as they often appeared splayed, with gaps between the rows of stereocilia (Figure 9I–K).

Figure 9 with 2 supplements see all
Scanning electron microscopy imaging of neuromasts following mechanical injury reveals disorganized hair-cell stereocilia bundles and damaged kinocilia.

(A–E) Representative images of tail neuromasts of control fish larvae. Each hair cell carries a kinocilium, which is visibly thicker than its neighboring actin-filled, mechanosensitive stereocilia: see panel C featuring both structures at higher magnification (the kinocilium diameter is 220 nm, while stereocilia measured 90–110 nm). The kinocilia of control neuromasts are long (10–15 µm) and bundled together, while the stereocilia bundles have an apparent staircase arrangement. (F–K) Representative images of damaged tail neuromasts immediately following noise exposure featuring short (F-H, yellow arrows), disorganized (G, H), and swollen (I-K, yellow arrowheads) kinocilia, and disorganized stereocilia. (K) Same stereocilia bundle as in J marked with an asterisk at higher magnification to highlight the difference in the diameter of the kinocilium (360 nm) and neighboring stereocilia (85–100 nm) for noise exposed hair cells, as compared to the control hair cells in C. Scale bars: A, B, D-J – 2 µm; C, K – 500 nm. (L–M) Kinocilia diameter at bundle level (L; Mann Whitney test ***p = 0.0007) and 3–5 µm above bundle level (M; Welch’s t test *p = 0.0243). Exposed NM data in L were not normally distributed (D'Agostino-Pearson test ****p < 0.0001). Error Bars = SD.

To evaluate the effect of mechanical overstimulation on hair bundle function in relation to hair bundle morphology, we assessed mechanotransduction by briefly exposing free-swimming larvae to the fixable mechanotransduction-channel permeable dye FM1-43X (Holmgren and Sheets, 2021; Toro et al., 2015). We treated control and mechanically overstimulated larvae with FM1-43X immediately and at several time points up to 4 hr following exposure, then fixed the larvae and co-labeled hair-cell stereocilia with fluorescently conjugated phalloidin (Figure 10A–C). We observed a significant reduction in the relative intensity of FM1-43 in all exposed neuromasts immediately following exposure (Figure 10 D and F; ****p < 0.0001). While phalloidin labeling of stereocilia revealed what appeared to be tapered hair bundles in some exposed neuromasts (Figure 10B’; yellow arrows), average stereocilia length obtained from 3D interpolated confocal image stacks was not significantly altered (Figure 10E). Remarkably, FM1-43FX uptake showed recovery within 30 min and fully recovered over several hours. (Figure 10C and D). The degree of FM1-43FX fluorescence recovery following mechanical damage appeared to correspond with recovery of neuromast morphology; following 4 hr, nearly all neuromasts exposed to strong wave stimulus showed ‘normal’ morphology and relative FM1-43X fluorescence that was comparable to control (Figures 10D, F, 4h recovery). This observed timeline of morphological recovery coincides with macrophage recruitment and phagocytosis hair-cell debris peaking 2 hr post exposure (Figure 6D and E) followed by full recovery of hair-cell number between 2 and 4 hr post exposure (Figure 6B). Additionally, we quantified proliferating neuromast cells by treating control and mechanically overstimulated to 5-ethynyl-2-deoxyuridine (EdU) larvae for 4 hr following exposure and saw no difference in the number of EdU positive cells per neuromast in each condition (Figure 11A–C). Cumulatively, these data support that most damaged hair cells are repaired within mechanically injured neuromasts.

Hair-cell mechanotransduction was significantly reduced but rapidly recovered following mechanical overstimulation.

(A–C) Representative images of hair-cell stereocilia (conjugated phalloidin, gray) and FM1-43FX fluorescence intensity of the corresponding neuromast in representative control (A) or mechanically overstimulated fish immediately (B, B’) or 4 hr (C) following exposure. Yellow arrows in (B’) indicate phalloidin labeling that appeared tapered. (D) Average relative FM1-43FX fluorescence intensity measurements in control and exposed neuromasts over 4 hr of recovery. FM1-43FX uptake was significantly reduced in exposed neuromasts immediately following mechanical overstimulation but appear to completely recover by 4 hr (Tukey’s multiple comparisons test ****p < 0.0001 (0h), p = 0.0579 (1 h), p = 0.8387 (2h), p = 0.8387 (4h)). Dashed lines indicate FM1-43FX fluorescence intensity measurements in exposed neuromasts parsed into ‘normal’ and ‘disrupted’ morphologies. (E) Average stereocilia length of centrally localized hair bundles in control and exposed neuromasts. Dashed lines indicate measurements in exposed neuromasts parsed into ‘normal’ and ‘disrupted’ morphologies. Error Bars = SD (F) Relative FM1-43FX fluorescence in both ‘normal’ and ‘disrupted’ exposed neuromasts was significantly reduced immediately following exposure but recovered over time (Tukey’s multiple comparisons test ****p < 0.0001, *p = 0.0328 (0h); ****p < 0.0001, **p = 0.0098 (1h); **p = 0.0025 control vs. dis, **p = 0.0089 normal vs. dis (2 h)). Each point represents an individual neuromast. Nearly all observed exposed neuromasts appeared morphologically normal following 4 hr; note only one neuromast data point in the disrupted category of the 4 hr recovery graph. Data were obtained from 26 to 32 neuromasts per condition over three trials.

Neuromasts show no change in cell proliferation following mechanical overstimulation.

(A,B) Representative cross-section images of EdU (magenta) labeling of proliferating neuromast cells. Fish were exposed to EdU for 4 hr following stimulus exposure. Supporting cells (SC) were expressing GFP. Scale bars: 5 µm (C) Average number of EdU + cells per neuromast were comparable in control and exposed larvae. Data were obtained from 33 to 34 neuromasts per condition over three trials (Two-way ANOVA. p = 0.4193). Bars represent 95% CI.

Discussion

To model mechanical injury resulting from noise trauma in the zebrafish lateral line, we describe here a method to mechanically overstimulate neuromasts of the posterior lateral line. Using this method, we observed: (i) hair-cell synapse loss in a subset of stimulus exposed neuromasts with intact morphology, (ii) morphological displacement, hair-cell loss, and afferent deinnervation in a subset of mechanically disrupted neuromasts, (iii) an inflammatory response that peaked 2–4 hr following stimulus exposure, (iv) kinocilia and hair bundle damage, and (v) reduced FM1-43 uptake in all immediately following exposure. Remarkably, mechanically injured neuromasts rapidly recover following exposure; neuromast morphology, innervation, and mechanotransduction showed significant recovery within an hour, and most neuromasts were completely recovered within 4 hr post exposure.

Zebrafish lateral-line as a model for sub-lethal mechanical damage and noise-induced synapse loss

Mechanical damage to the zebrafish lateral line induced by strong water wave stimulus is observable immediately following exposure, is specific to the lateral-line organ, and appears to be rapidly repaired. These observations contrast with a recently published noise damage protocol for larval zebrafish which used ultrasonic transducers (40 kHz) to generate small, localized shock waves (Uribe et al., 2018). They reported delayed hair-cell death and modest synapse loss that was not apparent until 48 hr following exposure, was not accompanied by decreased mechanotransduction, and was observed in the inner ear as well as the lateral-line organs. Some of the features of the damage they observed—delayed onset apoptosis and hair-cell death—may correspond to lethal damage following blast injuries. We propose features of the damage we observe with our stimulus protocol—reduced mechanotransduction, hair-cell synapse loss, and rapid inflammatory response—may correspond to sub-lethal noise-induced damage of hair-cell organs. Because of differences in the nature of the stimuli in these two studies, it is difficult to directly compare the pathological outcomes. Mechanical overstimulation in the Uribe et al., study was induced using ultrasonic (40 kHz) actuators. Such high frequencies are far outside those that are detected by lateral line neuromasts (Levi et al., 2015; Trapani and Nicolson, 2010) supporting that cavitation in the water medium is likely causing the observed damage. In contrast, our study delivered a stimulus of high intensity 60 Hz water waves directly to the fish. This frequency is within the range of sensitivity of lateral line neuromasts of larval zebrafish and evoked a lateral-line mediated behavior (Figure 1B and C) suggesting that the hair cells were being directly stimulated by the water motion. The present method more closely resembles the techniques that are typically used to study noise damage in the mammalian cochlea, where high-intensity acoustic energy causes hair cell and synaptic injury in specific regions of the cochlea that are best-responsive to the frequency of the stimulus. This idea is further supported by the observation that synapse loss in hair cells exposed to strong wave stimulation is greater when glutamate uptake is blocked (Figure 4E), suggesting a shared mechanism of glutamate excitotoxicity between noise-exposed mammalian ears and strong water wave stimulus exposed lateral-line organs (Kim et al., 2019; Sebe et al., 2017).

Disruption of neuromast morphology is a consequence of mechanical injury

Strong water wave exposure produced a percentage of pLL neuromasts that were morphologically ‘disrupted’. Several observations support that such ‘disrupted’ neuromasts represent mechanical injury to neuromast organs. One is that lhfpl5b mutant neuromasts, which lack mechanotransduction specifically in lateral-line hair cells, were comparably vulnerable to physical disruption as their wild-type siblings (Figure 2G; Erickson et al., 2019). This finding supports that hair-cell activity during stimulation does not underly the physical displacement of hair cells observed following strong water wave stimulus. Additionally, physical disruption of the neuromast affects the whole organ—hair cells and their adjacent supporting cells (Figure 2D). This observation contrasts with what is observed in mammalian ears exposed to high intensity noise, where mechanical injury to outer hair cells is localized to stereocilia disruption and gross displacement of hair cells is not found (Wang et al., 2011). Speculatively, displacement of hair cells in mechanically injured neuromasts may be due to loss of structural support from displaced supporting cells. As the lateral-line organs are superficially localized on the surface of the skin, some of the intense mechanical tension applied across the tail is likely coupled to neuromasts leading to the physical displacement of a subset of exposed neuromasts. Finally, we observed reduced FM1-43 uptake in ‘disrupted’ neuromasts (Figure 10F) and measurable changes to kinocilia likely reflecting mechanical damage (Figure 9; Wagner and Shin, 2019). One notable limitation to using this model is that the evaluation of subtle and functionally relevant damage to hair-cell stereocilia, such as loss of tip links and damage to the actin core, is a significant challenge in zebrafish neuromasts due to their small size. Nevertheless, it is remarkable how rapidly mechanically disrupted lateral-line neuromasts regain normal morphology and FM1-43 uptake (Figure 10). The superficial lateral line’s direct exposure to the environment may require more robust mechanisms for repair, unlike hair-cell organs of mammals which are encased in bone.

