1. Cell Biology
  2. Developmental Biology
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Lamellar projections in the endolymphatic sac act as a relief valve to regulate inner ear pressure

  1. Ian A Swinburne
  2. Kishore R Mosaliganti
  3. Srigokul Upadhyayula
  4. Tsung-Li Liu
  5. David G C Hildebrand
  6. Tony Y -C Tsai
  7. Anzhi Chen
  8. Ebaa Al-Obeidi
  9. Anna K Fass
  10. Samir Malhotra
  11. Florian Engert
  12. Jeff W Lichtman
  13. Tomas Kirchhausen
  14. Eric Betzig
  15. Sean G Megason  Is a corresponding author
  1. Harvard Medical School, United States
  2. Boston Children’s Hospital, United States
  3. Janelia Research Campus, Howard Hughes Medical Institute, United States
  4. Harvard University, United States
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Cite this article as: eLife 2018;7:e37131 doi: 10.7554/eLife.37131

Abstract

The inner ear is a fluid-filled closed-epithelial structure whose function requires maintenance of an internal hydrostatic pressure and fluid composition. The endolymphatic sac (ES) is a dead-end epithelial tube connected to the inner ear whose function is unclear. ES defects can cause distended ear tissue, a pathology often seen in hearing and balance disorders. Using live imaging of zebrafish larvae, we reveal that the ES undergoes cycles of slow pressure-driven inflation followed by rapid deflation. Absence of these cycles in lmx1bb mutants leads to distended ear tissue. Using serial-section electron microscopy and adaptive optics lattice light-sheet microscopy, we find a pressure relief valve in the ES comprised of partially separated apical junctions and dynamic overlapping basal lamellae that separate under pressure to release fluid. We propose that this lmx1-dependent pressure relief valve is required to maintain fluid homeostasis in the inner ear and other fluid-filled cavities.

https://doi.org/10.7554/eLife.37131.001

eLife digest

The most internal part of the human ear, the inner ear, is essential for us to hear and have a sense of balance. It is formed by a complex series of connected cavities filled by a liquid. When sound waves and changes in the position of the body make this liquid move, specialized ‘hair’ cells can detect these subtle movements; neurons then relay this information to the brain where it is decoded and interpreted.

For the inner ear to work properly, the body needs to finely regulate the pressure created by the liquid inside the cavities. For example, people with unstable pressure in their ears can experience deafness or problems with balance. A structure known as the endolymphatic sac, which is a balloon-like chamber connected to the rest of the inner ear by a thin tube, helps with this regulation. However, scientists are still unsure about how exactly the sac performs its role. One problem is that the inner ear is difficult to study because it is encased in one of the densest bones in the body.

Many other animals also have inner ears, from fish to birds and mammals. Here, Swinburne et al. examine the inner ear of zebrafish embryos because, in this fish, the ear starts working before the bones around it form; the structure is therefore accessible for injections and microscopy. Experiments show that when the pressure in the inner ear rises, the endolymphatic sac slowly fills up with the ear liquid, and then it rapidly deflates. Fish with mutations that stop the sac from deflating have overinflated sacs, which is a symptom also found in certain patients with hearing and balance disorders.

Looking into the details of these inflation-deflation cycles, Swinburne et al. found that the cells that form the sac have gaps between them, unlike a normal sheet of cells. A flap covers these gaps to keep the liquid in, but under pressure, the flap opens and the liquid can escape. These results show that the endolymphatic sac works as a pressure relief valve for the inner ear.

Ultimately, understanding how pressure is regulated in the ear could help patients with inner ear disorders. It could also serve as a template to investigate how eyes, kidneys and the brain, which all have liquid-filled cavities, control their internal pressure.

https://doi.org/10.7554/eLife.37131.002

Introduction

Understanding the mechanisms by which organs use water-filled cavities to compartmentalize biochemical and biophysical environments is a fundamental problem. Because water is nearly incompressible, several organs harness and cope with water as an object that transmits force. Hydrostatic pressure inflates the eye during development (Coulombre, 1956). Later, unstable ocular pressure from reduced production of aqueous humor or reduced drainage can lead to blindness, as occurs in hypertonic maculopathy or glaucoma (Costa and Arcieri, 2007; Leske, 1983). Hydrostatic pressure appears to drive expansion of brain ventricles during development (Desmond, 1985; Desmond and Levitan, 2002; Lowery and Sive, 2005). Later, unstable hydrostatic pressure in brain ventricles is correlated with hydrocephaly and mental disorders (Hardan et al., 2001; Kurokawa et al., 2000). Hydrostatic pressure inflates and controls the size of the developing ear (Abbas and Whitfield, 2009; Hoijman et al., 2015; Mosaliganti et al., unpublished). Later, unstable pressure in the ear can cause deafness and balance disorders like Pendred syndrome and Ménière’s disease (Belal and Antunez, 1980; Schuknecht and Gulya, 1983). This theme of harnessing hydrostatic pressure for normal development and controlling pressure for healthy physiology raises the question of how tissues regulate pressure. Tissue structures identified as important for pressure control include Schlemm’s canal in the eye, arachnoid granules and the choroid plexus in the brain, and the endolymphatic duct and sac in the ear (Johnstone et al., 2011; Kimura and Schuknecht, 1965; Naito, 1950; Orešković et al., 2017; Pollay, 2010; Symss and Oi, 2013; Tripathi, 1974). A cavity’s pressure could be managed via mechanisms involving molecular pores, molecular transporters, and the physical behavior of the tissue. Observing the mechanisms by which these tissue barriers control pressure has been limited by a range of obstacles such as optical accessibility and uncertainty in both the time- and length-scale on which they function. As such, not much is known about how these tissues manage fluctuating pressures because they have not been observed in vivo.

The inner ear is a prominent example of an organ whose tissue form determines its physiology. It is filled with endolymph, the composition of which differs from other fluids such as plasma, perilymph, and cerebral spinal fluid in that it contains high potassium, low sodium, and high electric potential (Lang et al., 2007). This endolymph composition is necessary to drive ion currents into hair cells to convert fluid movement, caused by either the head’s acceleration or by sound, into biochemical signals that underlie balance and hearing. The ear’s endolymphatic duct connects the endolymph in the semicircular canals, cochlea, and other chambers to the endolymphatic sac (ES, Figure 1A,B). An ES-like structure is present in basal vertebrates, including lamprey and hagfish (Hammond and Whitfield, 2006), suggesting it plays an ancient role in inner ear function. The epithelium of the ES, as well as that of the rest of the ear, has an apical surface facing the internal endolymph and a basal surface facing the external perilymph (Figure 1A). Most of the adult ear is enclosed in a bony labyrinth within the temporal bone and perilymph is located in the space between the epithelium and bone (Figure 1A). In adults, the endolymphatic duct and sac are partially encased within the temporal bone such that the distal end of the endolymphatic sac protrudes out of the temporal bone into the cranial cavity (Figure 1A). Excessive hydrostatic pressure can tear the inner ear’s epithelium, disrupting its electric potential (a pathology called endolymphatic hydrops). In contrast, low potassium or reduced endolymph production can lower hydrostatic pressure within the ear to the point where the ear chambers may collapse (Lang et al., 2007). Early work showed that ES ablation causes hydrostatic pressure to rise and the epithelium to tear (Kimura and Schuknecht, 1965; Naito, 1950), suggesting that the ES may maintain endolymph homeostasis. Recent work indicated that an explanted mouse ES gradually loses endolymph (Honda et al., 2017). While many molecular pores and transporters are expressed in the ES, it is unclear why it is organized as a dead-end tube and how it might reduce endolymph volume.

Figure 1 with 1 supplement see all
ES lumen slowly inflates and rapidly deflates every 0.3–4.5 hr.

(A) Illustration of the adult human inner ear showing cochlea, semicircular canals (SCCs), and endolymphatic duct and sac (ES, red arrowhead) and their organization of tissue (green), endolymph (beige), perilymph (magenta), and bone (black). Illustrations of the adult and larval zebrafish inner ear showing ES (indicated with red arrowheads, see also Figure 1—figure supplement 1 and Video 1 for how the zebrafish ES first forms). (B) Micrograph of larval zebrafish, sagittal view. (B’) In situ of foxi1 highlights position of ES (red arrowhead), n = 12. (C) Illustration of imaging setup. (D) Slices and select time points from 3D confocal time course showing a single inflation and deflation event from a live zebrafish embryo. Cell membranes (green) are labeled using ubiquitous membrane citrine transgenes. Perilymph (magenta) is labeled with 3 kDa dextran-Texas red. ES identified with dotted blue outline and yellow arrow. Lumen of inflated ES identified with dashed white outline in 64:36 panel. (E) Corresponding 3D meshes of the segmented ES lumen volume. (F) Quantification of segmented ES volumes (primary axis, green) and leak in fluorescence (secondary axis, magenta) over multiple cycles (see also Figure 1—figure supplement 1B,C and Videos 23). (G) Histogram of times between peak inflation volumes, compiled from eight different time courses and 54 inflations. Scale bars 100 μm for (B) and 10 μm for (D,E).

https://doi.org/10.7554/eLife.37131.003

The ES has not been studied much in zebrafish because it has not always been clear that zebrafish possess this structure or at what time it forms during development (Haddon and Lewis, 1996; Hammond and Whitfield, 2006). In situ hybridization images at 52 hr post fertilization (hpf) showing expression of ES markers, foxi1 and bmp4, restricted to an ES-like structure positioned on the dorsal side of the otic vesicle are the strongest and developmentally earliest evidence for zebrafish having an ES (Figure 1B’) (Abbas and Whitfield, 2009; Geng et al., 2013). However, its formation and physiological function remain unknown. Here, we present a detailed characterization of the ES in zebrafish performed using confocal microscopy, serial-section electron microscopy, adaptive optics lattice light-sheet microscopy, and a genetic mutant that together demonstrate that the ES contains a physical relief valve for regulating inner ear fluid pressure through regulated transepithelial fluid release.

Results

The endolymphatic sac exhibits cycles of inflation and deflation

We first established an imaging system to identify the developmental origins of the zebrafish ES. Extended time-lapse imaging required long-term immobilization with α-bungarotoxin that permits normal development without the reduction in ear growth caused by long-term treatment with tricaine, bright fluorescent transgenic fish with contrast from a membrane-localized fluorescent protein to minimize bleaching and photoxicity, and mounting in a submerged agarose canyon that permits positioning of the zebrafish ear close to the coverslip (400 μm wide, walls of canyon secure the yolk and head, Figure 1C) (Swinburne et al., 2015). The zebrafish ear develops from a collection of epithelial cells that form a closed fluid-filled ellipsoidal structure called the otic vesicle (Whitfield, 2015). We saw that ES morphogenesis begins at 36 hpf as an evagination in the dorsal epithelial wall of the otic vesicle (Figure 1—figure supplement 1A, Video 1, Figure 1A–B, and ES identity confirmed by expression of foxi1, Figure 1B’). Between 36 and 60 hpf, the ES grows and elongates. These findings established that the zebrafish ES is optically accessible during embryonic and larval stages and that ES morphogenesis begins at 36 hpf.

Video 1
Early ES development.

Video begins with schematic of experimental set-up and context of the presented field of view. Then, an annotated time point is presented of the upcoming video. The presented video is of a sagittal slice from a 4D time course of early ES development (white arrow points to ES in introduction). ES morphogenesis begins at 36 hr post fertilization (hpf) as an evagination in the dorsal-anterior-lateral epithelial wall of the otic vesicle. Fluorescence from membrane citrine, shown in grey. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.005

The zebrafish ear starts to sense body acceleration for the nervous system between 60 and 72 hpf, as demonstrated by the onset of the vestibulo-ocular reflex (Mo et al., 2010). We found that the ES begins to exhibit a physical behavior during the same time window. We observed that the lumen of the ES remains closed until 60 hpf, but between 60 and 65 hpf it begins cycles of slowly inflating and rapidly deflating (sagittal slices, Figure 1D, lumen volumes, Figure 1E, Figure 1—figure supplement 1, and Videos 23). Three-dimensional (3D) measurements showed that the ES lumen volume changes 5–20-fold through the course of each cycle (Figure 1E,F, and Figure 1—figure supplement 1B–C). The period between peak ES volumes exhibited a broad distribution (0.3–4.5 hr) with an average of 1.6 ± 0.8 hr (mean ± SD, histogram compiled from eight time courses and 54 peaks, Figure 1G). These initial observations suggested several potential causes of the inflation-deflation cycles of the ES including: a response to organ-wide increases in fluid pressure within the otic vesicle, a local tissue behavior wherein the ES inflates with fluid from the perilymph, or a local tissue behavior wherein cells in the ES periodically coordinate their movements to expand the ES volume by sucking fluid from the otic vesicle.

Video 2
Wild-type ES inflates and deflates.

Video begins with an illustration depicting the context of the presented field of view, which is a sagittal slice encompassing the developing ES from a 4D time course. Then, an annotated time point is presented of the upcoming video, a green arrow points to the ES, a dotted line outlines the ES lumen, the otic vesicle is labeled ventral to the ES, and the perilymph surrounds the ES structure, labeled in magenta. Video of sagittal slice from 4D dataset, quantified in Figure 1F. Fluorescence from membrane citrine, shown in green. Perilymph highlighted with fluorescence from 3 kDa dextran-Texas red, shown in magenta. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.006
Video 3
Wild-type ES inflates and deflates.

A video of two time courses, sagittal slices from 4D datasets, quantified in Figure 1—figure supplement 1B (left) and Figure 1—figure supplement 1C (right). Fluorescence from membrane citrine, shown in green. Perilymph highlighted with fluorescence from 3 kDa dextran-Texas red, shown in magenta. Scale bars are 10 μm.

https://doi.org/10.7554/eLife.37131.007

Loss of the epithelial barrier is sufficient for ES deflation

To assess whether ES inflations occur by transmission of endolymph pressure between the otic vesicle and the ES, we performed a single injection of a small volume of solution containing 3 kDa fluorescent dextran into the otic vesicle and followed its movement during inflation and deflation. We found that the duct is open and that endolymph flows from the otic vesicle into the ES during inflation (Figure 2A–D, Video 4). Additionally, upon deflation, endolymph rapidly leaked out of the ES and into the perilymphatic space (75:22 panel, Figure 2D). To determine whether pressure in the otic vesicle is transmitted to the ES for its inflation, we laser-ablated 2–3 cells within the wall of the otic vesicle distant from the ES at 64 hpf (Figure 2E). Shortly after ablation, the ES deflated and completely collapsed within 20 min (Figure 2F). These results indicate that fluid pressure in the otic vesicle inflates the ES volume, the ES tissue has elastic material properties, and a loss of epithelial integrity is sufficient for ES deflation.

Hydrostatic pressure transmits endolymph through duct to inflate the ES.

(A) Illustration of larval zebrafish highlighting sagittal plane of image acquisition (blue square). (B) Time points of individual sagittal slices of raw data from 3D time course (endolymph labeled yellow by single dye injection into otic vesicle, membrane citrine in cyan). (C) Illustration of larval zebrafish highlighting transverse perspective (blue box) for rendered volumes of ES. (D) Time points from time course of raw data rendered in 3D transverse view (endolymph in yellow, ES tissue outlined with dashed line) showing endolymph flowing through duct to ES and then out to perilymph (blue arrow). (E) Illustration of strategy for laser ablating otic vesicle cells with point-scanning 2-photon laser to ablate 2–3 targeted cells. (F) Slices from 4D confocal time course after laser ablation showing ES deflation. ES lumen is outlined with a white dashed line in (F), n = 4. (G) Time points from time course rendered in 3D transverse view (endolymph in yellow, perilymph in magenta, ES tissue outlined with dashed line), n = 8. Blue arrows indicate endolymph expulsion or perilymph leak in events (D,G). All scale bars 10 μm.

https://doi.org/10.7554/eLife.37131.008
Video 4
Endolymph periodically inflates ES and then released into periotic space.