Hair-cell overstimulation and synapse loss

In contrast to de-innervation and modest hair-cell loss that we observed in mechanically disrupted neuromasts, we saw significant loss of hair-cell synapses in neuromasts that were exposed to strong water wave stimulus but not mechanically disrupted i.e. ‘normal’ (Figure 4). Two notable observations were made in exposed neuromasts regarding synapse loss. First, loss of synapses in ‘normal’ exposed neuromasts was markedly more severe when synaptic glutamate clearance was inhibited (Figure 4G), suggesting that synapse loss may reflect moderate hair-cell damage resulting from overstimulation of hair cells and excess glutamate accumulation. Involvement of glutamate signaling as a key mediator of noise-induced synapse loss has recently been reported in mice; loss of glutamate signaling prevents noise-induced synapse loss and pharmacologically blocking postsynaptic Ca2+ permeable AMPA receptors protects against cochlear hair-cell synapse loss from moderate noise exposure (Hu et al., 2020; Kim et al., 2019). Second, synapse loss occurred in neuromasts that appeared to be fully innervated (Figures 3B and 4 B). This observation was initially surprising given that pharmacologically activating evolutionarily conserved Ca2+ permeable AMPARs has been shown to drive afferent terminal retraction in the zebrafish lateral line (Sebe et al., 2017). We propose that subtle damage to afferents in ‘normal’ exposed neuromasts may accompany synapse loss but not be apparent as loss of innervation, as single afferent processes innervate multiple hair cells of the same polarity within an individual neuromast (Dow et al., 2018; Faucherre et al., 2009). This idea is further supported by our observation that the relative frequency of synaptic debris (i.e. synaptic components that appear detached from afferent neurites) was higher in exposed neuromasts with ‘normal’ morphology relative to control (Figure 5C). We speculate that synaptic debris observed in exposed neuromasts with ‘normal’ morphology may be the result of excitotoxic damage at synaptic terminals (Sebe et al., 2017) while synaptic debris observed in ‘disrupted’ neuromasts may reflect mechanical disruption of supporting cells and retraction of afferent innervation from a subset of hair cells (Figure 3C and F).

In addition, glutamate signaling may not be the only driver of synapse loss resulting from excess stimulation. Studies in zebrafish and mice support that mitochondrial stress combined with excessive synaptic activity may also contribute to hair-cell synapse loss (Wang et al., 2018; Wong et al., 2019). Future work using this model to examine the effect of excess mechanical stimulation on mutants with reduced glutamate release or impaired glutamate clearance (to elevate or reduce glutamate in the synaptic cleft, respectively) combined with modified mitochondrial function may define the relative roles of glutamate excitotoxicity and hair-cell mitochondrial stress to synaptic loss.

Role of inflammation following mechanical injury to lateral-line organs

Our results indicate that mechanical injury to neuromasts evokes an inflammatory response. Prior studies of larval zebrafish have shown that macrophages reside near the borders of uninjured neuromasts and migrate into neuromasts after ototoxic injury (Hirose et al., 2017). We found that macrophages migrate into neuromasts within ~2 hr of mechanical injury, where they contact hair cells and, in some cases, engulf hair-cell debris. Although this macrophage response is similar to that which occurs after ototoxic injury to neuromasts (Carrillo et al., 2016; Hirose et al., 2017; Warchol et al., 2020), the extent of hair-cell loss after mechanical overstimulation is much less than the injury that occurs after ototoxicity. We observed macrophage entry in 30–40% of exposed neuromasts, despite modest hair-cell loss (Figure 6B and D). It is possible that the morphological changes characteristic of mechanically injured neuromasts are accompanied by the release of macrophage chemoattractants. In addition, studies of noise exposure to the mammalian cochlea indicate that high levels of synaptic activity (without accompanying hair-cell loss) can evoke macrophage migration to the synaptic region (Kaur et al., 2019). In either case, the signals responsible for such recruitment remain to be identified. The observation that macrophages had internalized immunolabeled hair-cell material further suggests that recruited macrophages engage in the phagocytosis of hair-cell debris, but it is not clear whether macrophages remove entire hair cells or target specific regions of cellular injury (e.g. synaptic debris; Figure 5). In any case, our data indicate that the macrophage response to mechanical injury of zebrafish lateral-line neurons is similar to that which occurs after noise injury to the mammalian cochlea (Warchol, 2019) and suggests that zebrafish may be an advantageous model system in which to identify the signals that recruit macrophages to sites of excitotoxic injury.

Hair-cell synapse morphology following mechanical overstimulation

Immediately following mechanical overstimulation, the most pronounced morphological change we observed in hair-cell synapses was significantly enlarged PSDs (Figure 8D and E). Speculatively, PSD enlargement may be a consequence of reduced glutamate release from hair cells following sustained intense stimulation. Mice and zebrafish fish lacking hair-cell glutamatergic transmission have enlarged postsynaptic structures (Kim et al., 2019; Sheets et al., 2012), indicating that glutamate may regulate postsynaptic size. While our data do not directly support this idea, we speculate that reduced glutamatergic transmission in mechanically overstimulated neuromasts may be a consequence of transiently impaired mechanotransduction (Figure 10B and D; Zhang et al., 2018). Alternatively, cholinergic efferent feedback, which has been shown to hyperpolarize lateral-line hair cells, may reduce hair-cell excitability during sustained strong wave exposure to protect against excess glutamate release, excitotoxic damage, and synapse loss (Carpaneto Freixas et al., 2021). Interestingly, presynaptic ribbons were not similarly enlarged, but instead showed a modest reduction in size following mechanical injury. Functional imaging of zebrafish lateral line has shown that a subset of hair cells in each neuromast are synaptically silent, and these silent hair cells can become active following damage (Zhang et al., 2018). As ribbon size has been observed to correspond with synaptic activity (Merchan-Perez and Liberman, 1996; Sheets et al., 2012), reduction in ribbon size may reflect recruitment of synaptically silent hair cells following mechanically induced damage. Future functional studies are needed to determine if mechanical overstimulation recruits more active hair-cell synapses, and to verify whether glutamate release from active synapses is reduced following mechanical overstimulation.

Lateral-line neuromasts fully recover following mechanical damage

Previous studies indicate mammalian cochlear hair cells have some capacity for repair following sub-lethal mechanical damage, including tip-link repair and regeneration of a subset of ribbons synapses (Indzhykulian et al., 2013; Jia et al., 2009; Kim et al., 2019). But such ability is limited, and our understanding of hair-cell repair mechanisms is incomplete. By contrast, we observe complete recovery of neuromast morphology and innvervation following mechanical trauma to the zebrafish lateral line. The cellular mechanisms responsible for such repair are not fully defined, but may involve regulation of neurite growth, glutamate signaling, inflammation, and/or neurotrophic factors (Kaur et al., 2019; Kim et al., 2019; Wan et al., 2014). Further study using this zebrafish model of mechanical overstimulation may provide insights that will assist in the development of methods for promoting complete hair-cell repair following damaging stimuli to the mammalian cochlea.

We also found that mechanical trauma resulted in a small degree of hair-cell loss in disrupted neuromasts (Figure 3D), but that hair-cell numbers had recovered after 2 hr (Figure 6B). The mechanism that mediates this recovery is not clear, but it is notable that a low level of hair-cell production normally occurs in lateral line neuromasts of larval zebrafish, as part of a process of ongoing turnover (Cruz et al., 2015; Williams and Holder, 2000). In the present study, we observed a small amount of proliferation in neuromasts of both mechanically-damaged and control fish. Since mechanical damage did not increase the level of cell proliferation in neuromasts relative to control (Figure 11), we believe this observed cell division is likely associated with the turnover process. Hair-cell regeneration in the vertebrate inner ear can also occur via direct phenotypic conversion of supporting cells into a replacement hair cells (Warchol, 2011). While it is conceivable that transdifferentiation of supporting cells could occur within 2 hr of mechanical injury, such transdifferentiation has not been previously demonstrated in zebrafish lateral line neuromasts (e.g. Thomas et al., 2015). Overall, these observations indicate that mechanical trauma does not increase the rate of hair-cell production within neuromasts, further supporting that neuromast recovery is largely due to hair-cell repair.

In summary, our data show that exposure of zebrafish lateral-line organs to strong water wave results in mechanical injury and loss of afferent synapses, but that these injuries rapidly recover. Our next steps will be to define the time course for synaptic recovery and to determine how lateral-line mediated behavior is affected by mechanically induced damage. Sub-lethal overstimulation of hair cells in the zebrafish lateral line provides a useful model for defining mechanisms of damage and inflammation and for identifying pathways that promote hair-cell repair following mechanically-induced injury.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Danio rerio)ABZIRCRRID: ZL1ZFIN ID: ZDB-GENO-960809–7
Strain, strain background (Danio rerio)TübingenZIRCRRID: ZIRC_ZL57ZFIN ID: ZDBGENO-990623–3
Genetic reagent (Danio rerio)lhfpl5bvo35/vo35Erickson et al., 2019RRID:ZIRC_ZL13656.05ZFIN ID: ZDB-GENO-200824–4
Genetic reagent (Danio rerio)TgBAC(neurod1:EGFP)Obholzer et al., 2008ZFIN ID: ZDB-ALT-080701–1
Genetic reagent (Danio rerio)Tg(myo6b:actb1-EGFP)Kindt et al., 2012ZFIN ID: ZDB-TGCONSTRCT-120926–1
Genetic reagent (Danio rerio)Tg(mpeg1:YFP)Roca and Ramakrishnan, 2013ZFIN ID: ZDB-ALT-130130–3
Sequence-based reagentlhfpl5b_ FErickson et al., 2019PCR primerGCGTCATGTGGGCAGTTTTC; Made by IDT
Sequence-based reagentlhfpl5b_RErickson et al., 2019PCR primerTAGACACTAGCGGCGTTGC; Made by IDT
Antibody(Ribbon label: Mouse monoclonal anti-Ribeye b IgG2a)Sheets et al., 2011N/A(1:10,000)
Antibody(Ribbon label: Mouse monoclonal anti-panCtBP IgG2a)Santa CruzCat. No. sc-55502(1:1000)
Antibody(PSD label: Mouse monoclonal anti-panMAGUK IgG1)NeuroMabK28/86, #75–029(1:500)
Antibody(Chicken polyclonal anti-GFP)Aves LabsCat. No. GFP-1020(1:500)
Antibody(Hair cell label: Rabbit polyclonal anti-Parvalbumin)Thermo FisherCat. No. PA1-933(1:500)
Antibody(Hair cell label: Rabbit polyclonal anti-Parvalbumin)AbcamCat. No. ab11427(1:2000)
Antibody(Hair cell label: Mouse anti-Otoferlin IgG2a)Developmental Studies Hybridoma BankHCS-1(1:500)
Antibody(Goat anti-Rabbit IgG Secondary Antibody, Pacific Blue)Thermo FisherCat. No. P-10994(1:400)
Antibody(Goat anti- Mouse IgG1 Antibody, Alexa Fluor 488)Thermo FisherCat. No. A-21121(1:1000)
Antibody(Goat anti-Chicken IgY Antibody, Alexa Fluor 488)Thermo FisherCat. No. A-11039(1:1000)
Antibody(Goat anti-Rabbit IgG Antibody, Dylight 549)Vector LaboratoriesCat. No. DI-1549–1.5(1:1000)
Antibody(Goat anti-Rabbit IgG Antibody, Alexa Fluor 555)Thermo FisherCat. No. A27039(1:1000)
Antibody(Goat anti- Mouse IgG2a Antibody, Alexa Fluor 647)Thermo FisherCat. No. A-21241(1:1000)
Peptide, recombinant proteinMluCINew England BiolabsCat. No. R0538
Chemical compound, drugDL-TBOATocrisCat. No.1223
Chemical compound, drugCopper(II) sulfate (CuSO4)Millipore SigmaCat. No. 451,657
Chemical compound, drug2.5 % Glutaraldehyde in 0.1 M Sodium Cacodylate Buffer, pH 7.4: SEMElectron Microscopy SciencesCat. No. 15,960
Chemical compound, drugParaformaldehyde; IHCMillipore SigmaCat. No. 158,127
Commercial assay or kitClick-iT EdU Cell Proliferation Kit for Imaging, Alexa Fluor 555 dyeThermo FisherCat. No. C10338
Software, algorithmFIJI is just ImageJNIHhttps://imagej.net/software/fiji/
Software, algorithmVolocityQuorum Technologieshttps://quorumtechnologies.com/index.php/component/content/category/31-volocity-software
Software, algorithmPrism (v9)Graphpad Softwarehttps://www.graphpad.com/
Software, algorithmAdobe IllustratorAdobehttps://www.adobe.com/
OtherFM1-43X; fixable analog of FM 1–43Thermo FisherCat. No. F353553 µM for 20 seconds
OtherDAPI nuclear stainThermo FisherCat. No. Cat. No. F353555 mg/ml stock; diluted (1:2000)

Zebrafish

All zebrafish experiments and procedures were performed in accordance with the Washington University Institutional Animal Use and Care Committee. Adult zebrafish were raised under standard conditions at 27–29°C in the Washington University Zebrafish Facility. Embryos were raised in incubators at 29 °C in E3 media 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgCl2; (Nüsslein-Volhard & Nüsslein-Volhard and Dahm, 2002) with a 14 hr:10 hr light:dark cycle. After four dpf, larvae were raised in 100–200 ml E3 media in 250 ml plastic beakers and fed rotifers daily. Sex of the animal was not considered in our studies because sex cannot be predicted or determined in larval zebrafish.

The transgenic lines TgBAC(neurod1:EGFP) (Obholzer et al., 2008), Tg(tnks1bp1:EGFP) (Behra et al., 2012), Tg(–6myo6b:βactin-EGFP) (Kindt et al., 2012), and Tg(mpeg1:YFP) (Roca and Ramakrishnan, 2013) were used in this study. Fluorescent larvae were identified at three dpf without anesthesia in E3 media. The mutant line lhfpl5bvo35/vo35 was also used (Erickson et al., 2019).

Genotyping

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To genotype lhfpl5bvo35/vo35 larvae and siblings after mechanical stimulation and immunohistochemical labeling, ~ 1 mm tail tissue was excised, and genomic DNA was extracted by incubation in a lysis buffer (10 mM Tris pH 8.0, 50 mM KCl, 0.3% NP-40, 0.3 % Tween-20). A genomic region of lhfpl5b was amplified by PCR using forward primer GCGTCATGTGGGCAGTTTTC and reverse primer TAGACACTAGCGGCGTTGC. The lhfpl5bvo35 mutation disrupts a MluCI restriction site (AATT), so PCR products were digested with MluCI, and homo- and heterozygotes were resolved by differences in band size on a 1–1.5% agarose gel.

Experimental apparatus

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Multi-well plates containing larvae were clamped to a custom magnesium head expander (Vibration & Shock Technologies, Woburn, MA) on a vertically oriented Brüel + Kjær LDS Vibrator, V408 (Brüel and Kjær, Naerum, Denmark). An additional metal plate was fitted to the bottom of the multi-well dish to fill a small gap between the bottoms of the wells and the head expander to eliminate flexing of the well plate relative to the head expander. Vibrometry of the well bottoms and the head expander with a laser-Dopper vibrometer (OFV-2600 and OFV-501, Polytec, Irvine, CA) confirmed that the well plate and head expander motion were equal at stimulus frequencies. This experimental apparatus was housed in a custom sound-attenuation chamber. An Optiplex 3,020 Mini Tower (Dell) with a NI PCIe-6321, X Series Multifunction DAQ (National Instruments) running a custom stimulus generation program (modified version of Cochlear Function Test Suite) was used to relay the stimulus signal to a Brüel + Kjær LDS PA100E Amplifier that drove a controlled 60 Hz vibratory stimulus along the larvae’s dorsoventral axis (vertically). Two accelerometers (BU-21771, Knowles, Itasca, IL) were mounted to the head expander to monitor the vertical displacement of the plate. The output of the accelerometers was relayed through a custom accelerometer amplifier (EPL Engineering Core). A block diagram for the EPL Lateral Line Stimulator can be found here: https://www.masseyeandear.org/research/otolaryngology/eaton-peabody-laboratories/engineering-core.

Mechanical overstimulation of lateral-line organs in free swimming larvae

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At seven dpf, free-swimming zebrafish larvae were placed in untreated six-well plates (Corning, Cat# 3736; well diameter: 34.8 mm; total well volume: 16.8 ml) with 6 ml E3 per well, pre-warmed to 29 °C. Up to 15 larvae were placed in each well. Individual wells were sealed with Glad Press ‘n Seal plastic food wrap prior to placing the lid on the plate. An additional metal plate was fitted to the bottom of the multi-well dish to fill a small gap between the bottoms of the wells and the head expander.

Mechanical water displacement exposures (stimulus parameters: 60 Hz, 40.3 ± 0.5 m/s2) were conducted at room temperature (22°C–24°C) up to 2 hr after dawn. The frequency selected for mechanical overexposure of lateral-line organs was based on previous studies showing 60 Hz to be within the optimal upper frequency range of mechanical sensitivity of superficial posterior lateral-line neuromasts (Weeg et al., 2002; Trapani et al., 2009, Levi et al., 2015). To confirm that 60 Hz was the optimal frequency to induce damage, we tested 45, 60, and 75 Hz at comparable intensities. We observed at 75 Hz no apparent damage to lateral line neuromasts while 45 Hz at a comparable intensity proved toxic that is it was lethal to the fish.

Exposures consisted of 20 min of stimulation followed by a 10-min break and 2 hr of uninterrupted stimulation. We also tested periodic exposures that consisted of a series of short pulses spanning 2 hr total: 2 20 min exposures each followed by 10 min of rest, followed by 30 min of stimulation, a 10 min break, and a final 20 min of stimulation. During the entire duration of exposure, unexposed control fish were kept in the same conditions as exposed fish i.e. placed in a multi-well dish and maintained in the same room as the exposure chamber. For experiments pharmacologically blocking glutamate uptake, fish were co-exposed to 10 µM DL-TBOA (Tocris; Cat. No.1223) + 0.1 % DMSO or 0.1 % DMSO alone. After exposure, larvae were either immediately fixed for histology, prepared for live imaging, or allowed to recover for up to 2 days in an incubator at 29 °C.

Ablation of lateral-line organ with CuSO4

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Free-swimming larvae were exposed to freshly made 3 µM CuSO4 solution in E3 for 1 hr, then rinsed and allowed to recover for 2 hr to ensure complete ablation of the lateral-line neuromasts. Neuromast ablation was confirmed by immunofluorescent labeling of hair cells. The effects of low-dose copper exposure are likely specific to lateral-line organs; a previous study in zebrafish determined exposure to low-dose CuSO4 for 1 hr did not alter the acoustic escape response, which is similar to the fast start response we observed but evoked by higher frequency stimulation (100–500 Hz) of the anterior macula of inner ear (Buck et al., 2012).

Fast-start escape response behavior assay

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Images of larval swimming behavior (1000 frames per second) were acquired with an Edgertronic SC1 high-speed camera (Sanstreak Corp). Image acquisition began 10 s following stimulus onset. All subsequent analysis was performed using ImageJ. To track swimming behavior, images were initially stabilized using the Image Stabilizer Plugin. In stabilized images, the position of individual larval heads (located via the pigmented eyes) in each frame were tracked using the Manual Tracking Plugin. Larvae were tracked over 10 s (10,000 frames total) per trial. ‘Fast start’ responses—defined as a c-bend of the body occurring within 15 ms followed by a counter-bend— were identified manually.

Whole-mount immunohistochemistry

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For visualization of zebrafish lateral-line hair cells, neurons, and synapses: 7–9 dpf larvae were briefly sedated on ice, transferred to fixative (4 % paraformaldehyde, 1 % sucrose, 37.5 µM CaCl2, 0.1 M phosphate buffer) in a flat-bottomed 2 ml Eppedorf tubes, and fixed for 5 hr at 4–8°C. Fixed larvae were permeabilized in ice-cold acetone for 5 minutes, then blocked in phosphate-buffered saline (PBS) with 2 % goat serum, 1 % bovine serum albumin (BSA), and 1 % DMSO for 2–4 hours at room temperature (RT; 22-24 oC). Larvae were incubated with primary antibodies diluted in PBS with 1 % BSA and 1 % DMSO overnight at 4–8°C, followed by several rinses in PBS/BSA/DMSO and incubation in diluted secondary antibodies conjugated to Pacific Blue (1:400), Alexa Fluor 488 (1:1000), Alexa Fluor 555 (1:1000), Alexa Fluor 647 (1:1000; Invitrogen), or DyLight 549 (1:1000; Vector Laboratories) for 2 hr at RT. In some experiments, fixed larvae were stained with 2.5 ug/ml 4′,6-diamidino-2-phenylindole (DAPI; Invitrogen) diluted in PBS to label all cell nuclei. Larvae were mounted on glass slides with elvanol (13% w/v polyvinyl alcohol, 33% w/v glycerol, 1% w/v DABCO (1,4 diazobicylo[2,2,2] octane) in 0.2 M Tris, pH 8.5) and #1.5 cover slips.

For visualization of inflammation and macrophage recruitment: 7 dpf larvae were sedated on ice, transferred to 4 % paraformaldehyde fixative in PBS, then fixed overnight at 4°C–8°C. The next day larvae were rinsed in PBS and blocked in PBS with 5 % normal horse serum (NHS), 1 % DMSO, and 1 % Triton x-100 for 2 hr at RT. Larvae were incubated with primary antibodies diluted in PBS with 5 % NHS and 1 % Triton-x 100 overnight at RT, rinsed several times in PBS, then incubated in diluted secondary antibodies listed above for 2 hr at RT. Larvae were mounted on glass slides with glycerol/PBS (9:1); coverslips were sealed with clear nail polish.

Primary antibodies

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The following commercial antibodies were used in this study: GFP (1:500; Aves Labs, Inc; Cat# GFP-1020), Parvalbumin (1:2000; Thermo Fisher; Cat# PA1-933), Parvalbumin (1:2000; Abcam; Cat# ab11427), Parvalbumin (1:500; Sigma-Aldrich; Cat# P3088), MAGUK (K28/86; 1:500; NeuroMab, UC Davis; Cat# 75–029), Otoferlin (1:500; Developmental Studies Hybridoma Bank/ HCS-1). Custom affinity-purified antibody generated against Danio rerio Ribeye b (mouse IgG2a; 1:2000; Sheets et al., 2011) was also used.

Hair-cell labeling

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To selectively label hair-cell nuclei, live zebrafish larvae were incubated with DAPI (5 mg/ml) diluted 1:2000 in E3 media for 4 min. Larvae were briefly rinsed three times in fresh E3 media, then immediately exposed to mechanical overstimulation.