Time course of otic vesicle injected with 3 kDa dextran-Texas red at 55 hpf. Panels are transverse volumes of same time course. Left, labeled endolymph presented in yellow. Right, labeled endolymph in yellow, membrane citrine in cyan. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.009

A core function of epithelial sheets is to act as a barrier that can prevent passive movement of molecules between an organ’s interior and exterior. Deflations could be driven by local breaks in the epithelial barrier combined with elastic tissue collapse or by an alternative mechanism such as cellular reabsorption of endolymph. To distinguish between these mechanisms, we used dye injections into the heart to label the perilymph that surrounds the otic vesicle and ES (Figure 1D). Quantification of fluorescence within the lumen of the ES revealed that perilymph dye begins to leak into the ES lumen at the onset of each deflation (Figure 1F, visible leakage in Figure 1D, Videos 23). In the example shown deflation coincided with an influx of fluorescence from the perilymph into the lumen of the ES in each of the nine major inflation-deflation cycles. This coincident timing suggests that both deflation and dye leakage were due to transient breaks in the epithelial barrier, an interpretation that is consistent with the rapid release of endolymph prior to deflation (Figure 2D, Video 4).

The observation that perilymph enters the deflating ES may seem counterintuitive, since the contents of a punctured high-pressure elastic vessel might be expected to primarily flow outwards. Estimation of the rates of advection and diffusion supports the presence of upstream movement of perilymph by diffusion. Due to limited spatial resolution and the absence of contrast for trafficking vesicles, we could not dismiss alternative mechanisms such as contributions from rapid transcytosis or cellular absorption. To determine whether endolymph efflux occurs at the same time as perilymph influx, we simultaneously labeled the two fluids with different colored dyes and imaged their localization with higher temporal resolution (Figure 2G, Video 5). We found a fluttering behavior underlying deflation in which endolymph leaked out (62:41:20), perilymph leaked in (62:55:20), leaked-in perilymph flushed out with more endolymph efflux (62:55:40), followed by complete deflation (63:12:00). It is difficult to explain the simultaneous inward and outward movement of fluid with the alternative interpretation that rapid transcytosis or cellular absorption processes deflate the ES. A plausible mechanism for this observed behavior involves a structure that normally resists flow driven by a pressure differential that rapidly opens during deflation to allow for fast efflux by advection followed by influx by diffusion before re-closing.

Video 5
Endolymph released into periotic space followed by perilymph leak-in.

Time course of otic vesicle injected with 3 kDa dextran-Texas red at 55 hpf, and perilymph labeled with 10 kDa dextran- Alexa Fluor 488. First three panels are transverse volumes while the fourth is a dorsal view of the same time course. Labeled endolymph presented in yellow, perilymph in magenta. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.010

Lmx1bb is essential for ES deflation

The uniqueness of the ES inflation and deflation cycles suggests specific genes might be involved. To identify pathways that contribute to the emergence of this physiology and to reveal ways in which it can malfunction, we examined a mutant caused by a premature stop codon in the transcription factor lmx1bb (Obholzer et al., 2012) that exhibited an enlarged ES. We found that the ES in lmx1bb mutants became greatly enlarged (>4 times the inflated wild-type ES volume) making it readily visible at 80 hpf by bright-field microscopy (Figure 3A). To determine if lmx1bb is expressed at an appropriate place and time for a mutation to be causing an ES defect, we imaged a transgenic reporter line, Tg(lmx1bb:eGFP)mw10, driven by the lmx1bb promoter McMahon et al., 2009.This reporter was expressed in ES cells beginning at 52–58 hpf, just before the first ES inflation (cyan, Figure 3B), consistent with lmx1bb expression being instrumental for development of the ability of the ES to release pressure. Earlier in the development of the otic vesicle, lmx1bb is expressed in portions of the nascent semicircular canals and sensory patches, regions of the inner ear that also exhibit abnormal development in the mutant (Obholzer et al., 2012; Schibler and Malicki, 2007). There is no precedent for ES development being dependent on those portions of the otic vesicles and there are many mutants with similar SCC or sensory defects that do not have ES phenotypes (Fekete, 1999; Malicki et al., 1996; Whitfield et al., 1996). Live imaging and perilymph tracking in lmx1bbjj410/jj410 mutant embryos revealed that the ES lumen over-inflates (Figure 3C–D, Figure 3—figure supplement 1, and Videos 67). As in the wild-type analysis, we quantified the presence of perilymph leaking into the ES lumen (secondary axes of Figure 3D and Figure 3—figure supplement 1A). In the mutant, however, we never observed perilymph entering the ES. Additionally, we imaged mutants where the endolymph was labeled and did not observe leakage out of the distended ES epithelium (Figure 3E). These findings suggest that the epithelial diffusion barrier remains intact in the mutant ES.

Figure 3 with 1 supplement see all
Lmx1bb is necessary for development of the ES’s ability to form breaks in its diffusion barrier and deflate.

(A) Lateral view of wild-type and lmx1bbjj410/jj410 mutant ears imaged by bright-field microscopy at 80 hpf, asterisk labels greatly enlarged mutant ES. Scale bar, 100 μm. (B) Slices from 3D confocal time course of an lmx1bb transcriptional reporter (cyan, Tg(lmx1bb:egfp)mw10/mw10; yellow, Tg(actb2:mem-mcherry2)hm29), n = 3. (C) Slices and select time points from 3D confocal time course of lmx1bbjj410/jj410 mutant embryos. Membrane (green) from ubiquitous membrane citrine transgenes. Perilymph (magenta) from 3 kDa dextran-Texas red, n = 4. (D) Quantification of segmented ES volumes (primary axis, green) and leak-in fluorescence (secondary axis, magenta) from lmx1bbjj410/jj410 time course in (C) (see also Figure 3—figure supplement 1 and Videos 56). (E) 3D transverse view (endolymph in yellow) from timelapse showing endolymph in dilated mutant ES, outlined with dashed blue line, n = 2. (F) Small regions with thin membranes (asterisks) form in the inflated ES of wild-type but not lmx1bb mutants. (G) Quantification of minimum epithelial thickness versus inflated ES volume in mutant (plotted in red, n = 9) and wild-type (plotted in black, n = 14). Compiled from 65 to 80 hpf embryos. (H) Uneven labeling from Tg(lmx1bb:egfp) reveals thin basal processes (white arrow). (I) Wild-type ES examples with sparsely labeled cells: membrane-labeled citrine (green) in a membrane-labeled cherry background (magenta), white arrows indicate lamellar projections, n = 15. (J) lmx1bbjj410/jj410 mutant ES examples with sparsely labeled cells: membrane cherry (magenta) in a membrane citrine background (green), n = 9. (K) Cartoon schematic of apico-basal organization of ES. (L) Supporting whole-mount immuno-stains for basal and apical markers (collagen and ZO-1, both magenta) in a membrane-labeled citrine background (green), n = 16, 15, 6, 32, 21, and 12 for (i-vi). Scale bars in (B-L), 10 μm.

https://doi.org/10.7554/eLife.37131.011
Video 6
Mutant ES over-inflates.

Video of sagittal slice from 4D dataset of lmx1bbjj410/jj410 mutant- quantified in Figure 3D. Fluorescence from membrane citrine shown in green. Perilymph highlighted with fluorescence from 3 kDa dextran-Texas red, shown in magenta. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.013
Video 7
Mutant ES over-inflates.

Video of sagittal slice from 4D dataset of lmx1bbjj410/jj410 mutant- quantified in Figure 3—figure supplement 1. Fluorescence from membrane citrine shown in green. Perilymph highlighted with fluorescence from 3 kDa dextran-Texas red, shown in magenta. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.014

Lamellar barriers appear to form an ES relief valve

The absence of ES deflation in the lmx1bb mutant suggests that a structural deficiency may be present. Indeed, by comparing the dorsal ES tissue between wild-type and lmx1bb mutants using confocal microscopy and membrane-localized fluorescent proteins, we observed that the ES tissue was much thinner in wild-type embryos during peak inflation than in the corresponding region in mutants inflated to a similar volume (thin region indicated by asterisk, Figure 3F,G, data in G compiled from 65 to 80 hpf inflation events). In wild-type embryos, this region often appeared as a thin sheet (1.0 ± 0.3 µm, mean ±SD) rather than thick as seen in the un-inflated wild-type ES or the lmx1bb mutants in which two distinct surfaces (apical and basal) were observed (6.2 ± 3.2 µm, the mutant, n = 9, is significantly thicker than wild-type, n = 14, with a Mann-Whitney-Wilcoxon one tailed p-value of 4 × 10−5). Imaging the uneven signal of the Tg(lmx1bb:eGFP) reporter revealed thin protrusions that extend along the basal side of neighboring ES cells (white arrow, Figure 3H). Sparse mosaic labeling of cells in the wild-type further confirmed the presence of thin basal protrusions in the ES, as well as diversity in their organization (Figure 3I). Similar labeling in the mutant revealed the absence of thin basal protrusions in the ES (Figure 3J). To determine the gross organization of the ES tissue, we stained for basal proteins collagen (Figure 3K–L) and laminin (not shown). We found that the basal surface of the ES tissue faces the perilymph outside surrounding the otic vesicle (Figure 3L). We could not resolve a clear difference in the staining pattern of collagen between wild-type and the lmx1bb410/410 mutant. Prior work suggested that Col1a2 may be a direct target of LMX1B in the limb (Haro et al., 2017) and additional studies will be necessary to determine whether collagen or other extracellular matrix components are abnormally expressed in the ES of lmx1bb410/410 mutants. We also examined localization of the apical marker ZO-1 and found that it stained the ES tissue that faces the endolymph-filled lumen (Figure 3K–L). This apical-basal organization was also present in the lmx1bb410/410 mutant (Figure 3K–L). The absence of local breaks in the mutant ES, the absence of regional thinning in the mutant inflated ES, and the absence of thin basal protrusions suggests that these thin areas may be structurally relevant to ES pressure relief.

To investigate this possibility, we examined the ultrastructure of these thin areas using serial-section electron microscopy (EM) at a resolution of 4.0 × 4.0 × 60.0 nm per voxel. We found the dorsal tissue of the inflated wild-type sac consists of an extremely thin shell of broadly overlapping lamellae extending from the basal sides of multiple adjacent cells (Figure 4; Video 8). These basal extensions appear to be the distal barrier in the ES because the endolymph they contain is continuous with the lumen of the ES and endolymphatic duct (extending ventrally from outlined ES lumen in third panel of Figure 4A). We term these ‘lamellar barriers’ because of their thin, plate-like structure and apparent function as a barrier that holds the elevated hydrostatic pressure of the endolymph. We observed lamellar sheets that extend for distances as long as a typical cell body but are as thin as 40 nm (Figure 4A,F; lamella from cell segmented in purple extends 7.5 μm in the x-y plane and 6.6 μm along the z-axis). In contrast to junctions between thin aveolar cells, which kiss at their tips with tight junctions, the lamellae from ES cells formed large zones of overlap and sometimes bifurcated to form a tongue-and-groove structure (Figure 4B, inset) or interwoven structures (Figure 4C,F, black mesh highlights area of overlap, dotted perimeter indicates full spread). These structures lacked electron-dense signal, indicating that they were unlikely to contain tight junctions between lamellae. We also identified an ear where the lamellar barriers were separated as if forced into an open configuration (Figure 4D). This may be an ES in the act of deflating via bursting or sliding of the lamellae. The full serial-section EM dataset also confirmed that the endolymphatic duct connects the lumen of the otic vesicle to the tip of the ES, where basal lamellae form a complete barrier (available at http://zebrafish.link/hildebrand16, ES-specific links presented in methods) (Hildebrand et al., 2017). Basal lamellae were present along the length of the endolymphatic duct. However, cell bodies in the duct remain tightly packed with apical junctions (Figure 4A,E) unlike the apical and lateral membrane separations exposing lamellae in the ES.

Lamellar protrusions at the tip of the ES exist in open and closed configurations.

(A) Select images from serial-section scanning electron microscopy of a 5.5 dpf zebrafish’s right inner ear. Dorsal is up, lateral left, medial right, ventral down, anterior top, and posterior bottom of the series z-stack. Cells forming lamellar barriers were labeled (color overlays, consistent across panels) to highlight connectivity of lamellae. Presented slices are a subset from the series, each separated by 960 nm (Video 8). The lumen of the ES is labeled and outlined with a white dashed line in third panel; perilymph (peri.). (B) Lamellae interdigitate and can form tongue-in-groove structures (inset). (C) Lamellae can interweave. (D) Example of lamellar junctions in an open configuration. (E) Cells in endolymphatic duct have basal lamellae, with the presented duct connecting with ES in panel A. (F) 3D rendering of ES segmentation from serial micrographs shown in panel (A) and Video 8. Black dotted-outline encompasses area of closed, endolymph-filled lamellae. Black mesh highlights areas of membrane overlap between lamellae that are spread open. (G) Electron-dense tight junctions (yellow arrows) present in cells that also have spread basal protrusions. An opening in the apical junctions creates a path from the duct to the basal protrusions (magenta arrow, slices in three panels 1.2 μm apart). (H, H’) lmx1bbjj410/jj410 embryos maintain apical junctions between ES cells (yellow arrows). (I) Mutant ES cells lack basal protrusions (red arrow). Scale bar in (A) is 1000 nm, (F) is 5 μm, and all other scale bars are 500 nm. (B–G) Serial section electron microscopy, n = 2, additional transmission EM, n = 3. (H–I) Serial section EM of mutant, n = 1, transmission EM, n = 3.

https://doi.org/10.7554/eLife.37131.015
Video 8
Serial-section electron micrographs of wild-type ES at 5.5 dpf.

Sections are 60 nm thick and color overlays highlight cells with lamellar barriers or basal lamellae. Second half explores organization of cells in space using cell segmentations.

https://doi.org/10.7554/eLife.37131.016

We identified electron-dense tight junctions between cells at the tip of the ES. These same cells also had lamellar projections (yellow arrows indicate tight junctions, Figures 4G and 3 serial EM sections, 1.2 μm apart, the diameter of the apical opening is ~1.2 μm). The electron-dense signal formed a ring sealing the apical side of these ES cells that was continuous except for an opening that connected fluid from the duct and sac to the exposed lamellae (magenta arrow directed from duct to sac, Figure 4G). Apical junctions were also present in the mutant ES of lmx1bbjj410/jj410 larvae (yellow arrows, Figure 4H,H’). Unlike the wild-type ES, however, the mutant ES appeared to completely lack openings in its apical junctions, as we were unable to identify any in mutants imaged by serial-section scanning EM or transmission EM (four total mutant ears). Consistent with the absence of thin protrusions in the sparsely labeled mutants (Figure 3J), we could not find long lamellar projections on the basal side of the mutant ES (Figure 4I). We were unable to identify apical openings in the wild-type ZO-1 immunostains (Figure 3L). This could be due to difficulty resolving small openings (~1 μm diameter) or to ZO-1 remaining present at the openings such that the apical barrier could reform after ES deflation to hold hydrostatic pressure. These data suggest that lmx1bb-dependent activity is necessary for openings to form in the apical junctions of ES tissue and for ES cells to extend thin basal protrusions.

Cells stretch during ES inflations

The stress from increased pressure in the otic vesicle likely causes an increased ES volume by inducing cell stretching or expansion of the lamellar barrier. To distinguish how the tissue behaves through cycles of ES inflation and deflation, we sought better resolution during live imaging using lattice light-sheet microscopy (LLSM), which generates thin light-sheets using Bessel beams to enhance axial resolution while minimizing phototoxicity and bleaching (Chen et al., 2014; Gao et al., 2014). When using LLSM to image the ES, emitted light passes through brain tissue that scatters and refracts light in a complex, spatially uneven manner. Recent advances in the application of adaptive optics (AO) to microscopy can compensate for these aberrations (Wang et al., 2014). A microscope combining live-cell lattice light-sheets and adaptive optics was built (Liu et al., 2018) and used here to image ES cycles with significantly improved spatial and temporal resolution and reduced photo-damage owing to 2–4 times less laser power being distributed over a large volume (the imaging plane is volumetrically ~105 times larger than the confocal point).

LLSM requires the illumination plane and optical axis to be perpendicular to one another, so we developed a mold for an agarose mount shaped like a volcano that positions the embryo in the desired orientation (Figure 5A). The LLSM produced slightly blurred images with low signal-to-noise (SNR, Figure 5B) without adaptive optics correction. Adding the correction with an AO-LLSM system adequately compensated for tissue aberrations and produced images with higher SNR and better contrast. The resulting high-quality images were then analyzed using our software, ACME, to reconstruct membrane signals (Figure 5C), segment cells from the lumen of the ES (Figure 5D,E), and track cells and lumen accurately through time (Mosaliganti et al., 2012). As with confocal live imaging, we observed cycles of the ES lumen inflating and deflating (Figure 5F,G; Videos 910).

AO-LLSM reveals dynamics of ES cells.

(A) Illustration of AO-LLSM mounting strategy for imaging ES using volcano mount. (B) Representative LLSM images without adaptive optics (AO), with AO, and with AO followed by deconvolution. (C) Three orthogonal views of ACME membrane reconstruction. (D) Three orthogonal views of raw fluorescence signal overlaid with cell segmentations and ES lumen segmentation (magenta), n = 4. (E) 3D rendering of segmented cells and ES lumen (magenta). (F) Volume measurements of segmented ES lumen, imaged every 30 s for over 3 hr (Videos 810). Blue arrowheads point to time points presented in G and I. (G) Top, dorsal perspective displaying 3D renderings of segmented cells. Bottom, maximum intensity projections (MIP) of 4.5 μm slab through tip of ES shows raw data of the ES for the same time points. (H) Secondary axis presents green cell’s thickness versus time (green cell in (G)). Again, primary axis is volume of ES lumen for comparison. (I) Top, 3D renderings of just green cell from (G) and magenta lumen highlight stretching of cell, dorsal-medial perspective. Bottom, centered cross-sectional view of raw data overlaid with green cell’s and ES lumen’s segmentations for the same time points. (J) Secondary axis is plot of grey cell’s thickness (grey in (G)). (K) Top, 3D renderings of only grey cell and magenta lumen. Bottom, centered cross-section view of raw data overlaid with grey cell’s and ES lumen’s segmentations for same time points. (L) Scatter plot of results from Spearman correlation test of cell thickness trajectories and ES lumen volume trajectories for individual inflation and deflation intervals that are monotonic (example intervals bracketed in H and J). Green region highlights significant correlation (p-value<10−3) between cells thinning during inflation or thickening during deflation. Green arrowhead points to test result for bracketed interval in H. Grey region highlights instances where there is no significant correlation between the trajectory of cell thickness and lumen volume. Grey arrowhead points to test result for bracketed interval in J (grey arrowhead). Magenta region highlights significant correlation between cells thinning during deflation or thickening during inflation. Magenta arrowhead points to test result for bracketed interval in J (magenta arrowhead). Y-axis was capped at 10 so that all values greater than or equal to 10 plot as 10, n = 99. All scale bars 10 μm.

https://doi.org/10.7554/eLife.37131.017
Video 9
Slab view of ES time course acquired with lattice light-sheet microscopy with adaptive optics.

15 sequential slices (300 nm slice spacing) were combined as a maximum intensity projections (MIP) to make a 4.5 μm slab. 7 sequential 4.5 μm slabs were tiled to consolidate the presentation of a complete 3D time course. The membrane citrine signal is green and the 3 kDa dextran-Texas red perilymph highlighter is magenta. Lower right panel is an annotated reference, with the basal surface of the ES labeled and outlined with a dotted yellow line, the apical interface enclosing endolymph outlined with a dotted blue line, the endolymph within the ES lumen indicated with a blue arrow, exposed basal lamellae indicated with a green arrow, and the perilymph labeled with magenta text. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.018
Video 10
3D rendering of tracked and segmented cells and ES lumen.

An anterior view on the left and dorsal view on the right. Segmented ES lumen is colored magenta. All other objects are ES cells. Labeled cubes indicate body axes. Same time course as Video 9.

https://doi.org/10.7554/eLife.37131.019

Cell bodies within the ES stretch in response to increasing pressure. While the increased resolution of AO-LLSM did not allow us to resolve overlapping basal lamellae (thickness ~40 nm), it did enable segmentation of individual cell bodies (lengths and thicknesses ~1–10 μm). We found that some cells stretched and thinned when the ES lumen was inflated and thickened upon deflation, likely a result of their elastic properties (Figure 5H,I). However, there were instances when the cells at the tip of the ES did not thin during inflation, or while they thinned during some events, they did not thin during others (Figure 5J,K). To quantify these correlations and obtain an overview of the range of behaviors, we determined the Spearman correlation coefficient between trajectories of lumen volume and cell thickness for intervals that spanned individual inflation and deflation events (Figure 5L, bracketed examples in Figure 5H,J). For 43 of 99 tested trajectories, either cell thinning significantly correlated (p-value less than 10−3) with inflation or cell thickening significantly correlated with deflation (green region, Figure 5L, example of significantly correlated interval, bracket with green arrowhead, Figure 5H, same data point indicated with green arrowhead in Figure 5L). For 42 of 99 tested trajectories, cell behavior did not significantly correlate with inflation or deflation (grey region, Figure 5L, example of uncorrelated inflation interval, bracket with grey arrowhead, Figure 5J, same data point indicated with grey arrowhead in Figure 5L). Unexpectedly, for 14 of 99 tested trajectories there was significant correlation between inflation and cell thickening or deflation and cell thinning (magenta region, Figure 5L, example of correlated interval, bracket with magenta arrowhead, Figure 5J, same data point indicated with magenta arrowhead in Figure 5L). While each cell’s behavior is varied, these data suggest that stretching of a subset of cells contributes to regulation of increased pressure by allowing the ES lumen to expand. Additionally, some cells appear to be pushed away from the ES lumen during inflation and pulled back into the ES epithelium during deflation. The complexity of the response is likely the result of features we did not resolve such as the dynamics of each cell’s basal lamellae, each cell’s residual apical and lateral adhesion, and each cell’s basal interface with the extracellular matrix.

Basal lamellae are dynamic

The low temporal resolution of our original confocal time courses made it unclear whether basal lamellae are static structures like floodgates, only opening to release endolymph, or dynamic. 3D rendering of the AO-LLSM signal shows that some lamellae are rapidly extending and retracting like they are crawling over neighboring cells or neighboring lamellae (Video 11). The basal lamellae are composed of two juxtaposed membranes and can overlay abutting membranes of adjacent cells or additional lamellae (Figure 4). While we observed stacked membranes by EM, they are still unresolved by AO-LLSM. However, one might expect patches of increased membrane signal several times the brightness of that of a lone cell membrane. In 3D renderings of the time course we see dynamic patches of increased intensity that can shift rapidly (Figure 6A, Video 11). By merging 3D renderings of 3 consecutive time points, 30 s apart, in red, blue, and then green, we observed the relative displacement of these lamellae that can crawl a few micrometers per minute (Figure 6B,C).

Basal lamellae are dynamic.

(A) 3D rendering of AO-LLSM data, three sequential time points 30 s apart (membrane citrine depicted in grey, Video 10). Bright patches on surface move (red, blue, green arrows). (B) Consecutive 3D renderings overlaid as red (71:18:00), blue (71:18:30), and then green (71:19:00). Immobile regions remain grey, while regions of displacement are red, blue, and green. Arrows point to moving lamellae. (C) Additional example of moving lamellae, again visualized by overlaying consecutive images in red (70:08:30), blue (70:09:00), and then green (70:09:30). Arrows point to moving lamellae. (D) Top, time points of 3D rendered segmentations spanning 30 min. Below, 4.5 μm MIP slabs of the raw data for the same time points. For both views, arrow points to same region where lumen segmentation is slowly exposed as thin lamellar region expands. (E) Top, time points of 3D rendered segmentations spanning 2 min. Below, 4.5 μm MIP slabs of the raw data for the same time points. For both views, arrow points to same region where lumen segmentation rapidly inflates as thin lamellar region expands. (F) Consecutive 3D renderings, same as (E), overlaid as red (71:46:00), blue (71:46:30), and then green (71:47:00). Arrows point to rapidly inflating lamellae. (G) Consecutive 3D renderings, overlaid as red (70:48:30), blue (70:49:00), and then green (71:49:30). Arrows point to rapidly inflating lamellae, same region as (F). (H) 4.2 μm MIP slabs of a time point from a time-lapse of wild-type embryos expressing myosin GFP (top) mCherry-utrophin (middle), and merged (myosin, green, utrophin, magenta, see Video 12), n = 4. (I) 4.2 μm MIP slabs of a time point from a time-lapse of lmx1bb crispant embryos expressing myosin GFP (see Video 12), n = 2. (J) Dilated mutant ES collapses following puncture of the otic vesicle with a tungsten needle, n = 5. All scale bars 10 μm.

https://doi.org/10.7554/eLife.37131.020
Video 11
3D rendering of membrane citrine signal.

The video begins with an annotated time point from the 3D rendering of signal from an AO-LLSM time course. A yellow dotted line highlights the rendered ES, green arrows point to basal lamellae, and the surrounding space is labeled as perilymph with magenta text. Dorsal view of ES, membrane citrine signal rendered in 3D. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.021

While the resolution of AO-LLSM is improved, it does not resolve the double membrane structure of lamellae or enable identification of the cell from which a lamella extends. Thus, in our surface-rendered segmentations, we give the cell membranes of each cell body a unique pseudo-color while the lamellae that are just exposed to the lumen and not adjacent to another cell body are collectively colored magenta (Figure 5E,G, corresponds to black dotted outline in Figure 4F). By visualizing the segmented objects in 3D we can identify when lamellar barriers are exposed based on when and where the magenta appears (Figures 5G and 6D, Video 10). Inflations of the ES lumen coincided with slow increases in the surface area of exposed lamellae (30 min expansion, Figure 6D). The expansion of thin membrane signal was also observable in the raw data (bottom portion of panel, Figure 6D). Some secondary loci included smaller lamellae that engaged and inflated much more quickly (1–2 min expansion, Figure 6E). We can visualize their sudden expansion by again merging consecutive time points, 30 s apart, with red, blue, and then green (Figure 6F). This presentation reveals that the same locus can rapidly expand multiple times (one minute rapid expansions an hour apart, Figure 6F,G). In summary, multiple sets of basal lamellae can simultaneously engage at different loci, either through slow spreading or rapid expansion.

Apical and basal myosin during ES inflation cycles

Cells and tissues modulate tension for crawling, retracting, bending, constricting, rounding-up, and resisting stretch by organizing an extensive variety of complexes of actin filaments, actin binding proteins, and myosin motors (Grill, 2011; Munjal and Lecuit, 2014; Rafelski and Theriot, 2004; Stewart et al., 2011). Myosin distribution correlates strongly with contractile forces (Fournier et al., 2010). Therefore, to begin to determine how myosin and actin might contribute to tension during ES inflation and deflation, we imaged myosin and actin using non-muscle myosin light chain fused to eGFP and utrophin, an actin binding protein, fused to mCherry (confocal microscopy, Figure 6H, Video 12). Myosin localized to the apical domains of ES cells as well as to dynamic puncta at the basal membrane throughout both inflation and deflation of the ES. During ES inflation, contraction of apical myosin likely counteracts the stress of hydrostatic pressure and maintains the integrity of apical junctions (Rauzi et al., 2010). Tension from apical myosin, strain from hydrostatic pressure, and the adhesion strength between junctional protein complexes likely determines the elastic limit of apical junctions. The crossing of this elastic limit may cause the small apical openings observed in Figure 4G, which ultimately communicates the otic vesicle’s hydrostatic pressure to the lamellar protrusions. The dynamic spots of basal myosin could indicate two possible activities: contraction to retract lamellar protrusions as observed during cell migration or contraction at focal adhesions. Utrophin localized strongest at apical domains of ES cells (Figure 6H), but unfortunately bleached too quickly to generate any meaningful insights into the potential of actin localization dynamics. It had been shown that utrophin based actin highlighters do not localize to actin within lamellae (Belin et al., 2014), which explains the relatively low amounts of basal utrophin signal. In contrast, fluorescent phalloidin labels actin filament enrichment at both apical and basal domains of ES cells (not shown). Our observations of consistent apical localization of myosin throughout ES cycling suggests that the same contractility both counteracts hydrostatic pressure during inflation and drives deflation when hydrostatic pressure drops as the relief valve is opened. We did not observe any sudden changes in myosin localization or behavior that would indicate regulation for either inflation or deflation. Analysis of myosin localization in lmx1bb crispants (CRISPR-Cas9 knockout embryos) revealed similar apical localization and dynamic basal puncta (Figure 6I, Video 12, representative of 2 time-lapses). To determine whether the mutant ES tissue has elastic material properties, we punctured the otic vesicle with a tungsten needle and found that the distended ES rapidly collapsed (5/5 mutant puncture experiments caused ES collapse, Figure 6K). Global inhibition of actin or myosin with cytochalasin D or (S)-nitro-blebbistatin correlated with ES deflation. However, it is currently impossible to determine whether this is owed to hydrostatic pressure not being maintained by the otic vesicle epithelium.

Video 12
Slab view of myosin-GFP during ES time course.

Seven sequential slices (600 nm slice spacing) were combined as a maximum intensity projections (MIP) to make a 4.2 μm slabs. 4 sequential 4.2 μm slabs, starting with the distal tip of the ES on the left, were tiled to consolidate the presentation of a complete 3D time course. On top is the time lapse of a wild-type embryo expression myosin-GFP. Below is an lmx1bb crispant expressing myosin-GFP. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.022

Given the finding that ES lamellae are dynamic with myosin puncta and phalloidin stain, we asked whether Rac1, known to promote actin polymerization in lamellipodial protrusions, was important for ES valve function (Waterman-Storer et al., 1999). Heat-shock induction, throughout the embryo, of a dominant negative Rac1 at 56 hpf resulted in otic vesicles that became leaky and subsequently collapsed between 60 and 70 hpf (Video 13, 19/34 leaky and collapsed ears versus 1/12 in a heat-shock gfp control)(Kardash et al., 2010). In contrast, heat shock induction of the dominant negative Rac1 at 32 hpf, prior to ES formation, did not result in leaking or ear collapse during the subsequent 15 hr (while 0/20 leaky or collapsed ears versus and 0/14 in a heat-shock gfp control). These results are consistent with an important role for the basal lamellipodia of the ES for maintaining barrier integrity during inflation and deflation cycles. Precise analysis of the roles of actin and myosin networks during ES inflation cycles will require either the identification of regulators specific to the ES or actomyosin perturbations specific to the ES.

Video 13
Representative heat shock dnRac1 time course.

Embryos were heat shocked at 55 hpf. At 57 hpf, α-bungarotoxin protein and 3 kDa dextran-Texas red were injected into the hearts. Time course began at 58 hpf, membrane citrine is green, perilymph is magenta. Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.023

Basal lamellae open for deflation

While we had evidence that lamellae can separate in our EM images and evidence that the epithelial barrier breaks during deflation, as seen in the leak in fluorescence from labeled perilymph, it remained uncertain whether separation of lamellae precipitated the deflation events. To observe evidence of the lamellar valve opening, we looked for instances when the membrane signal of lamellar barriers was disrupted prior to deflation events using AO-LLSM. We were able to witness instances when membrane signal from inflated lamellae was disrupted prior to deflation (yellow arrows, Figure 7A–D, lumen magenta, adjacent cells colored in at first at last time-points). These events can be accompanied by a thin lamellar protrusion sticking out into the perilymph (arrow, Figure 7D). Examination of the raw and rendered segmentations prior to and following breaks in membrane continuity showed that it preceded a deflation event (reduction in local volume of ES lumen, magenta, Figure 7A,C). To confirm that the small openings cause the rapid expulsion of endolymph during deflation events, we more closely examined time-lapse movies with labeled endolymph (Figure 7E, yellow endolymph volume before and after two rapid openings). Close examination of the path of endolymph release revealed a small opening through which endolymph travels out into the periotic space (x-y and y-z planes at relief valve openings, yellow arrows, Figure 7E). Narrow paths of pressure release were observed in all high time-resolution (20 s) time-lapse movies of endolymph dynamics (four fish in Video 14 exhibit 16 opening events).

Basal lamellae open prior to deflation.

(A,C) 3D renderings of segmented cells and ES lumen. (A) Dorsal view. (B) Raw data of membrane citrine AO-LLSM data spanning same time range as in (A). First and last time point are overlaid with segmentations for cells and lumen (magenta) neighboring the region of valve opening. In (A–D) arrows point to site of lamellae separating, dotted outlines indicate field of views in B and D, en. indicates endolymph lumen, peri. indicates perilymph. (C) Anterior view of another instance of lamellae separating. (D) Raw data spanning time range of (C). (E) Dorsal view of endolymph volume within ES lumen and time points before and after two release events. Middle and right panels show x-y and y-z ortho-planes of the isolated release sites, highlighted with yellow arrows (See Video 14), n = 8 (F) Illustrated pressure relief mechanism.

https://doi.org/10.7554/eLife.37131.024
Video 14
Endolymph time course at high time resolution reveal sites of release.