Live imaging of FM1-43 did not provide the temporal resolution needed to compare relative uptake and fast recovery 0–4 hr following exposure. We therefore examined FM1-43 uptake using the fixable analogue. To label with FM1-43X (n-(3-triethylammoniumpropyl)–4-(4-(dibutylamino)-styryl) pyridinium dibromide; ThermoFisher), free-swimming larvae were exposed to FM 1–43 FX at 3 µM for 20 seconds, then rinsed three times in fresh E3 as previously described (Toro et al., 2015) and immediately fixed (4 % paraformaldehyde, 4 % sucrose, 150 µM CaCl2, 0.1 M phosphate buffer). FM 1–43 FX mean signal intensity from maximum projection images was calculated using ImageJ as the integrated pixel intensity divided by the area of the neuromast region of interest. We verified that relative labeling of hair cells at 1–3 hours appeared comparable between live FM1-43 and FM 1–43 FX. We also verified loss of FM 1–43 FX uptake in larvae following brief treatment with 5 mM BAPTA to disrupt tip links (Kindt et al., 2012). Following fixation, stereocilia were labeled by incubation with phalloidin conjugated to Alexa Fluor 488 (Invitrogen) at 66 µM in PBS, washed, and mounted on slides with elvanol.

EdU labeling and quantification

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To label proliferating cells, we used the Click-iT EdU Cell Proliferation Kit for Imaging, Alexa Fluor 555 dye (Invitrogen). Following mechanical overstimulation, larvae were incubated in 500 µM EdU with 0.5 % DMSO in E3 for 4 hr at 28 °C then fixed in 4 % paraformaldehyde in PBS overnight at 4 °C. Larvae were washed in 3 % bovine serum albumin in PBS then permeabilized with 0.5 % Triton-X in PBS. GFP signal in Tg[tnks1bp1:GFP] fish was amplified using anti-GFP primary antibody (Aves), followed by a secondary antibody conjugated to Alexa Fluor 488. The EdU detection reaction was performed according to manufacturer guidelines; larvae were incubated in a reaction cocktail (4 mM CuSO4 and Alexa Fluor 555 azide in 1 X Click-iT reaction buffer with reaction buffer additive) for 1 hr at 25 °C. Larvae were washed, counterstained with Hoechst 33342, and mounted on slides with elvanol. Confocal images of neuromasts were acquired using an LSM 700 laser scanning confocal microscope with a 63 × 1.4 NA Plan-Apochromat oil-immersion objective (Carl Zeiss). The numbers of EdU+ cells per neuromast were quantified in ImageJ.

Confocal imaging

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Images of fixed samples were acquired using an LSM 700 laser scanning confocal microscope with a 63 × 1.4 NA Plan-Apochromat oil-immersion objective (Carl Zeiss). Confocal stacks were collected with a z step of 0.3 µm over 7–10 µm with pixel size of 100 nm (x-y image size 51 × 51 µm). Acquisition parameters were established using the brightest control specimen such that just a few pixels reached saturation in order to achieve the greatest dynamic range in our experiments. These parameters including gain, laser power, scan speed, dwell time, resolution, and zoom, were kept consistent between comparisons.

Confocal image processing and analysis

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All analysis was performed on blinded images. Digital images were processed using ImageJ software (Schneider et al., 2012). In order to quantitatively measure sizes and fluorescent intensities of puncta, raw images containing single immunolabel were subtracted for background using a 20-pixel rolling ball radius and whole neuromasts were delineated from Parvalbumin-labeled hair cells using the freehand selection and ‘synchronize windows’ tools. Puncta were defined as regions of immunolabel with pixel intensity above a determined threshold: threshold for Ribeye label was calculated using the Isodata algorithm (Ridler and Calvard, 1978) on maximum-intensity projections, threshold for MAGUK label was calculated as the product of 7 times the average pixel intensity of the whole neuromast region in a maximum-intensity projection. Particle volume and intensity were measured using the 3D object counter (Bolte and Cordelières, 2006) using a lower threshold and a minimum size of 10 voxels. To normalize for differences in staining intensity across experimental trials, all volumes were divided by the median control volume in each trial for each individual channel. The number of particles above lower threshold was quantified using the ImageJ Maximum Finder plugin with a noise tolerance of 10 on maximum-intensity projections. Intact synapses were manually counted and defined as adjoining or overlapping maxima of Ribeye and MAGUK labels. The number of synapses per hair cell was approximated by dividing the number of intact synapses within a neuromast by the number of hair cells in the neuromast. Innervation of neuromast hair cells was quantified during blinded analysis by scrolling through confocal z-stacks of each neuromast (step size 0.3 µm) containing hair cell and afferent labeling and identifying hair cells that were not directly contacted by an afferent neuron i.e. no discernable space between the hair cell and the neurite. Hair cells that were identified as no longer innervated showed measurable neurite retraction; there was generally >0.5 µm distance between a retracted neurite and hair cell. Stereocilia length measurements were obtained in ImageJ from interpolated 3D projections of z-stack images (step size 0.3 µm) containing phalloidin labeled hair bundles. Three independent measurements were obtained from the base to the tips of hair bundles at center of each neuromast, and the average length was calculated.

Quantitative data on macrophage response to mechanical injury were collected from the two caudal-most (‘terminal’) neuromasts. Confocal image stacks were obtained using a Zeiss LSM700 microscope and visualized using Volocity software. These image stacks were used to derive three metrics from each neuromast. First, the number of macrophages within 25 µm of a particular neuromast was determined by inscribing a circle of 25 µm radius, centered on the neuromast, and counting the number of macrophages that were either fully or partially enclosed by this circle. Next, the number of macrophages contacting a neuromast was determined by scrolling through the x-y planes of each image stack (1 µm interval between x-y planes, 15 µm total depth) and the counting macrophages that were in direct contact with Otoferlin-labeled hair cells. Finally, the number of macrophages that had internalized Otoferlin-labeled material (hair-cell debris) were counted and were assumed to reflect the number of phagocytic events. For each metric, the recorded number reflected the activity of a single macrophage, that is, a macrophage that made contacts with multiple hair cells and/or had internalized debris from several hair cells was still classified as a single ‘event.’

Subsequent image processing for display within figures was performed using Photoshop and Illustrator software (Adobe).

Scanning electron microscopy

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To image hair-cell bundles, zebrafish larvae were exposed to strong water wave stimulus, then anesthetized in 0.12 % tricaine in E3 and immediately fixed in 2.5 % glutaraldehyde in 0.1 M sodium cacodylate buffer (Electron Microscopy Sciences) supplemented with 2 mM CaCl2. Larvae were shipped overnight in fixative, then most of the fixative (~90–95%) was removed, replaced with distilled water, and samples were stored at 4 C. Next, larvae were washed in distilled water (Gibco), dehydrated with an ascending series of ethanol, critical point dried from liquid CO2 (Tousimis Aurosamdri 815), mounted on adhesive carbon tabs (Ted Pella), sputter coated with 5 nm of platinum (Leica EM ACE600), and imaged on Hitachi S-4700 scanning electron microscope. Kinocilia diameter measurements were performed using ImageJ.

Statistical analysis

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A hierarchical linear model analysis with fish used as random effects was used to compare each of the measures between the conditions and groups. Akaike Information Criterion (AIC) and Bayessian Information Criterion (BIC) were used to identify the best fitted covariance structure. Tukey’s adjustment was used for the alpha level to avoid type I error inflation due to multiple comparisons. Estimated marginal mean differences and 95 % Confidence Intervals around them were explored and reported for quantification of effect size for group differences. Graphs for data visualization and additional statistical analyses were performed Prism 8 (Graphpad Software Inc). Mixed model analysis was used to compare time-series data. Statistical significance between synaptic ribbon and PSD volumes with was determined by Kruskal-Wallis test (one independent variable) or Mann–Whitney U test (one independent variable) and appropriate post-hoc tests. Based on the variance and effect sizes reported in previous studies, the number of biological replicates were suitable to provide statistical power to avoid Type II error (Sebe et al., 2017; Uribe et al., 2018).

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 2, 3, 4, and 7.

References

  1. Book
    1. Coffin AB
    2. Kelley MW
    3. Manley GA
    4. Popper AN
    (2004) Evolution of sensory hair cells
    In: Manley GA, Fay RR, Popper AN, editors. Evolution of the Auditory System. New York: Springer-Verlag. pp. 55–94.
    https://doi.org/10.1007/978-1-4419-8957-4_3
  2. Book
    1. Nüsslein-Volhard C
    2. Dahm R
    (2002)
    Zebrafish: A Practical Approach (1st edn)
    Oxford University Press.

Decision letter

  1. Doris K Wu
    Reviewing Editor; NIDCD, NIH, United States
  2. Didier YR Stainier
    Senior Editor; Max Planck Institute for Heart and Lung Research, Germany

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

Mechanical insults can lead to sensory hair cell death but the underlying mechanisms are not well understood. Here, Holmgren et al. describes for the first time, a mechanical hair-cell damage model in the zebrafish lateral line system, which allowed a closer examination of hair cell response to mechanical injury and recovery.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "Mechanical overstimulation causes acute injury followed by fast recovery in lateral-line neuromasts of larval zebrafish" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below. Although all three reviewers agreed that the study was thorough but felt the findings do not provide sufficient advances to be considered further for publication in eLife.

Reviewer #1:

In the manuscript titled "Mechanical overstimulation causes acute injury followed by fast recovery in lateral-line neuromasts of larval zebrafish" by Holmgren et al., the authors develop a method to overstimulate hair cells and determine some of the consequences of this overstimulation. The overarching goal of this work is to develop a model for noise-induced hair-cell damage in the zebrafish. The authors use the lateral line for their studies and stimulate hair cells using an electrodynamic shaker which generate significant aqueous agitation. The authors demonstrate physical damage to hair cells of the lateral line that are dependent on position of the neuromast. The damage includes alteration of afferent synapses, afferent neurite retraction, limited damage to hair bundles and a decrease in mechanotransduction. After damage, they show macrophage recruitment and quick recovery of hair cell neuromasts, which is surprising.

The paper is interesting in that it brings a new capacity to the zebrafish animal model: mechanical overstimulation of the hair cell. Tempering this is a general feeling that the authors do not dig deep enough in the current form of the manuscript, but this could be remedied. More specifically, the authors are making a model in zebrafish for noise-induced damage, so they need to show that this model is similar to mammals in the way hair cells are damaged. This is done in the manuscript, but it is limited and should be expanded as suggested below.

– The authors use a vertically-oriented Brüel+Kjær LDS Vibrator to deliver a 60 Hz vibratory stimulus to damage lateral line hair cells. It is not made clear on why this frequency was selected. Did the authors choose this frequency because they screened a number of frequencies, and this is the one that did the most damage to hair cells or was it chosen for another reason? Or do all frequencies do the same amount of damage? The authors should screen a number of frequencies and choose the stimulus that does the most damage to hair cells. This would set the field in the best direction, should members of the community attempt this new technique. It is not necessary to repeat all of the experiments, but the authors should show which frequencies are best for inducing damage.

– The SEM images of the hair bundle are beautiful and do show damage to the hair bundle, but historically speaking older studies in mammals have shown that the actin core of the stereocilia is damaged. It would be critical to know if this was the case. Showing damage to the kinocilium and stereocilia splaying is a start, but readers of eLife would need to know if the actin cores are damaged. So, TEM should be used to find damage to the actin cores of stereocilia.

– I think the use of "Noise-exposed lateral line" as a term for mechanically overstimulated lateral line hair cells is not correct and could be misleading. The lateral line senses water motion, not sounds as the word noise would imply. Calling the stimulus "noise" should be removed throughout.

– Decreases in mechanotransduction are shown by dye entry. These results should be strengthened using microphonic potentials to determine the extent of damage. This experiment is not necessary but would improve the quality of the document.