Four embryos with 3 kDa dextran-Texas red injected into the otic vesicle. First three are 3-D rendered volumes from a dorsal view. The fourth fish is 3-D rendered from a transverse view (as in Videos 4 and 5). Scale bar is 10 μm.

https://doi.org/10.7554/eLife.37131.025

Discussion

We report identification of the ES as a pressure relief valve based on six lines of evidence. First, the lumen of the ES slowly inflates and then rapidly deflates every ~0.3–4.4 hr. Second, deflation of the ES coincides with a breach in its epithelial diffusion barrier. Third, laser ablation of the otic epithelium is sufficient to induce rapid deflation of the inflating ES. Fourth, the ultrastructure of the closed and opened relief valves in the ES shows that lamellar barriers can pull apart. Fifth, high temporal and spatial resolution time course imaging reveal lamellae that behave like lamellipodia: constantly crawling over one another before they separate to relieve pressure and excess volume. Sixth, perturbation of the valve’s development with a genetic mutation causes distension of the ES as found in common ear disorders such as Ménière’s disease, Pendred syndrome, and Enlarged Vestibular Aqueduct syndrome (Levenson et al., 1989Jackler and De La Cruz, 1989Belal and Antunez, 1980Schuknecht and Gulya, 1983).

Upon first observing the inflation and deflation cycles of the ES, we postulated that the underlying mechanism could either be a response to organ-wide hydrostatic pressure within the otic vesicle, a local tissue behavior where the ES inflates with fluid from the perilymph, or a local tissue behavior where cells in the ES periodically coordinate their movements to expand the ES volume. Our ablation experiments indicated that pressure is transmitted to the ES from the OV and that the tissue is elastic because it collapses quickly when the stress from hydrostatic pressure is removed (Figure 2E–F). The second and third models lack support: dye tracing shows that during inflation the ES is filled with endolymph from the otic vesicle, while perilymph only enters the ES during deflation when the epithelium is open. We did not observe coordinated cell movements within the ES other than stretching. Additionally, myosin appears to act consistently throughout the ES cycles likely providing a constant apical tension that both resists pressure-induced stretch during inflation and drives collapse during deflation by contributing to tissue elasticity. This constant activity precludes the need for sensors or signals to tell the valve when to inflate, when to open, and when to collapse. Furthermore, dynamic basal puncta of myosin may provide integrity to adhesion between basal lamellae with added tension (Ingber, 1997). Throughout our imaging experiments, whether the contrast was produced at plasma membranes, endolymph, or perilymph, we observed what appear to be small pockets of lumen that pinch off from the rest of the ES lumen (see Figures 2D,G, 5G, 6D–E and 7E, and Videos 4, 5, 9, 10, 11 and 14). These pockets were not present in the lmx1bb mutant, (Figure 3E). The simplest explanation is that fluid gets trapped in the lateral intracellular space as the apical junctions open but before the basal lamellae open or when the basal lamellae close after a deflation event. This is consistent with how the inclusions are often resolved in the next inflation cycle. Incorporating all of our observations, we surmise that increased pressure is managed through a combination of strain that stretches viscoelastic cells in the ES and adhesion distributed across the surface area of dynamic basal lamellae. Excess pressure and volume is then released when small openings form amongst compliant apical junctions of ES cells that transmits the stress of the ear’s hydrostatic pressure to the basal lamellae that hold until they separate to release excess pressure and volume. The elastic properties of the tissue then drive its deflation, basal lamellae reunite, and the homeostatic cycle begins again (Figure 7F).

A physical relief valve in the ES, composed of overlapping basal lamellae, had not been previously identified because of the lack of sufficient temporal and spatial resolution as well as the lack of a system with an optically accessible ES. Thin overlapping basal junctions are seen at the marginal folds of lymph and blood capillaries (Baluk et al., 2007). Capillaries flank the developing and adult ES and the similarity between the cross-sectional ultrastructure of lamellar barriers and capillaries, as well as vacuoles, could have led to inaccurate annotation of lamellar barriers (Kronenberg and Leventon, 1986). EM micrographs of the rat, guinea pig, tree frog, and human ES showed instances of thin cytoplasmic extensions enclosing large lumens, which were annotated as capillaries or large vacuoles (Bagger-SjobackSjöbäck et al., 1986; Bagger-Sjöbäck and Rask-Andersen, 1986; Dahlmann and von Düring, 1995; Kawamata et al., 1987; Møller et al., 2013; Qvortrup et al., 1999). More recent EM studies of the human ES, which used more refined techniques to prepare samples from adult cadavers, recognized similar structures as belonging to the distal portion of the ES and described them as interconnected lumen resembling a network of cisterns (Møller et al., 2013). While appearing more elaborate in humans, the structural unit of the relief valve seems to be conserved. However, because of insufficient spatial and temporal resolution, it remains to be determined whether these cells have overlapping lamellae and whether they function as a physical relief valve.

Primarily, pressure relief prevents endolymphatic hydrops—the buildup of pressure and potential tearing of the ear’s epithelium—that is associated with many inner ear pathologies. Secondarily, low-frequency deflations of the ES may drive intermittent longitudinal flow of endolymph from the cochlea and semicircular canals into the endolymphatic duct and sac. The presence of longitudinal flow had been proposed to explain the accumulation of debris and tracers within the ES and, while never directly observed in an unperturbed inner ear, longitudinal flow can be induced and observed through manipulations of the inner ear (Salt, 2001; Salt and DeMott, 1998; Salt and Plontke, 2010; Salt et al., 1986). The localized inflation and patterned formation of valve cells leads to a localized break at the ES lamellar barriers. The local break could create a transient pressure gradient that leads to flow from other parts of the inner ear towards the endolymphatic duct and sac. The intermittent and slow nature of this flow could explain conflicting interpretations of classic studies using tracer injections where tracer accumulated in the ES after long periods of time but significant flow was absent when observed on short time scales (Guild, 1927; Manni and Kuijpers, 1987; Salt, 2001; Salt et al., 1986). We observed longitudinal flow in action with the expulsion of leaked-in perilymph (Figure 2G, Video 5)

Lmx1bb is a LIM homeobox transcription factor. Mutations in LMX1B cause Nail Patella syndrome in humans and a third of Nail Patella syndrome patients develop glaucoma, a phenotype also observed in mouse models (Cross et al., 2014; Liu and Johnson, 2010; Mimiwati et al., 2006). The mechanism by which glaucoma arises in the disease and in mouse models is not clear; however, the trabecular meshwork and Schlemm’s canal appear abnormal in these cases. Giant vacuoles and pores in Schlemm’s canal may be used to relieve intraocular pressure, and their ultrastructure resembles the lamellar barriers of the ES (Gong et al., 2002). Like the ear, pulsatile pressure relief occurs in the eye, although at the higher frequency of the heart rate, and Schlemm’s canal is a major site of the pressure relief (Ascher, 1962; Johnstone et al., 2011). Additionally, up to half of Nail Patella syndrome patients suffer from kidney disease (Bennett et al., 1973). During podocyte maturation apical junctions are lost, thereby enabling development of the basal slit diaphragm that filters blood (Pavenstädt et al., 2003). In mice lacking LMX1B activity, podocytes fail to lose their apical junctions (Miner et al., 2002). Removal of apical junctions may thus be a specialized mechanism used in organs where elaborations of junction architectures allow controlled release of fluids from one compartment into another. Furthermore, the mouse Lmx1a mutant exhibits defects (Koo et al., 2009) like the fish lmx1bb mutant in that they exhibit similarly enlarged ears. Similarity in its protein sequence and expression suggests that mouse Lmx1a and fish lmx1bb likely have comparable activities. We speculate that LMX1 transcription factors may have a common set of targets in the ES, eye, and kidney involved in apical junction remodeling to regulate fluid flow across epithelia.

We find that apical junctions and basal lamellae in the endolymphatic sac behave like a pressure relief valve to regulate fluctuating volume and pressure that arises from excess endolymph in the inner ear (Figure 7F). The high temporal and spatial resolution of AO-LLSM, combined with image processing tools for segmenting, tracking, and quantifying cell geometries, was necessary to reveal the dynamic cell behaviors underlying the pulsing of the ES. While cells of the ES are immobile epithelial cells, the cell extensions resemble lamellipodia of crawling cells in thickness (40 nm), speed (as fast as ~1 μm per minute), and regulatory mechanism (Rac1) (Abercrombie, 1980). Because of limitations of imaging, it remains unclear how each cell’s basal lamellae, residual apical and lateral adhesion, and basal interface with the extracellular matrix contribute to the physical behavior of the ES. Of related interest is how the crawling lamellae adhere to their substrate in the ES to generate a barrier without gaps during the inflation phase. The force of this adhesion, in combination with the viscoelastic properties of the ES, likely determines the relief valve’s set point for volume and pressure homeostasis. For the inner ears of adult mammals, this set point is likely around 100–400 Pascals, although it is unknown how much the inner ear’s pressure and volume fluctuate (Park et al., 2012). In our mutant over-inflation videos we observed an instance of an incomplete deflation (Video 6) that contrasts with wild-type deflation cycles and mutant ES collapse after the otic vesicle was punctured (Figure 6J). This difference raises the questions of how pathological tearing of otic epithelium from endolymphatic hydrops might lead to hearing and balance symptoms and whether the organization of the endolymphatic duct and sac prevents an influx of low potassium perilymph from coming into direct contact with the stereocilia of sensory hair cells. Determination of the molecular basis of lamellar barrier organization and apical junction dynamics will be necessary to make progress on how the physical relief valve works and how it might fail to cause disease.

Materials and methods

Key resources table
Reagent type (species)
or resource
DesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Danio rerio)ABZIRC, Eugene, ORZFIN ID: ZDB-GENO-960809–7
Strain, strain background (Danio rerio)lmx1bb mutant, ale uchu (jj410 allele)ZIRC, Eugene, OR, PMID: 17574823jj410; ZFIN ID: ZDB-ALT-070426–3Schibler and Malicki, 2007
Strain, strain background (Danio rerio)Tg(actb2:mem-citrine-citrine)hm30Megason lab, PMID: 25303534hm30; ZFIN ID: ZDB-ALT-150209–1Xiong et al., 2014
Strain, strain background (Danio rerio)Tg(actb2:mem-citrine)/(actb2:Hsa.H2b-tdTomato)hm32Megason lab, PMID: 27535432hm32; ZFIN ID:
ZDB-ALT-161213–1
Aguet et al., 2016
Strain, strain background (Danio rerio)Tg(actb2:mem-citrine)/(actb2:Hsa.H2b-tdTomato)hm33Megason lab, PMID: 27535432hm33; ZFIN ID: ZDB-ALT-161213–2Aguet et al., 2016
Strain, strain background (Danio rerio)Tg(−5.0lmx1bb:d2eEGFP)mw10gift from Brian Link's lab, PMID: 19500562mw10; ZFIN ID: ZDB-ALT-091218–2McMahon et al., 2009
Strain, strain background (Danio rerio)Tg(actb2:mem-mcherry2)hm29Megason lab, PMID: 23622240hm29; ZFIN ID: ZDB-ALT-130625–1Xiong et al., 2013
Strain, strain background (Danio rerio)Tg(hsp70:rac1_T17N-p2a-mem-cherry2)hm35Megason lab, rac1 mutant plasmid gift from Raz lab, this paperhm35Kardash et al., 2010
Strain, strain background (Danio rerio)Tg(elavl3:GCaMP5G)a4598gift from Alexander Schier's lab, PMID: 23524393a4598; ZFIN ID: ZDB-ALT-130924–1Ahrens et al., 2013
Strain, strain background (Danio rerio)Tg(actb2:myl12.1-EGFP)e2212gift from C.P. Heisenberg's lab, PMID: 25535919e2212; ZFIN ID: ZDB-ALT-130108–2Compagnon et al., 2014
Strain, strain background (Danio rerio)Tg(actb2:mCherry-Hsa.UTRN)e119gift from C.P. Heisenberg's lab, PMID: 25535919e119; ZFIN ID: ZDB-ALT-151029–2Compagnon et al., 2014
Antibodymouse anti ZO-1Thermo Fisher Scientific, Waltham, MAZO1-1A12
Antibodyrabbit anti collagen IIAbcam, Cambridge, United Kingdomab209865
Antibodyrabbit anti lamininSigma-Aldrich, St. Louis, MOL9393
Recombinant DNA reagentpet-28b-Cas9-Hisgift from Alexander Schier's lab, PMID: 24873830addgene id: 47327Gagnon et al., 2014
Recombinant DNA reagentpmtb-t7-alpha-bungarotoxinMegason lab, PMID: 26244658addgene id: 69542Swinburne et al., 2015
Sequence-based reagentfoxi in situ probes,Danio rerioPCRtemplate + T7 reaction (Sigma)RefSeq:NM_181735Thisse and Thisse, 2014
Sequence-based reagentbmp4 in situ probes, Danio rerioPCRtemplate + T7 reaction (Sigma)RefSeq:NM_131342Thisse and Thisse, 2014
Sequence-based reagentlmx1bb sgRNA (exon 2), Danio rerioannealedoligos + SP6 reaction (NEB)GenBank:CR376762Gagnon et al., 2014
Sequence-based reagentlmx1bb sgRNA (exon 3),Danio rerioannealed oligos + SP6 reaction (NEB)GenBank:CR376762Gagnon et al., 2014
Peptide, recombinant proteincas9 proteinMegason labpurification scheme fromGagnon et al., 2014
Peptide, recombinant proteinalpha-bungarotoxinTocris Bioscience (Bristol, United Kingdom)Tocris catalog number 2133Swinburne et al., 2015
Commercial assay or kitmMessage mMachine T7 ULTRA kitThermo Fisher Scientific, Waltham, MAAM1345
Chemical compound, drugDextran, Texas Red, 3000 MWThermo Fisher Scientific, Waltham, MAD-3329
Chemical compound, drugDextran, Alexa Fluor 488, 10000 MWThermo Fisher Scientific, Waltham, MAD-22913
Chemical compound, drugtricaine methanosulfateSigma-Aldrich, St. Louis, MOE10521
Chemical compound, drugnonenyl succinic anhydrideElectron Microscopy Sciences, Hatfield, PA19050
Chemical compound, drugDMP-30Electron Microscopy Sciences, Hatfield, PA13600
Chemical compound, drug1,2,7,8-diepoxyoctane (97%)Sigma-Aldrich, St. Louis, MO139564
Chemical compound, drugSorensen's Phosphate BufferElectron Microscopy Sciences, Hatfield, PA11600–10
Chemical compound, drugglutaraldehyde, EM gradeElectron Microscopy Sciences, Hatfield, PA16220
Chemical compound, drugparaformaldehydeElectron Microscopy Sciences, Hatfield, PA15710
Chemical compound, drugpotassium ferricyanideSigma-Aldrich, St. Louis, MO702587
Chemical compound, drugosmium tetroxideElectron Microscopy Sciences, Hatfield, PA19140
Chemical compound, druguranyl acetateElectron Microscopy Sciences, Hatfield, PA22400
Chemical compound, drugmaleic acidSigma-Aldrich, St. Louis, MOm5757
Chemical compound, drugacetronitrileElectron Microscopy Sciences, Hatfield, PA10020
Chemical compound, drugTaab 812 ResinMarivac Ltd., Nova Scotia, Canada
Software, algorithmMovingROIExtract, convertFormathttps://github.com/krm15/AO-LLSM
Software, algorithmConvertToMegacapture, GoFigure2ContoursToMesheshttps://github.com/krm15/GF2Exchange
Software, algorithmItk-snapwww.itksnap.org
PMID: 16545965
Yushkevich et al., 2006
Software, algorithmFluorenderwww.sci.utah.edu/software/fluorender.html
PMID:23584131
Wan et al., 2012
Software, algorithmHandBrakehttps://handbrake.fr/
Software, algorithmLabVIEWNational Instruments
Software, algorithmMATLAB (R2014A)www.mathworks.com
Software, algorithmParaViewwww.paraview.org
Software, algorithmGoFigure2Xiong et al., 2013
Software, algorithmFIJI (imagJ)www.fiji.sc
PMID: 22743772
Schindelin et al., 2012
Software, algorithmcellPreprocess, multiscalePlanarityAndVoting3D, DistanceFromMask, resample, MorphologicalErosionOnLabelImageFilter, SizeThreshold, MembraneSegmentation, MembraneSegmentationWithMarkersImageFilter, CellSegmentationStatisticshttps://github.com/krm15/ACME/tree/MultithreadLookup
Software, algorithmZen softwarehttp://www.zeiss.com/microscopy/us/products/microscope-software/zen-lite.html
OtherVolcano mould,‘frosted extreme detail’https://www.shapeways.com/, this paperVolcano400
OtherFemtoJet 4xEppendorf, Hamburg, Germany5253000017
OtherNanojectDrummond Scientific, Broomall, PA3-000-204
OtherTungsten wireSigma-Aldrich, St. Louis, MO267554–9.5G

Zebrafish strains and maintenance

Zebrafish were maintained at 28.5°C using standard protocols (Westerfield, 1993). The Harvard Medical Area Standing Committee on Animals approved zebrafish work under protocol number 04487. Adult zebrafish, 3 months to 2 years of age, were mated to produce embryos and larvae. Adults were housed in a main fish facility containing a 19-rack Aquaneering system (San Diego, CA) and a 5-rack quarantine facility for strains entering the system from outside labs or stock centers. The systems’ lights turn on at 9am and go off at 11pm. Fish matings were set up the night before with males separated from females with a divider in false-bottom crossing cages in a pair wise, or 2 × 2 fashion to maximize embryo yield. The divider was pulled the following morning, shortly after the lights turned on. Egg production was monitored to establish the time of fertilization. Manipulations and observations of baby zebrafish were performed between fertilization and 5.5 days post fertilization.

These studies were performed using the AB wild-type strain, the lmx1bbjj410/jj410 mutant and the following transgenic lines: Tg(actb2:mem-citrine-citrine)hm30, Tg(actb2:mem-citrine)/(actb2:Hsa.H2b-tdTomato)hm32, Tg(actb2:mem-citrine)/(actb2:Hsa.H2b-tdTomato)hm33 (these three alleles were combined for maximal membrane signal, hm32 and hm33 are two separate alleles of the same construct that is composed of two divergent beta-actin promoters, one driving membrane citrine and the other driving histone tdTomato, actb2:Hsa.H2b-tdTomato of these divergent constructs tends to be silenced in transgenic fish and is not useful), Tg(−5.0lmx1bb:d2eEGFP)mw10, Tg(actb2:mem-mcherry2)hm29, Tg(hsp70:rac1_T17N-p2a-mem-cherry2)hm35, Tg(elavl3:GCaMP5G)a4598, Tg(actb2:myl12.1-EGFP)e2212, and Tg(actb2:mCherry-Hsa.UTRN)e119 (Aguet et al., 2016; Ahrens et al., 2013; Compagnon et al., 2014; McMahon et al., 2009; Obholzer et al., 2012; Schibler and Malicki, 2007; Xiong et al., 2014; Xiong et al., 2013).

In situ and fluorescent immunostains

The foxi1 in situs that confirm the identity of the ES tissue (Figure 1B’) is representative of 25 stained embryos. A similar ES tissue labeling was obtained when staining for bmp4 (12 embryos at 55hpf). In situs were performed as previously described (Thisse and Thisse, 2014). The foxi1 probe was made using T7 RNA polymerase (Sigma) off of a template made from cDNA using these PCR primers: ctccATGTTTCTGGAGGGAGAG and TAATACGACTCACTATAGGGagaGATCCGTCCCGGTTGTATATGAG. The bmp4 probe was made using T7 RNA polymerase (Sigma) off of a template made from cDNA using these PCR primers: GTCGAGACATCATGATTCCTGG and TAATACGACTCACTATAGGG aga GAGTCTCCGTTTAGCGGCAGC.

Whole-mount fluorescent immunostains were performed using standard protocols and a mouse monoclonal antibody against ZO-1 (ZO1-1A12, Thermo Fisher Scientific, Waltham, MA), a rabbit polyclonal antibody against collagen II (ab209865, Abcam), and a rabbit polyclonal antibody against laminin (L9393, Sigma). In Figure 3J, the 50 hpf wt collagen stain is representative of 16 stained and imaged embryos, the 50 hpf wt ZO-1 stain is representative of 22 stained and imaged embryos, the 80 hpf wt collagen stain is representative of 11 stained and imaged embryos, the 80 hpf wt ZO-1 stain is representative of 32 stained and imaged embryos, the 80 hpf mutant collagen stain is representative of 7 stained and imaged embryos, and the 80 hpf mutant ZO-1 stain is representative of 6 stained and imaged embryos. In all stained embryos, the membrane contrast comes from the membrane citrine transgenic alleles.

Crispants for lmx1bb in the Tg(actb2:myl12.1-EGFP)e2212 background were made using previously described approaches (Gagnon et al., 2014). sgRNA’s were designed and synthesized to target exon 2 (GTTGCTTGTCCCGACCGCAGCGG) and exon 3 (ACGGAGTGCCATCACCAGGCGG) of lmx1bb. These guides were pooled and combined with purified Cas9 protein and injected into 1 cell stage embryos. Analyses were performed on F0 crispants that exhibited the full mutant phenotype.

Immobilization and time-lapse imaging

Embryos were immobilized with either 50 pg of α-bungarotoxin mRNA injected into the 1 cell embryo or 2.8 ng of α-bungarotoxin protein injected into the heart at 59 hpf (Swinburne et al., 2015). α-bungarotoxin mRNA was synthesized from a linearized plasmid using the mMessage mMachine T7 ULTRA kit (Thermo Fisher Scientific). Subsequently, mRNA was purified using RNAeasy Mini Kit (Qiagen, Hilden, Germany). α-bungarotoxin protein was obtained from Tocris Bioscience (Bristol, United Kingdom). 2.3 nL injections were performed using Nanoject II (Drummond Scientific, Broomall, PA). For tracking perilymph,~10 ng of 3 kDa dextran-Texas red neutral (Thermo Fisher Scientific) was injected into the zebrafish heart at 59 hpf. For tracking endolymph,~1 ng of 3 kDa dextran-Texas red neutral was injected into the otic vesicle using a FemtoJet 4x (Eppendorf, Hamburg, Germany) to minimize the volume injected to ~0.2 nl. When labeling both endolymph and perilymph, 3 kDa dextran-Texas red neutral was used for endolymph and 10 kDa dextran- Alexa Fluor 488 (Thermo Fisher Scientific) was used for perilymph. Immobilization with α-bungarotoxin permits the ear to grow and develop normally (Swinburne et al., 2015).

Paralyzed embryos were cultured in Danieau buffer and mounted in an immersed 1.5% w/v agarose canyon mount or the volcano mount for AO-LLSM, that were generated from custom-made molds. The canyons were 0.4 mm wide and 1.5 mm deep. The volcano mold for AO-LLSM was printed by shapeways.com (New York City, NY). The inner width of the volcano was also 0.4 mm and was printed using their ‘Frosted Extreme Detail’ material. The embryo was placed dorso-laterally in the canyon or volcano so that dorsal portion of the left otic vesicle was either flush with a #1 coverslip placed over the canyon (embryos were tilted approximately 30 degrees laterally from their dorsal axes) or protruding from the mouth of the volcano mold. No coverslip was placed above embryos for AO-LLSM. Homemade hair loops were used to position embryos.

Mounted embryos were imaged on an upright LSM 710 (Carl ZEISS, Göttingen, Germany) with a C-apochromat 40x/NA 1.2 objective (ZEISS). The objective’s corrective ring was adjusted to account for the use of #1 coverslips (0.13–0.16 mm thick). Imaging took place within a homemade foam-core incubator maintained at 28.5°C. Lasers of wavelength 405, 488, 514, and 594 nm were used to image GFP, citrine, Texas red, and mCherry2. A typical imaging session used the following set up: citrine excited with 514 nm laser 5% (Ch1 filters 519–584 nm, gain 885), Texas red excited with 594 nm laser 15% (Ch2 filters 599–690 nm, gain 864), 1.27 μs pixel dwell, one line average, 137 μm pinhole, 1.2 zoom, 458/514/594 beam splitter, and 0.173 × 0.173×1.135 μm voxel scaling. Around 20–30 long-term time courses were attempted to establish the method. The data presented are representative of 8 wild-type time courses and four mutant time courses. Additionally, hundreds of ES distensions in mutants were observed during the cloning of the mutant and phenotype characterization at 72 and 96 hpf.

Laser ablations of epithelial cells in the otic vesicle were performed using a similar imaging set-up as for time-lapse confocal imaging. For the cell ablations a Mai-Tai HP 2-photon laser (Spectra-Physics, Santa Clara, CA) was used. After a target was chosen the 2-photon laser was tuned to 800 nm at 50% power, the pinhole was opened completely, the 690 + beam splitter was selected, and a spot scan was performed for 10,000 cycles. 2 or three spots were targeted in adjacent cells to ensure the wounds disrupted the epithelial barrier. After ablations, time-lapse microscopy was performed with 1-photon microscopy, as described above. The data presented are representative of 4 such experiments as well as seven ablations that were evaluated without time courses, 1 hr after ablation.

For puncturing with a tungsten needle, 78 hpf mutant ears were imaged on the confocal microscope, taken from the microscope to a dissecting microscope where the otic vesicle was rapidly punctured, remounted, and then reimaged on the confocal microscope within 2 min. 0.25 mm tungsten wire (Sigma) was sharpened by electrolysis to make tungsten needles.

For imaging with the adaptive optics lattice light-sheet microscope, zebrafish immobilized with α-bungarotoxin were mounted in a 1.5% low melt agarose volcano mold on a 5 mm coverslip and imaged starting at 68–72 hpf. We submerged the excitation and detection objectives along with the 5 mm coverslip in ~8 ml of 1X Danieau buffer at room temperature (22 ± 2°C). The zebrafish tissues expressing membrane citrine marker and 3 kDa dextran Texas Red fluid phase marker were excited using 488 nm and 560 nm lasers (488 nm operating at ~3 mW and 560 nm operating at 5 mW corresponding to ~16 µW and ~31 µW at the back aperture of the excitation objective) both sequentially exposed for 15 msec. The tissues were excited with 488 nm and 560 nm sequentially by dithering a multi-Bessel light-sheet arranged in a square lattice configuration (corresponding to an excitation inner/outer numerical aperture of 0.517/0.55, respectively). The optical sections were collected by scanning the detection objective with 300 nm steps. Each imaging volume consisted of 131 optical planes capturing 1024 × 1024 pixels, thereby capturing a volume of ~97 µm x 97 µm x 39 µm every 15.5 or 29.9 s using a dual Hamamatsu ORCA-Flash 4.0 sCMOS camera setup (Hamamatsu Photonics, Hamamatsu City, Japan). Prior to the acquisition of the time series data consisting of 300–530 time points, the imaged volume was corrected for optical aberrations (manuscript in preparation). The imaged volumes were deconvolved with experimentally measured point-spread functions measured with 0.17 µm tetraspec beads (Thermo Fisher) excited with 488 nm and 560 nm lasers in MATLAB using Lucy-Richardson algorithm on HHMI Janelia Research Campus’ computing cluster and locally with a 3.3 GHz 32-Core Intel Xeon E5 with 512 GB memory. The LLSM was operated using a custom LabVIEW software (National Instruments, Woburn, MA) on a 3.47 GHz Intel Xeon X5690 workstation with 96 GB memory running Microsoft Windows seven operating system. The images presented are from one AO-LLSM time course that is representative of three other similar acquisitions also obtained using AO-LLSM.

Serial-section electron microscopy

‘Wild-type’ Tg(elavl3:GCaMP5G)a4598 and ‘mutant’ lmx1bbjj410/jj410 larvae at 5.5 days post-fertilization (or 4.5 days for the mutant that begins to suffer from pleiotropic toxicity) were briefly anesthetized with 0.02% tricaine methanesulfonate (Sigma-Aldrich, St. Louis, MO), and subsequently preserved by dissection and immersion into an aldehyde fixative solution (2% paraformaldehyde and 2.5% glutaraldehyde in 0.08M Sorenson's phosphate buffer, Electron Microscopy Sciences, Hatfield, PA). Each specimen was then post-fixed and stained en bloc with reduced osmium solution (1% osmium tetroxide (Electron Microscopy Sciences) and 1.5% potassium ferricyanide (Sigma-Aldrich, St. Louis, MO)) followed by uranyl acetate (1% uranyl acetate (Electron Microscopy Sciences) in 0.05 M maleate buffer). Finally, samples were dehydrated with serial dilutions of acetonitrile in distilled water, infiltrated with serial dilutions of epoxy resin in acetonitrile (Electron Microscopy Sciences), and embedded in low-viscosity resin (Koehler and Bullivant, 1973). The epoxy resin for infiltrations and embedding was composed of 63% nonenyl succinic anhydride (Electron Microscopy Sciences), 35.5% 1,2,7,8-diepoxyoctane (97%, Sigma), and 1.5% 2,4,6-tri(dimethylaminomethyl) phenol (DMP-30, Electron Microscopy Sciences), v/v.

Serial sections through the endolymphatic sac were continuously cut with a nominal thickness of 60 nm using a 45° diamond knife (Diatome, Biel, Switzerland) affixed to an ultramicrotome (Leica EM UC6, Leica Microsystems, Wetzlar, Germany) and collected with an automated tape-collecting ultramicrotome (Hayworth et al., 2014). Field-emission scanning EM of back-scattered electrons was conducted either on a Zeiss Merlin (ZEISS) or a FEI Magellan XHR 400L (Thermo Fisher Scientific) with an accelerating voltage of 5.0 kV and beam current of 1.6–7 nA (Hildebrand et al., 2017). Image registration was performed with Fiji TrakEM2 alignment plug-ins (Saalfeld et al., 2010; Saalfeld et al., 2012; Schindelin et al., 2012). Manual image segmentation was performed with ITK-SNAP (Yushkevich et al., 2006). The data presented are from a single wild-type and single mutant sample because of the time and effort required technology development as well as in sample preparation, acquisition, and processing.

Additional wild-type and lmx1bbjj410/j4410 mutant specimens were analyzed by transmission electron microscopy at the Harvard Medical School EM facility. Samples were fixed in 2.5% paraformaldehyde, 2.5% glutaraldehyde, 0.03% picric acid in 0.1M sodium cacodylate buffer, pH 7.4. Samples were stained for three hours in 1% osmium tetroxide and 1.5% potassium ferrocyanide, washed, and then stained with 1% uranyl acetate in maleate buffer, pH 5.2 for one hour. Samples were then embedded in Taab 812 Resin (Marivac Ltd., Nova Scotia, Canada). Blocks were sectioned with a Leica ultracut microtome to a thickness of 80 nm. Sections were collected on carbon coated, formvar coated, copper slot grids. Sections were further stained with with 0.2% lead citrate prior to viewing on a JEOL 1200EX (JEOL USA, Peabody, MA) using an AMT 2 k CCD camera (Advanced Microscopy Techniques, Corp., Woburn, MA).

Quantification and statistical analyses

Time-lapse confocal data sets were converted from Zeiss’s LSM format to a series of image files (*.png) with a header file containing information on the imaging set up (*.meg). This was performed with a script called lsmtomegacapture available in our open source ‘in toto image analysis tool-kit’ (ITIAT, https://wiki.med.harvard.edu/SysBio/Megason/GoFigureImageAnalysis). The image data sets were then loaded into GoFigure2, open-source software developed in the Megason lab for image analyses with a database (http://www.gofigure2.org; Xiong et al., 2013). In GoFigure2 the ES was manually contoured and these contours were exported as XML-files in the GFX format. Exported contours were modeled in 3D and volumes were quantified using a script called GoFigure2ContoursToMeshes, which uses the power crust reconstruction algorithm (Amenta et al., 2001). 3D meshes were generated in the VTK format and viewed using ParaView (KitWare) (Henderson, 2007). Leak in fluorescence was measured by placing a sphere with radius 1 μm into the lumen of the ES and GoFigure2 recorded the integrated fluorescence intensity.