– In figure 2, PSD labeling is not clear.

Reviewer #2:

Holmgren et al. describe the development of a model for hair cell noise damage using the zebrafish lateral line line system. Using an electrodynamic shaker, the authors induce quantifiable damage and death of hair cells after a two-hour treatment. They describe gross morphological changes of hair cells, changes in innervation and synapse distribution. In addition, they describe disruption of stereocilia and kinocilia, as well as reduced mechanotransduction-dependent uptake of FM1-43 dye. Damage is no longer detectable several hours after insult, demonstrating recovery.

1. While the findings are carefully measured and described the effects of insult on hair cells are relatively minor, with a change in hair cell number, extent of innervation or synapses per hair cell (Figures3 and 4) in the range of 10% reduction compared to control. One potential value of the model would be to use it to discover underlying pathways of damage or screen for potential therapeutics. However, with these modest changes it is not clear that there will be enough power to determine effects of potential interventions.

2. The most dramatic phenotype after shaking is a physical displacement of hair cells, described as disrupted morphology. However, it is not clear what the underlying cause of this change. Are only posterior neuromasts damaged in this way? Is it a wounding response as animals are exposed to an air interface during shaking? It is also not clear to what extent this displacement reveals more general principles of the effects of noise on hair cells. Additional discussion of underlying causes would be welcome.

3. Because afferent neurons innervate more than one neuromast and more than one hair cell per neuromast, measurements of innervation of neuromasts (Figure 3) or synapses per hair cell (Figure 4) cannot be assumed to be independent events. That is, changes in a single postsynaptic neuron may be reflected across multiple synapses, hair cells, and even neuromasts. This needs to be accounted for in experimental design for statistical analysis.

4. The SEM analysis provides compelling snapshots of apical damage but could be supplemented by quantitative analysis with antibody staining or transgenic lines where kinocilia are labeled. The amount of reduced FM1-43 labeling is one of the more dramatic effects of the shaking insult, suggesting widespread disruption to mechanotransduction that could be related to this apical damage. Further examination of the recovery of mechanotransduction would be interesting.

5. A previous publication by Uribe et al.2018 describes a somewhat similar shaking protocol with somewhat different results – more long-lasting changes in hair cell number, presynaptic changes in synapses, etc. It would be worth discussing potential differences across the two studies.

Reviewer #3:

Holmgren et al. describe a novel model of reversible mechanical damage to zebrafish neuromast hair cells. The authors demonstrate that when zebrafish are exposed to strong currents, neuromast morphology, hair cell number, innervation, and MET function suffer various types and degrees of damage, from which the NMs recover within 2 days. Additionally, they show macrophage recruitment to damaged neuromasts, where they may be phagocytosing synaptic debris. Based on various mechanistic and phenotypic commonalities (involvement of ROS, stereocilia and synapse phenotype), the authors argue that this model is a good approximation of noise-induced hair cell damage in mammals.

Overall impact:

This reviewer agrees that a "noise" damage model in the zebrafish would be a powerful tool to better understand the mechanisms underlying noise-induced hearing loss. However, due to various weaknesses of the data (detailed below), the main claims of the paper are not sufficiently supported. In addition, noise-induced hearing loss has been previously modeled in the zebrafish model. The present model, therefore, does not provide a significant methodological innovation. Based on this, and the fact that addressing all the concerns listed below likely exceeds the scope of a reasonable revision, this manuscript is believed to lack the impact and novelty to be recommended for publication in eLife.

– As the authors point out, zebrafish hair cells can be regenerated. With that in mind, and to make the relevance for mammalian hair cell repair clear, a clear distinction between mechanisms mediated by "repair" or "regeneration" needs to be made. The authors discuss that proliferative hair cell generation can be excluded based on the short time period, but suggest that transdifferentiation might be involved. Recovery of NM hair cell number occurs within the same 2 hour period in which NM morphology and hair cell function improved, making it difficult to determine the extent to which "regeneration" contributed to the recovery. The amount of transdifferentiation has to be shown experimentally (lineage tracing?).

– The classification of "normal" vs "disrupted" is vague and not quantitative. The examples shown in the paper seem to be quite clear-cut, but this reviewer doubts that was the case throughout all analyzed samples. Formulate clear benchmarks and criteria for the disrupted phenotype (even when blind analysis is performed).

– Sustained and periodic exposure: These two exposure protocols not only differ with respect to sustained vs periodic, they also differ in total exposure time (Figure 2B). This complicates the interpretation, especially considering the authors own finding that a pre-exposure is protective.

– The data on the mitochondrial ROS aspect seems not well integrated into the overall story.

– It is surprising that the hair bundle morphology was not assessed after recovery. This is crucial. Overall, it would be good to see some quantification of the SEM data, e.g. kinocilia length and number of splayed bundles.

– Behavioral recovery (measured as number of "fast start" responses) was also not assessed. This is essential for determining the functional relevance of the recovery.

– This reviewer is not yet convinced that this damage model displays enough commonalities to mammalian noise damage to justify the ubiquitous use of the term "noise" throughout the manuscript. It would be more prudent to use a more careful term along the lines of "mechanical overstimulation-induced damage".

– Overall, there was a lack of experimental and analysis detail in the Results section. For example, how was afferent innervation quantified? Just counting GFP labeled contacts to hair cells? There was also inconsistency in the use of two variations of the mechanical damage protocol, the time points at which repair was assessed, and whether the damage was quantified in all neuromasts or in normal vs. disrupted neuromasts separately, making the data difficult to interpret.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Mechanical overstimulation causes acute injury and synapse loss followed by fast recovery in lateral-line neuromasts of larval zebrafish" for further consideration by eLife. Your revised article has been evaluated by Didier Stainier (Senior Editor) and a Reviewing Editor.

All three reviewers appreciated the fact that you have made a good effort in revising the manuscript. However, the story has also changed from a noise-induced damage to a mechanical over-stimulation paradigm. While the Ihfp15b mutant results are interesting, it is not entirely clear how these results are applicable to noise-induced hearing loss in mammals. A point that could be better addressed in the Discussion. After an extensive discussion with the three reviewers, they have consolidated their main suggestions as follow:

Essential revisions

1) It would be important to determine whether the hair cells with severe bundle damage recover their bundle morphology or are replaced by transdifferentiation of SCs.

2) Statistical analyses of results should be incorporated per comments from Reviewer#2. It is important to know if the conclusions hold up across total neuromasts and fish. In this regard, this is a reference suggested by the Reviewer: Aarts E, Verhage M, Veenvliet JV, Dolan CV, van der Sluis S. A solution to dependency: using multilevel analysis to accommodate nested data. Nat Neurosci. 2014 Apr;17(4):491-6.

Reviewer #1:

The authors embark on a comprehensive characterization of damages suffered by the zebrafish lateral line organ in response to mechanical overstimulation. The ultimate goal is to establish this as a novel model for noise-induced hearing loss. The study describes significant overlap in the damage characteristics between mammals and fish, justifying this protocol as a (limited) but very useful model (due to the well-known advantages of the zebrafish system) to study the basic mechanisms of hair cell damage incurred by mechanical overstimulation.

The revision includes some valuable additions in response to the previous review, but many suggestions were deemed intractable in the given time period. The manuscript is therefore improved, but not to a level that this reviewer judges had hoped for.

– The quantification of changes in SEMs of hair bundle morphology following damage was a significant improvement from the original submission, but imaging or quantification of recovery, which would be an important addition to the manuscript, is still lacking. It would also be nice to see a quantification of splayed bundles and kinocilia length before and after recovery. The confocal images demonstrating the changes to the kinocilium following damage were a nice addition and might provide an easier way to quantify changes during the recovery process.

– The authors decided that further investigation into the distinction of repair vs recovery (beyond SC EdU incorporation) is outside the scope of this study, but this reviewer thinks that it should have been further explored. The authors claim that they only see a modest increase in SC EdU incorporation (>0.5 SC/neuromast) following injury, but the decrease in HC number is also small (looks to be <1 HC/neuromast from graph). Directly addressing transdifferentiation of SCs through lineage tracing techniques would be informative.

Reviewer #2:

I appreciate the extensive revisions Holmgren et al. have made to their previous manuscript, addressing many of the criticisms of the previous review. They include a clever new experiment using lhfpl5b mutants that have mechanosensory deficits specific to lateral line hair cells. Using these mutants, they demonstrate that the morphological disruption they observe after shaker treatment occurs in these mutants to the same degree as in wt fish.

However, this result does call into question whether the model is a functional one for noise damage. If mechanotransduction is not necessary, are the phenotypes observed really excitotoxicity? Do the reported changes in innervation and synapses also not require functional mechanotransduction?

I fear I was not clear enough on my previous point (#3 in response to reviewer 2):

Because afferent neurons innervate more than one neuromast and more than one hair cell per neuromast, measurements of innervation of neuromasts (Figure 3) or synapses per hair cell (Figure 4) cannot be assumed to be independent events. That is, changes in a single postsynaptic neuron may be reflected across multiple synapses, hair cells, and even neuromasts. This needs to be accounted for in experimental design for statistical analysis.

As measurements of innervation or synapse density are not independent, they cannot be treated as distinct instances (n's) in the statistical analysis. In these experiments the independent instances are going to be the number of fish, not hair cells. This issue extends to analysis in Figures7, 8 as well.

I am somewhat confused about the conclusions from figure 4. Setting aside the issue of independence, the differences in synapses per hair cell are variably measured as significant or insignificant, suggesting that the analysis is underpowered. In addition, shouldn't the relevant comparisons in Figure 4E be between DMSA and TBOE, that is comparing each of control, shaken normal and shaken disrupted to the corresponding drug treatment?

Reviewer #3:

The goal of the manuscript is to present a new paradigm for damaging hair cells of the zebrafish lateral line using vibrational stimuli. A successful method would be important to the community as this goal has been difficult to achieve, and it would open up new avenues of research. The authors addressed some of my concerns from the last review but not all. The experiments they did perform are a screen of frequencies that damage the hair cells of the lateral line and improved the quality of an image (Figure 4). This is appreciated especially during the time of Covid.

They argue that 2 suggested experiments are not necessary: microphonic potentials and damage to the core of actin of stereocilia. I feel that they would improve the paper's quality, but I see their point about not including the microphonic potentials; however, they should at some level look at the damage to the actin core. This could be by TEM as suggested or it could be by phalloidin labeling. Both of these methods can be used to see gaps in the core of actin in mouse hair cells and maybe those of zebrafish.

Another issue is the use of the word "current". According to the dictionary (OED) current is defined as "That which runs or flows, a stream; spec. a portion of a body of water, or of air, etc. moving in a definite direction." This is the way I see the word used in the scientific literature as well. I don't think what the authors are providing is a current. They should use a different term, perhaps, "vibratory stimulus" or "water stimulus."

https://doi.org/10.7554/eLife.69264.sa1

Author response

[Editors’ note: the authors resubmitted a revised version of the paper for consideration. What follows is the authors’ response to the first round of review.]

Reviewer #1:

In the manuscript titled "Mechanical overstimulation causes acute injury followed by fast recovery in lateral-line neuromasts of larval zebrafish" by Holmgren et al., the authors develop a method to overstimulate hair cells and determine some of the consequences of this overstimulation. The overarching goal of this work is to develop a model for noise-induced hair-cell damage in the zebrafish. The authors use the lateral line for their studies and stimulate hair cells using an electrodynamic shaker which generate significant aqueous agitation. The authors demonstrate physical damage to hair cells of the lateral line that are dependent on position of the neuromast. The damage includes alteration of afferent synapses, afferent neurite retraction, limited damage to hair bundles and a decrease in mechanotransduction. After damage, they show macrophage recruitment and quick recovery of hair cell neuromasts, which is surprising.

The paper is interesting in that it brings a new capacity to the zebrafish animal model: mechanical overstimulation of the hair cell. Tempering this is a general feeling that the authors do not dig deep enough in the current form of the manuscript, but this could be remedied. More specifically, the authors are making a model in zebrafish for noise-induced damage, so they need to show that this model is similar to mammals in the way hair cells are damaged. This is done in the manuscript, but it is limited and should be expanded as suggested below.

– The authors use a vertically-oriented Brüel+Kjær LDS Vibrator to deliver a 60 Hz vibratory stimulus to damage lateral line hair cells. It is not made clear on why this frequency was selected. Did the authors choose this frequency because they screened a number of frequencies, and this is the one that did the most damage to hair cells or was it chosen for another reason? Or do all frequencies do the same amount of damage? The authors should screen a number of frequencies and choose the stimulus that does the most damage to hair cells. This would set the field in the best direction, should members of the community attempt this new technique. It is not necessary to repeat all of the experiments, but the authors should show which frequencies are best for inducing damage.

The frequency selected for mechanical overexposure of lateral-line organs was based on previous studies showing 60 Hz to be within the optimal upper frequency range of mechanical sensitivity of superficial posterior lateral-line neuromasts, with maximal response between 10-60 Hz, but a suboptimal frequency for hair cells of the anterior macula in the ear (Weeg and Bass 2002, Trapani et al., 2009, Levi et al., 2015). To confirm that 60 Hz was the optimal frequency to induce damage, we tested 45, 60, and 75 Hz at comparable intensities. We observed at 75 Hz no apparent damage to lateral line neuromasts while 45 Hz at a comparable intensity proved toxic i.e. it was lethal to the fish. We have updated the Results and Method Details to include our rationale for choosing 60 Hz.

– The SEM images of the hair bundle are beautiful and do show damage to the hair bundle, but historically speaking older studies in mammals have shown that the actin core of the stereocilia is damaged. It would be critical to know if this was the case. Showing damage to the kinocilium and stereocilia splaying is a start, but readers of eLife would need to know if the actin cores are damaged. So, TEM should be used to find damage to the actin cores of stereocilia.

Our main goal of this initial manuscript was to survey morphological and functional changes in mechanically injured lateral line organs with an emphasis on inflammation and synapse loss. We agree TEM studies showing damage to the actin core of the stereocilia will be important to determine whether mechanical damage to neuromast hair bundles fully mimics mammalian stereocilia damage, but these experiments will require significant time to perform and optimize. We have expanded our analysis of hair-bundle morphology in this study and intend to pursue deeper analysis of hair bundle damage, i.e. examination of the stereocilia actin core, in future follow-up studies.

– I think the use of "Noise-exposed lateral line" as a term for mechanically overstimulated lateral line hair cells is not correct and could be misleading. The lateral line senses water motion, not sounds as the word noise would imply. Calling the stimulus "noise" should be removed throughout.

We have removed the term “noise” throughout the manuscript and replaced it with either “strong water current stimulus” or “mechanical overstimulation” where appropriate.

– Decreases in mechanotransduction are shown by dye entry. These results should be strengthened using microphonic potentials to determine the extent of damage. This experiment is not necessary but would improve the quality of the document.

While we agree that microphonic recordings would provide further support for reduced mechanotransduction, quantitative FM1-43 uptake in zebrafish lateral line hair cells is a well-established proxy for microphonic measurements. In a previous study using the same protocol utilized in our manuscript, FM1-43 labeling intensity was shown to directly correspond with microphonic amplitude (Toro et al., 2015). Moreover, the fixable analogue of FM1-43 (FM143FX) gave us comparable relative measurements of uptake as live FM1-43 and provided the additional advantage of high temporal resolution and the ability to simultaneously assay entire cohorts of control and overstimulated fish (which is not possible with microphonic measurements or live FM1-43 imaging), as we could expose groups of fish briefly to the dye at determined time intervals following overstimulation, then immediately place in fixative.

– In figure 2, PSD labeling is not clear.

We assume the reviewer meant PSD labeling in Figure 4 and we agree it is difficult to discern. We have changed the hair-cell label from gray to blue in the images so that the green PSD labeling is clear.

Reviewer #2:

Holmgren et al. describe the development of a model for hair cell noise damage using the zebrafish lateral line line system. Using an electrodynamic shaker, the authors induce quantifiable damage and death of hair cells after a two-hour treatment. They describe gross morphological changes of hair cells, changes in innervation and synapse distribution. In addition, they describe disruption of stereocilia and kinocilia, as well as reduced mechanotransduction-dependent uptake of FM1-43 dye. Damage is no longer detectable several hours after insult, demonstrating recovery.

1. While the findings are carefully measured and described the effects of insult on hair cells are relatively minor, with a change in hair cell number, extent of innervation or synapses per hair cell (Figures3 and 4) in the range of 10% reduction compared to control. One potential value of the model would be to use it to discover underlying pathways of damage or screen for potential therapeutics. However, with these modest changes it is not clear that there will be enough power to determine effects of potential interventions.

One advantage of the zebrafish model is the ability to overstimulate large cohorts of larvae, thereby providing enough power to uncover modest but significant changes resulting from moderate damage to hair cells. While not as well suited for unbiased large-scale screens of therapeutics, our overexposure protocol provides the opportunity to determine the role of specific cellular pathways (e.g. metabolic stress, inflammation, and glutamate excitotoxicity) in hair-cell damage and synapse loss following mechanically-induced damage via genetic or pharmacological manipulation of these pathways. Additionally, as the hair cell synapses fully repair following stimulus-induced loss, the zebrafish model has the potential for identifying novel pathways for repair through transcriptomic profiling (for an example, see Mattern et al., Front. Cell Dev. Biol., 2018). Cumulatively, these future experimental directions will provide important mechanistic information that could be used toward the development of targeted therapeutic interventions.

2. The most dramatic phenotype after shaking is a physical displacement of hair cells, described as disrupted morphology. However, it is not clear what the underlying cause of this change. Are only posterior neuromasts damaged in this way? Is it a wounding response as animals are exposed to an air interface during shaking? It is also not clear to what extent this displacement reveals more general principles of the effects of noise on hair cells. Additional discussion of underlying causes would be welcome.

We agree that the underlying causes of the physical displacement of posterior lateral-line neuromasts warranted further investigation and we have expanded appropriate sections of the results. To determine if excessive hair-cell activity plays a role in the displacement of neuromasts we have exposed lhfpl5b mutant—fish that have intact hair cell function in the ear, but no mechanotransduction in hair cells of the lateral line—to mechanical overstimulation. We observed comparable disruption of neuromasts lacking mechanotransduction, supporting that displacement of lateral-line hair cells is due to mechanical damage and does not require intact mechnotransduction. Further, when examining the adjacent supporting cells in disrupted neuromasts, we observed they are similarly displaced and elongated. We conclude that observed disruption of hair cells is a consequence of mechanical displacement of the entire neuromast organ. We have added additional discussion of this phenomenon to the Results and Discussion sections of the manuscript.

3. Because afferent neurons innervate more than one neuromast and more than one hair cell per neuromast, measurements of innervation of neuromasts (Figure 3) or synapses per hair cell (Figure 4) cannot be assumed to be independent events. That is, changes in a single postsynaptic neuron may be reflected across multiple synapses, hair cells, and even neuromasts. This needs to be accounted for in experimental design for statistical analysis.

We agree that changes in single postsynaptic neurons, which innervate groups of hair cells of the same polarity within a neuromast, could be reflected across multiple synapses. Additionally, it is plausable that excitotoxic events at the postsynapse, while not contributing to apparent neurite retraction, could be contributing to synapse loss across multiple innervated hair cells. We have updated the manuscript to reflect the potential contribution of postsynaptic signaling to synapse loss and added experiments pharmacologically blocking glutamate uptake.

4. The SEM analysis provides compelling snapshots of apical damage but could be supplemented by quantitative analysis with antibody staining or transgenic lines where kinocilia are labeled. The amount of reduced FM1-43 labeling is one of the more dramatic effects of the shaking insult, suggesting widespread disruption to mechanotransduction that could be related to this apical damage. Further examination of the recovery of mechanotransduction would be interesting.

To supplement the SEM snapshots of severe apical damage, we have expanded the SEM image analysis with quantitative data on kinocilia morphology. We have also added confocal images of hair bundles using antibody labeling of acetylated tubulin in a transgenic line expressing β-actin-GFP in hair cells. We agree that correlative studies of mechanotransduction recovery relative to hair-bundle morphology would be interesting, and we intend to examine this question in a future follow-up study.

5. A previous publication by Uribe et al.2018 describes a somewhat similar shaking protocol with somewhat different results – more long-lasting changes in hair cell number, presynaptic changes in synapses, etc. It would be worth discussing potential differences across the two studies.

We agree we did not adequately address the considerable differences between our mechanical damage protocol for the zebrafish lateral line and the damage protocol described by Uribe et al., 2018. We have provided a more direct comparison in the Results section and addressed the differences in our protocols in-depth in the Discussion section.

Our damage protocol uses a stimulus within the known frequency range of lateral-line hair cells (60 Hz) that is applied to free-swimming larvae and evokes a behaviorally relevant response (fast start response). The damage is observable immediately following noise exposure, is specific to posterior lateral-line neuromasts, and appears to be rapidly repaired. Some features of the damage we observe—reduced mechanotransduction and hair-cell synapse loss—may correspond to mechanically induced damage of hair cell organs in other species. Notably, hair cell synapse loss in seemingly intact neuromasts is exacerbated by pharmacologically blocking synaptic glutamate clearance, supporting that the 60 Hz frequency stimulus is overstimulating neuromast hair cells directly and suggesting that the mechanism of synapse loss may be similar to inner hair cell synapse loss reported in mice following moderate noise exposures.

By contrast, the damage protocol published by Uribe et al. used ultrasonic transducers (40-kHz) to generate small, localized shock waves rather than directly stimulate neuromast hair cells. The damaged they reported—delayed hair-cell death and modest synapse loss with no effect on hair-cell mechanotransduction—was not apparent until 48 hours following exposure and not specific to the lateral-line organ. Some of the features of the damage they observed—delayed onset apoptosis and hair-cell death—may correspond to damage reported in mice following blast injuries.

Reviewer #3:

Holmgren et al. describe a novel model of reversible mechanical damage to zebrafish neuromast hair cells. The authors demonstrate that when zebrafish are exposed to strong currents, neuromast morphology, hair cell number, innervation, and MET function suffer various types and degrees of damage, from which the NMs recover within 2 days. Additionally, they show macrophage recruitment to damaged neuromasts, where they may be phagocytosing synaptic debris. Based on various mechanistic and phenotypic commonalities (involvement of ROS, stereocilia and synapse phenotype), the authors argue that this model is a good approximation of noise-induced hair cell damage in mammals.