Automated image analysis of the AO-LLSM data was performed using modified versions of the ACME software, python shell scripts, and ITIAT scripts (Mosaliganti et al., 2012). Starting with 3D tif files, the ACME pipeline includes the following steps, parameters used in (bold):

  1. convertFormat inputFile.tif outputFile.mha spacingX(0.097) spacingY(0.097) spacingZ(0.3)

  2. cellPreprocess inputFile.mha outputFile.mha cellRadius (0.1)

  3. resample inputFile.mha outputFile.mha spacingFactorX(1.5) spacingFactorY(1.5) spacingFactorY(0.5)

  4. multiscalePlanarityAndVoting3D inputFile.mha outputFile.mha iSigmaFiltering(0.25) iSigmaVoting(0.5) alpha(0.3) beta(0.25) gamma(1000)

  5. membraneSegmentation inputResample.mha inputTensorVoting.mha outputSegmentedLabelImage.mha threshold(1.0)

Correcting of segmentations and tracking were done using iterative implementations of the ITIAT scripts membraneSegmentationWithMarkersImageFilter and MorpholigcalErosionLabelImageFilter with manual error corrections done in ITK-SNAP (Yushkevich et al., 2006).

3D rendering of segmented labels and overlays of segmented labels and raw membrane signal were performed using the software ITK-SNAP (www.itksnap.org, [Yushkevich et al., 2006]). 3D visualization of raw membrane data was performed using the software FluoRender (www.sci.utah.edu/software/fluorender.html, [Wan et al., 2012]). Videos were compiled and encoded using a combination of ImageJ and HandBrake ([Schindelin et al., 2012], www.handbrake.fr).

For tracking cell thickness, we used the ITIAT scripts CellSegmentationStatistics to map the coordinates of each cell’s centroid, sizeThresh to extract label image files of just the ES lumen, and DistanceFromMask to calculate the physical distance from cell centroids to the surface of the ES lumen labe, which was then multiplied by two to estimate the height of each cell. Cell and lumen volumes were also determined using CellSegmentationStatistics.

The outputs from DistanceFromMask and CellSegmentationStatistics were loaded into MATLAB for both graphing and further analysis. For determining the Spearman correlation coefficient between lumen volumes we first parsed the data matrix into time windows when lumen volume exhibited approximately monotonic increasing or decreasing values. For these time windows we used MATLAB’s statistical toolbox to determine the Spearman correlation coefficient and its associate p-value. The syntax for this analysis was: [RHO,PVAL]=corr(LumenVolume,CellThickness,'type','Spearman'). This test was performed on 99 data windows. MATLAB was also used to perform a Mann-Whitney-Wilcoxon test on the significance cell thickness differences between wild-type and mutant images (Figure 3).

Data and software availability

The image processing scripts described above are available through Github at the following repositories: https://github.com/krm15/ACME/tree/MultithreadLookup (Mosaliganti, 2016; copy archived at https://github.com/elifesciences-publications/ACME), https://github.com/krm15/AO-LLSM (Mosaliganti, 2017a; copy archived at https://github.com/elifesciences-publications/AO-LLSM) and https://github.com/krm15/GF2Exchange (Mosaliganti, 2017b; copy archived at https://github.com/elifesciences-publications/GF2Exchange)

The full data-set of serial-section EM images are publicly available at resolutions of 4.0 × 4.0×60 nm3 per voxel for the closed valve and 18.8 × 18.8×60 nm3 per voxel for the open valve. To examine the ES anatomy viewers should navigate to http://zebrafish.link/hildebrand16/data/es_closed (closed valve) or http://zebrafish.link/hildebrand16/data/es_open (open valve). This set of multi-resolution serial-section EM image volumes was co-registered (Wetzel et al., 2016) to allow one to navigate the lumen of the endolymphatic duct from the lumen of the otic vesicle to the tip of the ES.

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Decision letter

  1. Gary L Westbrook
    Reviewing Editor; Vollum Institute, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

[Editors’ note: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration. The first decision letter after peer review is shown below.]

Thank you for submitting your work entitled "Lamellar junctions in the endolymphatic sac act as a relief valve to regulate inner ear pressure" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Philine Wangemann (Reviewer #1).

Summary:

The reviewers thought the work was interesting and using impressive technology. Although the results are potentially novel with a possible link to human disease, you will see that the reviewers were not completely convinced of the proposed mechanism, i.e. that breaking of the epithelium drives deflation of the endolymphatic sac. A general concern was that the interpretations go beyond the data. In terms of specific comments, the reviewers wanted better validation of the tight function results, and a better demonstration of the presence or absence of basement membrane. Linking opening of the "lamellar junctions" to the deflation process is key to the overall conclusions, but not completely convincing. For example, the videos were not clear to the reviewers in terms of resolving lamella opening. We recognize that this could be technically difficult, but the conclusions should match what can be discerned from the data. The reviewers did not think the Rac1 experiments were informative and thus may want to consider removing them from the manuscript.

Reviewer #1:

This manuscript describes periodic openings of epithelial junctions in the endolymphatic sac of developing zebrafish after the onset of sensory function. The use of high-resolution 3D light microscopy to investigate the dynamic behavior of cells during development opens a new dimension in our understanding of organ development. I would like to make a few comments:

1) The authors observed thinning of epithelial cells and openings of junctions between epithelial cells of the endolymphatic sac and argue that they observed a novel form of junction which they would like to call 'lamellar junction'. I think that the possibility that dynamic behavior of tight junctions resulted in the observed barrier openings has been dismissed too easily and that a new name is therefore not warranted at this time. The authors' main argument for a novel kind of junction appears to rest on the assumption that the thin regions where cell height approaches a thickness of 1 micrometer are too thin to accommodate tight junctions that separate an apical from a basolateral membrane of the epithelium. Accordingly, the authors write: "In wild-type embryos this region often appeared as thin as a single membrane (1.0+/-0.3 micrometer, mean +/- SD) rather than being thick enough to identify the characteristic apical and basal sides of an epithelium". I would like to point out that other very thin epithelia such as alveolar epithelia have no problem to form tight junctions that separate an apical from a basolateral membrane. The thinness of the cells may therefore not be suitable as demission criterion. Before a novel form of an epithelial junction should be claimed and named, it would be necessary to perform more work. It would be necessary to demonstrate the absence of typical electron-dense structures between cell membranes, absence of congregated tight junction proteins such as claudins, occludins and ZO-1 and lack of segregation of marker proteins such as Na+/K+ ATPase. Great care would need to be taken to avoid fixation artifacts. Demonstration of uniform expression of marker proteins such as Na+/K+ ATPase may provide the necessary evidence to address the thin folds of cell membrane as basal lamellae, a term that implies that these thin folds consist of exclusively of basolateral membrane.

2) Video 5 supports the concept that the ES is continually inflating, however, Video 6 does not. In Video 6, there appears to be at least one set back in volume expansion after time 76:00:00. Observation of Video 6 questions whether the time course in Figure 3D is representative.

3) Comparison of Video 1-4 to Video 8 leads to the impression that cells are swollen in Video 8 which may indicate that AO-LLSM itself or the preparation until the imaging began was more damaging to the tissue than conventional confocal microscopy. Cell swelling including a loss of cytoskeletal integrity and vacuolization may contribute to differences in the apparent elastic properties. Although the centroid-based evaluation of elastic behavior is elegant, the authors are correct to recognize the shortcomings of the technique and to refrain from postulating the presence of multiple cell types distinguished by differences in elastic properties.

4) It is unclear to me what is happening in Video 4? At 76:05.00 a luminal segment appears to get detached before it disappears at 76:17.30. Is the ES fragmenting or are epithelial cells engulfing large vacuoles possibly in a pathologic process?

5) The experiments employing the dominant negative Rac1 mutant are not necessary to arrive at the conclusion that "cytoskeletal regulators are […] used for lamellipodial extension". Nevertheless, the experiments employing Rac1 mutants should be quantified and presented in Volume vs Time graphs. In support of the conclusion, it is important to demonstrate that volumes increase in the Rac1 mutant at least as much if not more than in controls.

Reviewer #2:

The manuscript from Swinburne et al. is an interesting and well-performed piece of work analysing the mechanisms of inflation and deflation of the zebrafish Endolymphatic Sac. The authors combine high resolution life imaging techniques, 3D cell reconstructions, quantitative analysis and EM studies and show for the first time the dynamics of the ES epithelium during inflation and deflation. The authors find lamellar protrusions that might work as a valve and upon breakage contribute to the release of ES pressure and deflation. The authors link the action of Lmx1b gene in this process. The findings are relevant for the understanding the homestasis of endolympha in the inner ear. Disruption of ES function might be involved in relevant inner ear diseases. The work is primarily descriptive at this early stage.

1) One on the main points of the manuscript is that rupturing of the lamellae from ES cells allow the leakage out of fluid from the endolymph to the perilymph, being the main mechanism for deflation of the ES. The leakage is only shown in Figure 2E (75.22) and Video 4. To me, the leakage is not very clear. It seems that the cell membranes are displaced and if there is some leakage is very small and probably cannot account for the large shrinkage of ES volume in deflation. As inflation is due to passage from the main otic vesicle cavity to the ES, the deflation could be also explained by involving a contraction and returning of fluid backwards to the main cavity or to re-absorption by ES cells. Also, it is shown in Figure 1 that at the time of deflation, perilymph can enter the ES. Could this be explained because the ES fluid flows back to the vesicle and then fluid can enter from the perilymph to the ES? This possibility should be addressed by injecting dye in the ES and imaging the flow of the dye. The main point of the paper is to convince that breakage of the epithelium drives deflation, but the real contribution to this event to the deflation is less clear. Major epithelium reorganization takes place during the inflation and deflation that are little explored.

2) In line with this, imaging of F-actin or myosinII during the cycles of inflation and deflation would greatly improve the paper. Is the epithelium contracting during deflation or vice-versa is breakage relevant for releasing tension during deflation?

3) Regarding the Lmx1bb role in inner ear. From the paper, it seems that Lmx1bb is only expressed in the ES which is not and thus the overall expression should be shown. Has the Lmx1bb mutant other defects that should be taking into account? The authors try to make a functional relationship between the phenotype in the ES and the lamellae observed but altogether this data is not too strong. The authors mention that in the Lmx1bb mutant the ES cannot deflate and is continually enlarged. The authors claim that inhibition of deflation, reveals that the diffusion barrier does not break but this is not shown. Moreover, there is not evidences that Lmx1bb is necessary for ES epithelial barrier. From the images Figure 3E it seems that the organization of the ES epithelium is different, but as the authors have performed EM of Lmx1bb mutant ES, they should indicate if lamellae are present. Only one panel is shown of the mutant but should provide a better analysis of the epithelial changes. This data would tell better if indeed loss of the lamellae and increase in the junctional organization of the ES epithelium impairs release of the fluid. In discussion the authors claim that Lmx1b might be regulating the loss of junctional adhesions. Are the separations between cells lost in Lmx1bb mutant? If so how the ES expands? Can the authors image ZO1 localization in the mutants and wild-type during inflation and deflation?

4) The authors mention that the cells stretch upon inflation and perform cell reconstruction with their ACME software. Is the volume of ES cells changing overtime? In Figure 5H changes of ES lumen volume are correlated with changes in ES cell thickness but that should be correlated with mean ES cell volumes. Are ES cells losing fluid during inflation?

The stretching and destretching favours the idea that is not only pressure release by epithelium breakage what triggers deflation but the building up of tension a possible main drive for inflation and deflation of the ES. Again, imaging contractility in wild-type and Lmx1bb mutant might be relevant to discern these two possibilities.

5) I am not too convinced that the data on Rac1 is relevant and informative. If more functional studies on the action of the cytoskeleton is provided, then this data might acquire more relevance, if not I don´t see what adds to the paper.

6) A direct evidence that lamellae opening triggers the leakage is not clear. On one side, the authors mention that they observe separation of lamellae (but this would just be part of the stretching mechanism). On the other side, they indicate that there is leakage out (not clear in Figure 2E). Figure 7 tries to make the link. They show 3D renderings of cells Figure 7A but then indicate that in pink the endolymph is depicted. It is not clear in Figure 7 B and D what is a cell and what is the lumen. The breakage of the epithelium should be indicated in the Figure clearly.

Overall, the authors draw conclusions that lamellae are relevant for opening the epithelium and driving deflation. However, as this data is not straight forward, the lamellae might be relevant for stretching during inflation and contraction drive the inflation- deflation cycles.

Reviewer #3:

This manuscript describes a previously unknown phenomenon of the endolymphatic sac (ES) during zebrafish inner ear development. The authors demonstrated that the ES underwent periodic cycles of inflation and deflation during development. Cells in the ES changed shape dynamically to form lamella projections, which served as a pressure valve to release fluid within the otic vesicle after it has inflated to a certain point. The authors showed that knockout of lmx1b, a gene expressed in the ES, cells within the sac did not undergo cell shape changes and the otic vesicle simply swelled without cycling through the inflation-deflation process. These results are interesting and provide insights into several pathological conditions pertaining to the disruption of fluid homeostasis of the inner ear.

Perhaps, the most puzzling part of the results is the description of perilymph entering into the ES while endolymph was being released under pressure. Have the authors entertained other possibilities or artifacts to account for a rise in the level of fluorescent dyes during deflation? I noticed that during the inflation phase, fluorescent vesicles/droplets were seen in the ES cells, suggesting the possibility that perilymph was being transported into the ear. This observation was not described or discussed in the manuscript. Could the fluorescent dye in the perilymph be transported into the ES cells or endolymph during the inflation phase but somehow the dye concentration increases within the endolymph after the initial endolymph release rather than diffusion of dyes/perilymph into the vesicle at the time of deflation?

Presumably, there is a basement membrane lining the otic vesicle and it is not clear what happens to the basement membrane at the ES and the lamella region. It does not look like there is a basement membrane lining the lamellae from the EM pictures. How would the presence or absence of a basement membrane affect the lamellae opening and closing? Immunostaining for the basement membrane would at least be helpful to determine whether the basement membrane is intact at the endolymphatic duct region but absent or discontinuous at the ES.

1) The ES was described to grow and move towards a more central and medial position during morphogenesis (subsection “The endolymphatic sac exhibits cycles of inflation and deflation”). It would be good to have a video documenting this process and it will complement results shown in Video 1.

2) Based on Video 4 and the summary diagram in Figure 7E, I got the impression that the release valve is at the tip of the ES. However, the light sheet data indicate that the lamella region is highly dynamic suggesting that the pressure relief at the ES could occur at any region of the ES. If correct, then a summary diagram that reflects the entire data is warranted.

3) ES cells in Lmx1b mutant do not undergo lamella formation, which was attributed to the presence of tight junctions. Since tight junctions are normally found in epithelial cells, a more global assessment of tight junctions between wildtype and mutant ES using antibodies to tight-junction proteins or tight-junction proteins coupled to Gfp is warranted.

4) Is Lmx1b expressed in the entire ES? How does the different cellular behaviors described in Figure 5 relate to the expression of lmx1b?

5) Video 4. Please clarify that the labeling of the endolymph at the beginning of the video is the result of a single injection to the otic vesicle rather than two separate injections to the vesicle and ES. It also seems like there could be two events of fluid release from the ES. If so, the text and panels in the figure should be revised accordingly.

[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]

Thank you for resubmitting your work entitled "Lamellar projections in the endolymphatic sac act as a relief valve to regulate inner ear pressure" for further consideration at eLife. Your revised article has been favorably evaluated by Gary Westbrook (Senior Editor), a Reviewing Editor, and three reviewers.

The reviewers are pleased with the revision, but had a few minor issues that need your attention, as outlined below. Your responses will be monitored by the Senior Editor before acceptance.

Reviewer #1:

This revised manuscript is a huge improvement over the previous submission. The authors have gone beyond their call of duty in addressing the issues raised in the previous submission. The additional results of simultaneous labeling of perilymph and endolymph as well as the identification of the Lmx1b-positive cells in the endolymphatic sac provided clarity and added volume to this manuscript.

Further minor suggestions to improve this manuscript are listed as follow:

1) The new color scheme is very helpful. How about making the color scheme consistent in Figure 3I and J as well? Adding a white arrow in 3I and adding genotype information on Figure 3C, D and E will also help readers to get through this information-packed figure.

2) Figure 3L, Anti-collagen staining pattern in WT seems qualitatively different between 50 and 80 hpf. Is this a reproducible result? If so, the staining pattern of the lmx1b mutant resembles the WT in 50 hpf rather than 80 hpf, which suggests an arrest of development in the mutant. These points should be clarified in the text.