Overall impact:

This reviewer agrees that a "noise" damage model in the zebrafish would be a powerful tool to better understand the mechanisms underlying noise-induced hearing loss. However, due to various weaknesses of the data (detailed below), the main claims of the paper are not sufficiently supported. In addition, noise-induced hearing loss has been previously modeled in the zebrafish model. The present model, therefore, does not provide a significant methodological innovation. Based on this, and the fact that addressing all the concerns listed below likely exceeds the scope of a reasonable revision, this manuscript is believed to lack the impact and novelty to be recommended for publication in eLife.

– As the authors point out, zebrafish hair cells can be regenerated. With that in mind, and to make the relevance for mammalian hair cell repair clear, a clear distinction between mechanisms mediated by "repair" or "regeneration" needs to be made. The authors discuss that proliferative hair cell generation can be excluded based on the short time period, but suggest that transdifferentiation might be involved. Recovery of NM hair cell number occurs within the same 2 hour period in which NM morphology and hair cell function improved, making it difficult to determine the extent to which "regeneration" contributed to the recovery. The amount of transdifferentiation has to be shown experimentally (lineage tracing?).

We agree that the distinction between "repair" and "regeneration" needs to be made when discussing this model of mechanical damage to zebrafish hair cell organs. We have tried to clarify that most of what we observe regarding recovery—restoration of neuromast shape, mechanostransduction, afferent contacts, and synapse number —reflect mechanisms of repair following mechanical damage (and, in the case of synapse loss, overstimulation) rather than regeneration. However, one feature of damage that may reflect rapid regeneration is restoration of hair cells number following mechanical injury. To experimentally determine whether proliferation contributed to hair cell generation, we assessed the incorporation of the thymidine analog EdU during a 4 hour recovery following mechanical overexposure in a transgenic line expressing GFP in neuromast supporting cells and observe a modest but not statistically significant increase in the number of proliferating supporting cells in neuromasts exposed to strong current stimulus, suggesting recovery of lost hair cells is not primarily due to renewed proliferation.

The number of hair cells that are lost and recover within several hours are low, i.e., typically ~1 hair cell/neuromast. We observed this consistently in all of our experiments, but the mechanisms responsible are not clear. Based on previous studies of hair cell regeneration in the lateral line, the recovery time appears too rapid to be caused by renewed proliferation, a notion that is further supported by our Edu studies. On the other hand, it is possible that a few supporting cells may undergo the initial phases of phenotypic change into hair cells during this short time period, and we speculate that such transdifferentiation may be responsible for the observed recovery. We should emphasize that this is a new observation and, at present, we do not fully understand the underlying mechanism. However, the focus of the present study is on mechanical damage, synaptic loss, and subsequent repair. We believe that it is important to report our consistent findings of low level hair cell loss and recovery, but a detailed characterization of the mechanism would require considerable effort and would best be the topic of a future study.

– The classification of "normal" vs "disrupted" is vague and not quantitative. The examples shown in the paper seem to be quite clear-cut, but this reviewer doubts that was the case throughout all analyzed samples. Formulate clear benchmarks and criteria for the disrupted phenotype (even when blind analysis is performed).

We have defined measurable criteria for "normal" vs "disrupted" neuromasts that we have added to the Method Details section: “We defined exposed neuromast morphology as “normal” when hair cells appeared radially organized with a relatively uniform shape and size, with ≤7 µm difference observed when comparing the lengths from apex to base of an opposing pair of anterior/posterior hair cells. Length was measured from a fixed point at the center of the hair bundle to the basolateral end of each opposing hair cell. We defined neuromasts as “disrupted” when hair cells appeared elongated and displaced to one side, with >7 µm difference observed when comparing the lengths of an opposing pair of anterior/posterior hair cells. Generally, the apical ends of the hair cells were displaced posteriorly, with the basolateral ends oriented anteriorly.”

– Sustained and periodic exposure: These two exposure protocols not only differ with respect to sustained vs periodic, they also differ in total exposure time (Figure 2B). This complicates the interpretation, especially considering the authors own finding that a pre-exposure is protective.

To clarify—pre-exposure was not protective to hair-cell survival. Rather, in preliminary experiments, pre-exposure appeared to reduce larval mortality, and we have clarified that observation in the text of the Results and the Methods Details sections. We agree with the reviewer that comparing the two protocols based on differences in time distribution is complicated in that they also differ in total exposure time. For the purpose of clarity, we now focus on the sustained exposure in the main figures and created supplemental figures for the reduced damage still observed using periodic exposure, specifying that reduced damage may be the result of periodic time distribution of stimulus and/or less cumulative time exposed to the stimulus.

– The data on the mitochondrial ROS aspect seems not well integrated into the overall story.

We agree that the ROS story was not well integrated and incomplete. We have removed the data describing mpv17-/- mutants and mitochondrial disfunction from this manuscript. A more comprehensive report of mpv17-/- mutant mitochondrial function and morphological analysis of neuromasts following noise exposure will be described in a follow-up manuscript.

– It is surprising that the hair bundle morphology was not assessed after recovery. This is crucial. Overall, it would be good to see some quantification of the SEM data, e.g. kinocilia length and number of splayed bundles.

We have expanded the SEM image analysis to quantitatively access kinocilia morphology following exposure. We agree that assessment of recovery using live-imaging of hair bundles paired with subsequent SEM analysis will be informative, and we intend to perform those experiments in a future study.

– Behavioral recovery (measured as number of "fast start" responses) was also not assessed. This is essential for determining the functional relevance of the recovery.

We attempted to measure behavior recovery of lateral-line function by measuring “fast-start” responses immediately and several hours after recovery, and discovered that i) strong water current provided stimulation that was too intense to reveal subtle behavioral changes following lateral-line damage and recovery, and ii) when testing larvae immediately following sustained strong current exposures, it was difficult to discern if fewer “fast-start” responses were due to lateral-line organ damage or larval fatigue.

We agree that behavioral recovery is important to assay but acknowledge assessing lateral-line mediated behavior following mechanical damage will require a more sensitive testing paradigm that stimulates the lateral-line sensory organ with a relatively gentile, calibrated water flow stimulus. We are currently performing a follow-up study to this paper using a testing paradigm developed by a postdoctoral associate in our lab (Kyle Newton) that analyses subtle changes in larval orientation to water flow (rheotaxis) mediated by the lateral-line organ. Using this behavior paradigm, we will directly correlate morphological and functional recovery over time.

– This reviewer is not yet convinced that this damage model displays enough commonalities to mammalian noise damage to justify the ubiquitous use of the term "noise" throughout the manuscript. It would be more prudent to use a more careful term along the lines of "mechanical overstimulation-induced damage".

We have removed the term “noise” throughout the manuscript and replaced it with either “strong water current stimulus” or “mechanical overstimulation” where appropriate.

– Overall, there was a lack of experimental and analysis detail in the Results section. For example, how was afferent innervation quantified? Just counting GFP labeled contacts to hair cells?

Innervation of neuromast hair cells was quantified during blinded analysis by scrolling through confocal z-stacks of each neuromast (step size 0.3 µm) containing hair cell and afferent labeling and identifying hair cells that were not directly contacted by an afferent neuron i.e. no discernable space between the hair cell and the neurite. Hair cells that were identified as no longer innervated showed measurable neurite retraction; there was generally >0.5 µm distance between a retracted neurite and hair cell. We have added this information to the Methods Detail section.

There was also inconsistency in the use of two variations of the mechanical damage protocol, the time points at which repair was assessed, and whether the damage was quantified in all neuromasts or in normal vs. disrupted neuromasts separately, making the data difficult to interpret.

We have revised our figure legends to clearly indicate when we are assessing damage in all exposed neuromasts (pooled) to control vs. comparative analysis of normal vs. disrupted neuromasts relative to control. In addition, we now focus on the sustained exposure in the main figures, which was the exposure protocol used for the time points in which repair and recovery were assessed.

[Editors’ note: what follows is the authors’ response to the second round of review.]

All three reviewers appreciated the fact that you have made a good effort in revising the manuscript. However, the story has also changed from a noise-induced damage to a mechanical over-stimulation paradigm. While the Ihfp15b mutant results are interesting, it is not entirely clear how these results are applicable to noise-induced hearing loss in mammals. A point that could be better addressed in the Discussion. After an extensive discussion with the three reviewers, they have consolidated their main suggestions as follow:

Essential revisions

1) It would be important to determine whether the hair cells with severe bundle damage recover their bundle morphology or are replaced by transdifferentiation of SCs.

Our revised manuscript provides data that indicate disrupted neuromasts, which appear to correspond with damaged hair bundles in our SEM imaging experiments, are repaired rather than replaced following mechanical damage.

We performed time series experiments examining mechanotransduction and neuromast morphology 0-4 hours following strong water wave exposure, which are now reported in the revised Figure 10. We observed hair-cell mechanotransduction, as measured with FM1-43X uptake, was significantly reduced immediately following exposure. However, uptake of FM1-43X began to recover within 30 minutes and was fully restored after several hours. Notably, reduced FM1-43X uptake did not appear to correlate with stereocilia length. Recovery of FM1-43X fluorescence instead coincided with a relative decrease in the number of neuromasts that appeared “disrupted”, suggesting restoration of overall neuromast morphology. In addition, we expanded our evaluation of cell proliferation following 4 hours recovery and observed no significant increase in supporting cell EdU labeling in exposed neuromasts (Figure 11). Together, these observations suggest that neuromast hair cells disrupted by strong mechanical stimuli are generally repaired rather than replaced.

2) Statistical analyses of results should be incorporated per comments from Reviewer#2. It is important to know if the conclusions hold up across total neuromasts and fish. In this regard, this is a reference suggested by the Reviewer: Aarts E, Verhage M, Veenvliet JV, Dolan CV, van der Sluis S. A solution to dependency: using multilevel analysis to accommodate nested data. Nat Neurosci. 2014 Apr;17(4):491-6.

To accommodate nested data and ensure statistical rigor, our data has been reanalyzed by Dr. Dorina Kallogjeri, a research statistician affiliated with our department. Using an unconditional model, she determined ~35% of variability in hair cell number ~25% variability in innervation could be accounted for by the fish in the experiment. She therefore used a multilevel model to test statistical differences between conditions and groups. Tukey’s adjustment was used for the α level to avoid type I error inflation due to multiple comparisons.

In addition to updating our statistics and figures in the manuscript, we have included our raw data and her analysis in our resubmission.

Reviewer #1:

The authors embark on a comprehensive characterization of damages suffered by the zebrafish lateral line organ in response to mechanical overstimulation. The ultimate goal is to establish this as a novel model for noise-induced hearing loss. The study describes significant overlap in the damage characteristics between mammals and fish, justifying this protocol as a (limited) but very useful model (due to the well-known advantages of the zebrafish system) to study the basic mechanisms of hair cell damage incurred by mechanical overstimulation.

The revision includes some valuable additions in response to the previous review, but many suggestions were deemed intractable in the given time period. The manuscript is therefore improved, but not to a level that this reviewer judges had hoped for.

– The quantification of changes in SEMs of hair bundle morphology following damage was a significant improvement from the original submission, but imaging or quantification of recovery, which would be an important addition to the manuscript, is still lacking. It would also be nice to see a quantification of splayed bundles and kinocilia length before and after recovery. The confocal images demonstrating the changes to the kinocilium following damage were a nice addition and might provide an easier way to quantify changes during the recovery process.

Quantification of phalloidin-labeled hair bundle length during recovery and in relation to FM1-43 uptake is now reported in revised Figure 10.