3) Figure 3 legend, 3L, eLife probably will require a more specific n number than 6-32.

4) Subsection “Apical and basal myosin during ES inflation cycles”, Do the authors mean video 13 instead of video 11?

5) Discussion section, it may be misleading to say Lmx1a mutant exhibits ES defects like the fish lmx1bb mutant because Lmx1a mouse mutants have no endolymphatic duct or sac. One could say they exhibit similar enlarged ear defects.

6) Introduction, extra brackets.

Reviewer #2:

The authors have improved their excellent manuscript and provided additional data following the reviewers' suggestions. The term "lamellar projections" is indeed a better choice than the initially "lamellar junctions".

Reviewer #3:

The authors have made great efforts to answer the reviewer’s points. They have performed all the experiments requested to address the data that was not clear and that could help to discard other possible scenarios. The experiments have indeed helped to improve substantially the paper, together with major rewriting of the text and incorporation of more videos. The Results section flows very smoothly now and it is to thank that they have also extended the discussion with the three different possibilities of how the cycles of inflation and deflation might be regulated. The role of Lmx1b, as well as of the apical cell remodeling is also more clear in this version.

The paper combines a large set of different techniques, that altogether make a beautiful and also interesting piece of work exploring how hydrostatic pressure might be regulated in the inner ear. This work could be extended to other organs in which fluid homeostasis has to be tightly regulated. The possible role of Lmx1bb in controlling remodeling of apical junctions and lamellae might be relevant in other cellular contexts.

I only suggest that the authors mention in methods section whether the analysis in lmx1bb crispr mutants was done in F1 embryos.

https://doi.org/10.7554/eLife.37131.030

Author response

[Editors' note: the author responses to the re-review follow.]

Reviewer #1:

This manuscript describes periodic openings of epithelial junctions in the endolymphatic sac of developing zebrafish after the onset of sensory function. The use of high-resolution 3D light microscopy to investigate the dynamic behavior of cells during development opens a new dimension in our understanding of organ development. I would like to make a few comments:

1) The authors observed thinning of epithelial cells and openings of junctions between epithelial cells of the endolymphatic sac and argue that they observed a novel form of junction which they would like to call 'lamellar junction'. I think that the possibility that dynamic behavior of tight junctions resulted in the observed barrier openings has been dismissed too easily and that a new name is therefore not warranted at this time. The authors' main argument for a novel kind of junction appears to rest on the assumption that the thin regions where cell height approaches a thickness of 1 micrometer are too thin to accommodate tight junctions that separate an apical from a basolateral membrane of the epithelium. Accordingly, the authors write: "In wild-type embryos this region often appeared as thin as a single membrane (1.0+/-0.3 micrometer, mean +/- SD) rather than being thick enough to identify the characteristic apical and basal sides of an epithelium". I would like to point out that other very thin epithelia such as alveolar epithelia have no problem to form tight junctions that separate an apical from a basolateral membrane. The thinness of the cells may therefore not be suitable as demission criterion.

We did not mean to suggest that this data is a proof that the ES cells are not polarized. We have reworded the presentation of our observations.

subsection “Lamellar barriers appear to form an ES relief valve”:

“In wild-type embryos, this region often appeared as a thin sheet (1.0 ± 0.3 µm, mean ± SD) rather than thick as seen in the un-inflated wild-type ES or the lmx1bb mutants in which two distinct surfaces (apical and basal) were observed (6.2 ± 3.2 µm, the mutant, n = 9, is significantly thicker than wild-type, n =14, with a Mann-Whitney-Wilcoxon one tailed p-value of 4 x 10-5).”

We agree with this criticism regarding the potential role of tight junction dynamics and now fully address the potential role for dynamic tight junctions throughout the manuscript.

Subsection “Lamellar barriers appear to form an ES relief valve”:

“In contrast to junctions between thin aveolar cells, which kiss at their tips with tight junctions, the lamellae from ES cells formed large zones of overlap and sometimes bifurcated to form a tongue-and-groove structure (Figure 4B, inset) or interwoven structures (Figure 4C,F, black mesh highlights area of overlap, dotted perimeter indicates full spread). These structures lacked electron-dense signal, indicating that they were unlikely to contain tight junctions between lamellae.”

Subsection “Lamellar barriers appear to form an ES relief valve”:

“We identified electron-dense tight junctions between cells at the tip of the ES. These same cells also had lamellar projections (yellow arrows indicate tight junctions, Figure 4G, 3 serial EM sections, 1.2 μm apart, the diameter of the apical opening is ~1.2μm). The electron-dense signal formed a ring sealing the apical side of these ES cells that was continuous except for an opening that connected fluid from the duct and sac to the exposed lamellae (magenta arrow directed from duct to sac, Figure 4G). Apical junctions were also present in the mutant ES of lmx1bbjj410/jj410 larvae (yellow arrows, Figure 4H,H’). Unlike the wild-type ES, however, the mutant ES appeared to completely lack openings in its apical junctions, as we were unable to identify any in mutants imaged by serial-section scanning EM or transmission EM (4 total mutant ears).”

A revised Figure 7F.

“Throughout our imaging experiments, whether the contrast was produced at plasma membranes, endolymph, or perilymph, we observed what appear to be small pockets of lumen that pinch off from the rest of the ES lumen (see Figures 2D, G, 5G, 6D-E, 7E, and Videos 4, 5, 9, 10, 11, 14). These pockets were not present in the lmx1bb mutant, (Figure 3E). The simplest explanation is that fluid gets trapped in the lateral intracellular space as the apical junctions open but before the basal lamellae open or when the basal lamellae close after a deflation event. This is consistent with how the inclusions are often resolved in the next inflation cycle. Incorporating all of our observations, we surmise that increased pressure is managed through a combination of strain that stretches viscoelastic cells in the ES and adhesion distributed across the surface area of dynamic basal lamellae. Excess pressure and volume is then released when small openings form amongst compliant apical junctions of ES cells that transmits the stress of the ear’s hydrostatic pressure to the basal lamellae that hold until they separate to release excess pressure and volume. The elastic properties of the tissue then drive its deflation, basal lamellae reunite, and the homeostatic cycle begins again (Figure 7F).”

Before a novel form of an epithelial junction should be claimed and named, it would be necessary to perform more work. It would be necessary to demonstrate the absence of typical electron-dense structures between cell membranes, absence of congregated tight junction proteins such as claudins, occludins and ZO-1 and lack of segregation of marker proteins such as Na+/K+ ATPase. Great care would need to be taken to avoid fixation artifacts. Demonstration of uniform expression of marker proteins such as Na+/K+ ATPase may provide the necessary evidence to address the thin folds of cell membrane as basal lamellae, a term that implies that these thin folds consist of exclusively of basolateral membrane.

We find this criticism valid and have now shown that the epithelium is indeed still polarized with apical junction near the lumen containing endolymph and laminin and collagen adjacent to basal lamellae. Additionally, we now refer to the lamellae as lamellar barriers rather than lamellar junctions, so as not to make unsubstantiated interpretations regarding their molecular organization. Indeed, the cell behavior appears more complicated than we initially presented, as apical junctions appear to still be present and potentially function as an additional site of pressure regulation. There are likely three sites of resistance to flow in response to volume and pressure increases: the endolymphatic duct, the apical junctions of the endolymphatic sac, and the dynamic basal lamellae. For various technical reasons, the live dynamics of these three structures were not and still are not simultaneously observable. We have added a discussion on the nature of the cell behavior relative to its polarity and discuss the potential of a multi-tiered relief valve.

We addressed the organization of the ES tissue on subsection “Lamellar barriers appear to form an ES relief valve”:

“To determine the gross organization of the ES tissue, we stained for basal proteins collagen (Figure 3K-L) and laminin (not shown). We found that the basal surface of the ES tissue faces the perilymph outside surrounding the otic vesicle (Figure 3J,K). We also examined localization of the apical marker ZO-1 and found that it stained the ES tissue that faces the endolymph-filled lumen (Figure 3K-L). This apical-basal organization was also present in the lmx1bb410/410 mutant (Figure 3K-L). The absence of local breaks in the mutant ES, the absence of regional thinning in the mutant inflated ES, and the absence of thin basal protrusions suggests that these thin areas may be structurally relevant to ES pressure relief.”

2) Video 5 supports the concept that the ES is continually inflating, however, Video 6 does not. In Video 6, there appears to be at least one set back in volume expansion after time 76:00:00. Observation of Video 6 questions whether the time course in Figure 3D is representative.

The growth rate in Figure 3D does show reduction in the rate of ES growth. We have a follow-up study that is looking into the pressure at which tissue tearing occurs.

3) Comparison of Video 1-4 to Video 8 leads to the impression that cells are swollen in Video 8 which may indicate that AO-LLSM itself or the preparation until the imaging began was more damaging to the tissue than conventional confocal microscopy. Cell swelling including a loss of cytoskeletal integrity and vacuolization may contribute to differences in the apparent elastic properties. Although the centroid-based evaluation of elastic behavior is elegant, the authors are correct to recognize the shortcomings of the technique and to refrain from postulating the presence of multiple cell types distinguished by differences in elastic properties.

It is always a concern that the act of observation perturbs the object being observed. A major design advantage of AO-LLSM is that light energy is distributed across the focal plane. For instance, the total power of the AO-LLSM acquisitions was 11 µW for the 488nm light-sheet and 22 µW for the 560nm light-sheet at the back aperture. These laser powers were lower than those used for the confocal experiments (~50µW for both the 514nm laser and the 594nm laser). Furthermore, the power of the lattice light-sheet is distributed over a much larger volume within the depth of focus (~105 times more volume) of the detection objective, thus leading to gentler excitation that resulted in little to no photo-damage over duration of imaging. The only difference in sample preparation was the absence of a coverslip on top of the embryo during AO-LLSM. The observed membrane curvature is likely the result of better resolution, better signal to noise, and a different (oblique) angle of observation (necessitated to minimize the tissue through which the excitation and emission light passed, in contrast to the shared excitation and emission path of standard laser scanning microscopy). We have observed photo-toxicity in the past during normal confocal imaging, where we used higher power than what we report in this paper to compensate for different transgenics expressing low levels of fluorescent proteins. In these cases, the ES does not inflate when cells appear like soap bubbles and become opaque as they die. Following the AO-LLSM imaging sessions, embryos appeared healthy (tissue was still transparent, and development continued).

In Liu et al., 2018 there are many examples of healthy cells undergoing AO-LLSM imaging.

We briefly address this concern in the text, subsection “Cells stretch during ES inflations”:

“A microscope combining live-cell lattice light-sheets and adaptive optics was built (Liu et al., 2018 in press) and used here to image ES cycles with significantly improved spatial and temporal resolution and reduced photo-damage owing to 2–4 times less laser power being distributed over a large volume (the imaging plane is volumetrically ~105 times larger than the confocal point).”

4) It is unclear to me what is happening in Video 4? At 76:05.00 a luminal segment appears to get detached before it disappears at 76:17.30. Is the ES fragmenting or are epithelial cells engulfing large vacuoles possibly in a pathologic process?

It appears that fluid can get trapped in a lateral space between apical junctions and basal lamellae. We have added more movies exhibiting this phenomenon. We have no reason to think these are pathological, but likely highlight the simultaneous existence of apical adhesion and a basal lamellar barrier. Two barriers likely make the relief valve more robust and we now discuss this in the Discussion section:

“Throughout our imaging experiments, whether the contrast was produced at plasma membranes, endolymph, or perilymph, we observed what appear to be small pockets of lumen that pinch off from the rest of the ES lumen (see Figures 2D, G, 5G, 6D-E, 7E, and Videos 4, 5, 9, 10, 11, 14). These pockets were not present in the lmx1bb mutant, (Figure 3E). The simplest explanation is that fluid gets trapped in the lateral intracellular space as the apical junctions open but before the basal lamellae open or when the basal lamellae close after a deflation event. This is consistent with how the inclusions are often resolved in the next inflation cycle. Incorporating all of our observations, we surmise that increased pressure is managed through a combination of strain that stretches viscoelastic cells in the ES and adhesion distributed across the surface area of dynamic basal lamellae. Excess pressure and volume is then released when small openings form amongst compliant apical junctions of ES cells that transmits the stress of the ear’s hydrostatic pressure to the basal lamellae that hold until they separate to release excess pressure and volume. The elastic properties of the tissue then drive its deflation, basal lamellae reunite, and the homeostatic cycle begins again (Figure 7F).”

5) The experiments employing the dominant negative Rac1 mutant are not necessary to arrive at the conclusion that "cytoskeletal regulators are […] used for lamellipodial extension". Nevertheless, the experiments employing Rac1 mutants should be quantified and presented in Volume vs Time graphs. In support of the conclusion, it is important to demonstrate that volumes increase in the Rac1 mutant at least as much if not more than in controls.

The Rac1 dominant negative likely impedes the ability of the basal lamellae to hold hydrostatic pressure. When this occurs depends on when openings are present in the apical junctions. The time of apical junction opening formation seems to vary quite a bit between each wild-type ES based on when labeled endolymph can fill the lateral space (Video 4, Video 5 and Video 14). Therefore, it seems incorrect to assume that the ES would inflate to the same volumes as controls. We have added myosin movies that complement this experiment. Additionally, we have provided more detail to the discussion and the cartoon model in Figure 7F regarding the two barriers to pressure release, apical junctions and basal lamellae.

Reviewer #2:

The manuscript from Swinburne et al. is an interesting and well-performed piece of work analysing the mechanisms of inflation and deflation of the zebrafish Endolymphatic Sac. The authors combine high resolution life imaging techniques, 3D cell reconstructions, quantitative analysis and EM studies and show for the first time the dynamics of the ES epithelium during inflation and deflation. The authors find lamellar protrusions that might work as a valve and upon breakage contribute to the release of ES pressure and deflation. The authors link the action of Lmx1b gene in this process. The findings are relevant for the understanding the homestasis of endolympha in the inner ear. Disruption of ES function might be involved in relevant inner ear diseases. The work is primarily descriptive at this early stage.

1) One on the main points of the manuscript is that rupturing of the lamellae from ES cells allow the leakage out of fluid from the endolymph to the perilymph, being the main mechanism for deflation of the ES. The leakage is only shown in Figure 2E (75.22) and Video 4. To me, the leakage is not very clear. It seems that the cell membranes are displaced and if there is some leakage is very small and probably cannot account for the large shrinkage of ES volume in deflation.

We have added more images and movies showing leakage. The ES is flanked by blood vessels, that are fenestrated and it is likely that the expelled dye is rapidly taken away and diluted within the vasculature. The rapid and transient nature of the leaked endolymph is likely the result of dilution and advection into the blood stream. See improved Figure 2, Figure 7, Video 4, Video 5 and Video 14.

As inflation is due to passage from the main otic vesicle cavity to the ES, the deflation could be also explained by involving a contraction and returning of fluid backwards to the main cavity or to re-absorption by ES cells. Also, it is shown in Figure 1 that at the time of deflation, perilymph can enter the ES. Could this be explained because the ES fluid flows back to the vesicle and then fluid can enter from the perilymph to the ES? This possibility should be addressed by injecting dye in the ES and imaging the flow of the dye. The main point of the paper is to convince that breakage of the epithelium drives deflation, but the real contribution to this event to the deflation is less clear. Major epithelium reorganization takes place during the inflation and deflation that are little explored.

We have now added a representative time-lapse movie where both the endolymph and perilymph are labeled, in two different colors. These movies clearly show the expulsion of fluid from the ES into the perilymphatic space (as in the prior Figure 2E and Video 4). We show more instances of the fluid movement and have made an effort to clarify both the presentation in the figure as well as the movies.

Additionally, we have now examined the localization of myosin throughout the inflation and deflation cycle, and there is no evidence for an active pumping behavior that has greater contractility than that present during inflation. Additionally, puncturing of wild-type and mutant otic vesicles is sufficient to cause the elastic collapse of the ES. We now discuss how constant tension could contribute to both the resistance to inflation as well as elastic collapse during deflation.