– The authors decided that further investigation into the distinction of repair vs recovery (beyond SC EdU incorporation) is outside the scope of this study, but this reviewer thinks that it should have been further explored. The authors claim that they only see a modest increase in SC EdU incorporation (>0.5 SC/neuromast) following injury, but the decrease in HC number is also small (looks to be <1 HC/neuromast from graph). Directly addressing transdifferentiation of SCs through lineage tracing techniques would be informative.

It is notable that prior studies of hair cell regeneration in the zebrafish lateral line suggest that the underlying cellular mechanisms are slightly different from those that occur in the inner ears of other nonmammalian vertebrates. The ears of birds and amphibians can generate replacement hair cells either by asymmetric division of supporting cells or by nonmitotic transdifferentiation of supporting cells (Robertson et al., 2004). In contrast, regeneration of zebrafish lateral line hair cells appears to occur via the symmetric division of supporting cells, and no prior studies, to our knowledge, have shown evidence for either transdifferentiation or production by asymmetric proliferation.

To address the issue of hair cell addition, we examined supporting cell proliferation (via EdU labeling) along with cell fate using a transgenic line that expressed a stable fluorophore (GFP) in supporting cells. In these fish, transdifferentiation of supporting cells should result in the colocalization of GFP with the hair-cell marker otoferlin, either immediately after or at short time intervals after mechanical injury. It should be noted that this method will label both hair cells that arise from transdifferentiation as well as newly-differentiated hair cells that were created by the proliferation of (GFP-expressing) supporting cells. In mechanically-damaged fish, we observed such GFP colocalization within 1-2 hair cells in a small fraction of neuromasts (3/109 neuromasts; n=41 fish, examined at 0-2 hr after exposure). However, we also observed similar colocalization in unexposed (control) fish (1/49 neuromasts, n=24 fish), suggesting that addition of new hair cells (by whatever mechanism) is similar in both damaged and control fish. In addition, supporting cell proliferation (as quantified by EdU labeling; Figure 11) did not significantly differ in mechanically injured vs. control fish. Together, these observations suggest that the mechanical trauma did not affect the rate at which new hair cells are generated. Finally, since hair cells in zebrafish neuromasts undergo a slow rate of turnover (e.g., Williams and Holder, 2000) it is expected that a low level of hair cell addition will continuously occur in control and mechanically overstimulated larvae.

Reviewer #2:

I appreciate the extensive revisions Holmgren et al. have made to their previous manuscript, addressing many of the criticisms of the previous review. They include a clever new experiment using lhfpl5b mutants that have mechanosensory deficits specific to lateral line hair cells. Using these mutants, they demonstrate that the morphological disruption they observe after shaker treatment occurs in these mutants to the same degree as in wt fish.

However, this result does call into question whether the model is a functional one for noise damage. If mechanotransduction is not necessary, are the phenotypes observed really excitotoxicity? Do the reported changes in innervation and synapses also not require functional mechanotransduction?

We believe the data showing significant synapse loss in neuromasts with overall intact morphology (i.e. “normal”) and our observation that blocking glutamate uptake significantly worsens synapse loss specifically in “normal” neuromasts provide strong evidence that synapse loss is a consequence of excitotoxicity.

I fear I was not clear enough on my previous point (#3 in response to reviewer 2):

Because afferent neurons innervate more than one neuromast and more than one hair cell per neuromast, measurements of innervation of neuromasts (Figure 3) or synapses per hair cell (Figure 4) cannot be assumed to be independent events. That is, changes in a single postsynaptic neuron may be reflected across multiple synapses, hair cells, and even neuromasts. This needs to be accounted for in experimental design for statistical analysis.

As measurements of innervation or synapse density are not independent, they cannot be treated as distinct instances (n's) in the statistical analysis. In these experiments the independent instances are going to be the number of fish, not hair cells. This issue extends to analysis in Figures7, 8 as well.

To accommodate nested data and ensure statistical rigor, we used a multilevel model to test statistical differences between groups.

I am somewhat confused about the conclusions from figure 4. Setting aside the issue of independence, the differences in synapses per hair cell are variably measured as significant or insignificant, suggesting that the analysis is underpowered. In addition, shouldn't the relevant comparisons in Figure 4E be between DMSA and TBOE, that is comparing each of control, shaken normal and shaken disrupted to the corresponding drug treatment?

We have included the appropriate comparisons in the updated Figure 4.

Reviewer #3:

The goal of the manuscript is to present a new paradigm for damaging hair cells of the zebrafish lateral line using vibrational stimuli. A successful method would be important to the community as this goal has been difficult to achieve, and it would open up new avenues of research. The authors addressed some of my concerns from the last review but not all. The experiments they did perform are a screen of frequencies that damage the hair cells of the lateral line and improved the quality of an image (Figure 4). This is appreciated especially during the time of Covid.

They argue that 2 suggested experiments are not necessary: microphonic potentials and damage to the core of actin of stereocilia. I feel that they would improve the paper's quality, but I see their point about not including the microphonic potentials; however, they should at some level look at the damage to the actin core. This could be by TEM as suggested or it could be by phalloidin labeling. Both of these methods can be used to see gaps in the core of actin in mouse hair cells and maybe those of zebrafish.

While we were unable to discern damage to the actin core in zebrafish neuromast hair bundles using phalloidin labeling, we were able to characterize qualitative changes as well as quantify average stereocilia length in projected 3D image stacks. We used these data to determine how stereocilia morphology corresponds with FM1-43 uptake recovery in the revised Figure 10.

Overall, the present study’s aims were to evaluate mechanical damage in the zebrafish lateral line and to establish this injury paradigm as a model for the study of synaptic repair. A more detailed analysis of actin injury and repair would require considerable TEM imaging which, while we hope may be performed in future studies, is beyond the scope of the present study.

Another issue is the use of the word "current". According to the dictionary (OED) current is defined as "That which runs or flows, a stream; spec. a portion of a body of water, or of air, etc. moving in a definite direction." This is the way I see the word used in the scientific literature as well. I don't think what the authors are providing is a current. They should use a different term, perhaps, "vibratory stimulus" or "water stimulus."

We have replaced the term “current” with “water wave” in the manuscript.

https://doi.org/10.7554/eLife.69264.sa2

Article and author information

Author details

  1. Melanie Holmgren

    Department of Otolaryngology, Washington University School of Medicine, St Louis, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2541-4854
  2. Michael E Ravicz

    1. Eaton-Peabody Laboratory, Massachusetts Eye and Ear, Boston, United States
    2. Department of Otolaryngology–Head and Neck Surgery, Harvard Medical School, Boston, United States
    Contribution
    Methodology, Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9978-3444
  3. Kenneth E Hancock

    1. Eaton-Peabody Laboratory, Massachusetts Eye and Ear, Boston, United States
    2. Department of Otolaryngology–Head and Neck Surgery, Harvard Medical School, Boston, United States
    Contribution
    Software
    Competing interests
    No competing interests declared
  4. Olga Strelkova

    1. Eaton-Peabody Laboratory, Massachusetts Eye and Ear, Boston, United States
    2. Department of Otolaryngology–Head and Neck Surgery, Harvard Medical School, Boston, United States
    Contribution
    Data curation, Formal analysis
    Competing interests
    No competing interests declared
  5. Dorina Kallogjeri

    Department of Otolaryngology, Washington University School of Medicine, St Louis, United States
    Contribution
    Formal analysis
    Competing interests
    No competing interests declared
  6. Artur A Indzhykulian

    1. Eaton-Peabody Laboratory, Massachusetts Eye and Ear, Boston, United States
    2. Department of Otolaryngology–Head and Neck Surgery, Harvard Medical School, Boston, United States
    Contribution
    Data curation, Formal analysis, Funding acquisition, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2076-6818
  7. Mark E Warchol

    1. Department of Otolaryngology, Washington University School of Medicine, St Louis, United States
    2. Department of Neuroscience, Washington University School of Medicine, St Louis, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0445-6318
  8. Lavinia Sheets

    1. Department of Otolaryngology, Washington University School of Medicine, St Louis, United States
    2. Department of Developmental Biology, Washington University School of Medicine, St. Louis, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Supervision, Writing - original draft
    For correspondence
    sheetsl@wustl.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5231-2450

Funding

National Institute on Deafness and Other Communication Disorders (R01DC016066)

  • Lavinia Sheets

National Institute on Deafness and Other Communication Disorders (R01DC017166)

  • Artur A Indzhykulian

National Institute on Deafness and Other Communication Disorders (R01DC006283)

  • Mark E Warchol

Washington University School of Medicine in St. Louis

  • Lavinia Sheets

Amelia Peabody Charitable Fund

  • Lavinia Sheets

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

Acknowledgements

This work was supported by the National Institute on Deafness and Other Communication Disorders R01DC016066 (LS), R01DC017166 (AAI), and R01DC006283 (MEW), Washington University Dept. of Otolaryngology (LS), and the Amelia Peabody Charitable Fund (LS). We thank Valentin Militchin (WashU) and Evan Foss (Mass Eye and Ear) for engineering support and Mark Rutherford for thoughtful feedback on the manuscript.

Ethics

This study was performed with the approval of the Institutional Animal Care and Use Committee of Washington University School of Medicine in St. Louis (protocol no. 20–0158) and in accordance with NIH guidelines for use of zebrafish.

Senior Editor

  1. Didier YR Stainier, Max Planck Institute for Heart and Lung Research, Germany

Reviewing Editor

  1. Doris K Wu, NIDCD, NIH, United States

Publication history

  1. Preprint posted: July 17, 2020 (view preprint)
  2. Received: April 9, 2021
  3. Accepted: October 18, 2021
  4. Accepted Manuscript published: October 19, 2021 (version 1)
  5. Version of Record published: October 29, 2021 (version 2)

Copyright

© 2021, Holmgren et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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    Insulin-induced hypoglycemia is a major treatment barrier in type-1 diabetes (T1D). Accordingly, it is important that we understand the mechanisms regulating the circulating levels of glucagon. Varying glucose over the range of concentrations that occur physiologically between the fed and fuel-deprived states (8 to 4 mM) has no significant effect on glucagon secretion in the perfused mouse pancreas or in isolated mouse islets (in vitro), and yet associates with dramatic increases in plasma glucagon. The identity of the systemic factor(s) that elevates circulating glucagon remains unknown. Here, we show that arginine-vasopressin (AVP), secreted from the posterior pituitary, stimulates glucagon secretion. Alpha-cells express high levels of the vasopressin 1b receptor (V1bR) gene (Avpr1b). Activation of AVP neurons in vivo increased circulating copeptin (the C-terminal segment of the AVP precursor peptide) and increased blood glucose; effects blocked by pharmacological antagonism of either the glucagon receptor or V1bR. AVP also mediates the stimulatory effects of hypoglycemia produced by exogenous insulin and 2-deoxy-D-glucose on glucagon secretion. We show that the A1/C1 neurons of the medulla oblongata drive AVP neuron activation in response to insulin-induced hypoglycemia. AVP injection increased cytoplasmic Ca2+ in alpha-cells (implanted into the anterior chamber of the eye) and glucagon release. Hypoglycemia also increases circulating levels of AVP/copeptin in humans and this hormone stimulates glucagon secretion from human islets. In patients with T1D, hypoglycemia failed to increase both copeptin and glucagon. These findings suggest that AVP is a physiological systemic regulator of glucagon secretion and that this mechanism becomes impaired in T1D.