Subsection “Apical and basal myosin during ES inflation cycles”:

“Cells and tissues modulate tension for crawling, retracting, bending, constricting, rounding-up, and resisting stretch by organizing an extensive variety of complexes of actin filaments, actin binding proteins, and myosin motors (Grill, 2011; Munjal and Lecuit, 2014; Rafelski and Theriot, 2004; Stewart et al., 2011). Myosin distribution correlates strongly with contractile forces (Fournier et al., 2010). Therefore, to begin to determine how myosin and actin might contribute to tension during ES inflation and deflation, we imaged myosin and actin using non-muscle myosin light chain fused to eGFP and utrophin, an actin binding protein, fused to mCherry (confocal microscopy, Figure 6H, Video 12). Myosin localized to the apical domains of ES cells as well as to dynamic puncta at the basal membrane throughout both inflation and deflation of the ES. During ES inflation, contraction of apical myosin likely counteracts the stress of hydrostatic pressure and maintains the integrity of apical junctions (Rauzi et al., 2010). Tension from apical myosin, strain from hydrostatic pressure, and the adhesion strength between junctional protein complexes likely determines the elastic limit of apical junctions. The crossing of this elastic limit may cause the small apical openings observed in Figure 4G, which ultimately communicates the otic vesicle’s hydrostatic pressure to the lamellar protrusions. The dynamic spots of basal myosin could indicate two possible activities: contraction to retract lamellar protrusions as observed during cell migration or contraction at focal adhesions. Utrophin localized strongest at apical domains of ES cells (Figure 6H), but unfortunately bleached too quickly to generate any meaningful insights into the potential of actin localization dynamics. It had been shown that utrophin based actin highlighters do not localize to actin within lamellae (Belin et al., 2014), which explains the relatively low amounts of basal utrophin signal. In contrast, fluorescent phalloidin labels actin filament enrichment at both apical and basal domains of ES cells (not shown). Our observations of consistent apical localization of myosin throughout ES cycling suggests that the same contractility both counteracts hydrostatic pressure during inflation and drives deflation when hydrostatic pressure drops as the relief valve is opened. We did not observe any sudden changes in myosin localization or behavior that would indicate regulation for either inflation or deflation. Analysis of myosin localization in lmx1bb crispants (CRISPR-Cas9 knockout embryos) revealed similar apical localization and dynamic basal puncta (Figure 6I, Video 12, representative of 2 time-lapses). To determine whether the mutant ES tissue has elastic material properties, we punctured the otic vesicle with a tungsten needle and found that the distended ES rapidly collapsed (5/5 mutant puncture experiments caused ES collapse, Figure 6K). Global inhibition of actin or myosin with cytochalasin D or (S)-nitro-blebbistatin correlated with ES deflation. However, it is currently impossible to determine whether this is owed to hydrostatic pressure not being maintained by the otic vesicle epithelium.”

Discussion section

“Additionally, myosin appears to act consistently throughout the ES cycles likely providing a constant apical tension that both resists pressure-induced stretch during inflation and drives collapse during deflation by contributing to tissue elasticity. This constant activity precludes the need for sensors or signals to tell the valve when to inflate, when to open, and when to collapse. Furthermore, dynamic basal puncta of myosin may provide integrity to adhesion between basal lamellae with added tension (Ingber, 1997).”

2) In line with this, imaging of F-actin or myosinII during the cycles of inflation and deflation would greatly improve the paper. Is the epithelium contracting during deflation or viceversa is breakage relevant for releasing tension during deflation?

We agree that the epithelium is not passive in the process of inflation and deflation. We have imaged both utrophin and myosin dynamics. Myosin is present within the apical domain of ES cells throughout both inflation and deflation. Basal myosin is dynamic during both inflation and deflation. All our data suggests that contractility that resists the stress of hydrostatic pressure during inflation is used for the elastic behavior of the ES during deflation. See the changes to the text highlighted above. A full study of the role of actin and myosin’s roles in the ES is beyond the scope of this paper and is an ongoing effort.

3) Regarding the Lmx1bb role in inner ear. From the paper, it seems that Lmx1bb is only expressed in the ES which is not and thus the overall expression should be shown. Has the Lmx1bb mutant other defects that should be taking into account?

We now address expression of lmx1bb in other tissues.

Subsection “Lmx1bb is essential for ES deflation”:

“Earlier in the development of the otic vesicle, lmx1bb is expressed in portions of the nascent semicircular canals and sensory patches, regions of the inner ear that also exhibit abnormal development in the mutant (Obholzer et al., 2012; Schibler and Malicki, 2007). There is no precedent for ES development being dependent on those portions of the otic vesicles and there are many mutants with similar SCC or sensory defects that do not have ES phenotypes (Fekete, 1999; Malicki et al., 1996; Whitfield et al., 1996).”

The authors try to make a functional relationship between the phenotype in the ES and the lamellae observed but altogether this data is not too strong. The authors mention that in the Lmx1bb mutant the ES cannot deflate and is continually enlarged. The authors claim that inhibition of deflation, reveals that the diffusion barrier does not break but this is not shown.

The evidence that the diffusion barrier does not break in the mutant was presented in Figure 3D and supplemental figure for Figure 3—figure supplement 1. On the secondary axis we plotted the leak in fluorescence of perilymph into the ES lumen. Unlike in the wild-type, there was never any leak-in fluorescence. Additionally, we performed new time-lapse experiments where the endolymph was labeled in mutants. We also improved the writing to emphasize the evidence and clarify our interpretation.

Subsection “Lmx1bb is essential for ES deflation”:

“As in the wild-type analysis, we quantified the presence of perilymph leaking into the ES lumen (secondary axes of Figure 3D and Figure 3—figure supplement 1A). In the mutant, however, we never observed perilymph entering the ES. Additionally, we imaged mutants where the endolymph was labeled and did not observe leakage out of the distended ES epithelium (Figure 3E). These findings suggest that the epithelial diffusion barrier remains intact in the mutant ES.”

Moreover, there is not evidences that Lmx1bb is necessary for ES epithelial barrier. From the images Figure 3E it seems that the organization of the ES epithelium is different, but as the authors have performed EM of Lmx1bb mutant ES, they should indicate if lamellae are present. Only one panel is shown of the mutant but should provide a better analysis of the epithelial changes. This data would tell better if indeed loss of the lamellae and increase in the junctional organization of the ES epithelium impairs release of the fluid. In discussion the authors claim that Lmx1b might be regulating the loss of junctional adhesions. Are the separations between cells lost in Lmx1bb mutant? If so how the ES expands? Can the authors image ZO1 localization in the mutants and wild-type during inflation and deflation?

We do not state or find any evidence that Lmx1bb is necessary for the ES barrier. Our findings support a role for lmx1bb in weakening or removing apical junctions to create openings that connect the duct to the basal lamellae. We have performed more EM of the mutants and now present images indicating that tight junctions are present in the wild-type and mutant. However, in the wild-type, there are small apical openings that are not present in the mutant. We have added more EM images and whole mount stains. Unfortunately, it is not currently possible to live-image tight junction proteins in the mutants and wild-type. Ubiquitous overexpression of ZO1-GFP sickens embryos. We are working hard to develop technology to perform these experiments, but this is likely a long-term endeavor.

Subsection “Lamellar barriers appear to form an ES relief valve”:

Sparse mosaic labeling of cells in the wild-type further confirmed the presence of thin basal protrusions in the ES, as well as diversity in their organization (Figure 3I). Similar labeling in the mutant revealed the absence of thin basal protrusions in the ES (Figure 3J).

The following additions have been made to the Results section:

“We identified electron-dense tight junctions between cells at the tip of the ES. These same cells also had lamellar projections (yellow arrows indicate tight junctions, Figure 4G, 3 serial EM sections, 1.2 μm apart, the diameter of the apical opening is ~1.2μm). The electron-dense signal formed a ring sealing the apical side of these ES cells that was continuous except for an opening that connected fluid from the duct and sac to the exposed lamellae (magenta arrow directed from duct to sac, Figure 4G). Apical junctions were also present in the mutant ES of lmx1bbjj410/jj410 larvae (yellow arrows, Figure 4H,H’). Unlike the wild-type ES, however, the mutant ES appeared to completely lack openings in its apical junctions, as we were unable to identify any in mutants imaged by serial-section scanning EM or transmission EM (4 total mutant ears). Consistent with the absence of thin protrusions in the sparsely labeled mutants (Figure 3J), we could not find long lamellar projections on the basal side of the mutant ES (Figure 4I). We were unable to identify apical openings in the wild-type ZO-1 immunostains (Figure 3L). This could be due to difficulty resolving small openings (~1μm diameter) or to ZO-1 remaining present at the openings such that the apical barrier could reform after ES deflation to hold hydrostatic pressure. These data suggest that lmx1bb-dependent activity is necessary for openings to form in the apical junctions of ES tissue and for ES cells to extend thin basal protrusions.”

4) The authors mention that the cells stretch upon inflation and perform cell reconstruction with their ACME software. Is the volume of ES cells changing overtime? In Figure 5H changes of ES lumen volume are correlated with changes in ES cell thickness but that should be correlated with mean ES cell volumes. Are ES cells losing fluid during inflation?

We do not see any evidence of gross loss of fluid by the ES cells. While cell volume measurements from microscopy data exist in the literature, they are generally flawed in that anisotropy in the point spread function makes volume measurements very susceptible to systematic error depending on how anisotropic cells are oriented relative to the imaging plane. Together, our imaging and segmentation are not accurate enough to detect subtle changes in cell volume (~10%).

The stretching and destretching favours the idea that is not only pressure release by epithelium breakage what triggers deflation but the building up of tension a possible main drive for inflation and deflation of the ES. Again, imaging contractility in wild-type and Lmx1bb mutant might be relevant to discern these two possibilities.

We agree that tension is likely critical for both deflation and inflation. This perspective complements our original focus on hydrostatic pressure, as pressure creates the stress that is balanced by tension. Indeed, ablation and puncturing experiments demonstrate that the tissue is elastic. This is most likely because of actomyosin contractility. Imaging myosin through the inflation and deflation cycles shows a constant presence at both apical and basal domains of the ES cells. We now address this perspective in greater detail in the Discussion section:

“Upon first observing the inflation and deflation cycles of the ES, we postulated that the underlying mechanism could either be a response to organ-wide hydrostatic pressure within the otic vesicle, a local tissue behavior where the ES inflates with fluid from the perilymph, or a local tissue behavior where cells in the ES periodically coordinate their movements to expand the ES volume. Our ablation experiments indicated that pressure is transmitted to the ES from the OV and that the tissue is elastic because it collapses quickly when the stress from hydrostatic pressure is removed (Figure 2E-F). The second and third models lack support: dye tracing shows that during inflation the ES is filled with endolymph from the otic vesicle, while perilymph only enters the ES during deflation when the epithelium is open. We did not observe coordinated cell movements within the ES other than stretching. Additionally, myosin appears to act consistently throughout the ES cycles likely providing a constant apical tension that both resists pressure-induced stretch during inflation and drives collapse during deflation by contributing to tissue elasticity. This constant activity precludes the need for sensors or signals to tell the valve when to inflate, when to open, and when to collapse. Furthermore, dynamic basal puncta of myosin may provide integrity to adhesion between basal lamellae with added tension (Ingber, 1997). Throughout our imaging experiments, whether the contrast was produced at plasma membranes, endolymph, or perilymph, we observed what appear to be small pockets of lumen that pinch off from the rest of the ES lumen (see Figures 2D, G, 5G, 6D-E, 7E, and Videos 4, 5, 9, 10, 11, 14). These pockets were not present in the lmx1bb mutant, (Figure 3E). The simplest explanation is that fluid gets trapped in the lateral intracellular space as the apical junctions open but before the basal lamellae open or when the basal lamellae close after a deflation event. This is consistent with how the inclusions are often resolved in the next inflation cycle. Incorporating all of our observations, we surmise that increased pressure is managed through a combination of strain that stretches viscoelastic cells in the ES and adhesion distributed across the surface area of dynamic basal lamellae. Excess pressure and volume is then released when small openings form amongst compliant apical junctions of ES cells that transmits the stress of the ear’s hydrostatic pressure to the basal lamellae that hold until they separate to release excess pressure and volume. The elastic properties of the tissue then drive its deflation, basal lamellae reunite, and the homeostatic cycle begins again (Figure 7F).”

5) I am not too convinced that the data on Rac1 is relevant and informative. If more functional studies on the action of the cytoskeleton is provided, then this data might acquire more relevance, if not I don´t see what adds to the paper.

Good suggestion, we have added actin and myosin experiments that complement the Rac1 experiments.

6) A direct evidence that lamellae opening triggers the leakage is not clear. On one side, the authors mention that they observe separation of lamellae (but this would just be part of the stretching mechanism). On the other side, they indicate that there is leakage out (not clear in Figure 2E). Figure 7 tries to make the link. They show 3D renderings of cells Figure 7A but then indicate that in pink the endolymph is depicted. It is not clear in Figure 7 B and D what is a cell and what is the lumen. The breakage of the epithelium should be indicated in the Figure clearly.

These openings are small (as seen in Figure 4D) and are indicated with yellow arrows. We have added additional labels for endolymph and perilymph. Additionally, we have added data from new time-lapses using labeled endolymph, with higher time resolution, that make the narrow paths of endolymph release more clear (Figure 7E, Video 14).

Overall, the authors draw conclusions that lamellae are relevant for opening the epithelium and driving deflation. However, as this data is not straight forward, the lamellae might be relevant for stretching during inflation and contraction drive the inflation- deflation cycles.

We now discuss this more in the Discussion section. We also discuss fluid being trapped between the apical junctions and basal lamellae (Figure 2, 7, Video 4, Video 5 and Video 14), which indicates that both the apical junction and basal lamellae are barriers to pressure release.

Reviewer #3:

This manuscript describes a previously unknown phenomenon of the endolymphatic sac (ES) during zebrafish inner ear development. The authors demonstrated that the ES underwent periodic cycles of inflation and deflation during development. Cells in the ES changed shape dynamically to form lamella projections, which served as a pressure valve to release fluid within the otic vesicle after it has inflated to a certain point. The authors showed that knockout of lmx1b, a gene expressed in the ES, cells within the sac did not undergo cell shape changes and the otic vesicle simply swelled without cycling through the inflation-deflation process. These results are interesting and provide insights into several pathological conditions pertaining to the disruption of fluid homeostasis of the inner ear.

Perhaps, the most puzzling part of the results is the description of perilymph entering into the ES while endolymph was being released under pressure. Have the authors entertained other possibilities or artifacts to account for a rise in the level of fluorescent dyes during deflation? I noticed that during the inflation phase, fluorescent vesicles/droplets were seen in the ES cells, suggesting the possibility that perilymph was being transported into the ear. This observation was not described or discussed in the manuscript. Could the fluorescent dye in the perilymph be transported into the ES cells or endolymph during the inflation phase but somehow the dye concentration increases within the endolymph after the initial endolymph release rather than diffusion of dyes/perilymph into the vesicle at the time of deflation?

We never observe rapid loss of the cytoplasmic signal of perilymph, just the slow accumulation over time. After labeling perilymph, the first deflation events coincide with little to no accumulation of perilymph dye within the cells, yet still the dye leaked into the endolymph. We modeled the respective roles of advective as well as diffusive flow of the perilymph dye and indeed, the dye could diffuse through a small opening to enter the ES lumen during deflation.

We have also performed new experiments where both the endolymph and perilymph are labeled with different colored fluorescent dyes. These data, as well as an improved discussion of the path of pressure relief, address these concerns.

Figure 2G, Video 5, subsection “Loss of the epithelial barrier is sufficient for ES deflation”:

“To determine whether endolymph efflux occurs at the same time as perilymph influx, we simultaneously labeled the two fluids with different colored dyes and imaged their localization with higher temporal resolution (Figure 2G, Video 5). We found a fluttering behavior underlying deflation in which endolymph leaked out (62:41:20), perilymph leaked in (62:55:20), leaked-in perilymph flushed out with more endolymph efflux (62:55:40), followed by complete deflation (63:12:00).”

Presumably, there is a basement membrane lining the otic vesicle and it is not clear what happens to the basement membrane at the ES and the lamella region. It does not look like there is a basement membrane lining the lamellae from the EM pictures. How would the presence or absence of a basement membrane affect the lamellae opening and closing? Immunostaining for the basement membrane would at least be helpful to determine whether the basement membrane is intact at the endolymphatic duct region but absent or discontinuous at the ES.

We have now immunostained for laminin and collagen and found that they are present but potentially at a reduced amount at the tip of the ES, consistent with the EM micrographs.

1) The ES was described to grow and move towards a more central and medial position during morphogenesis (subsection “The endolymphatic sac exhibits cycles of inflation and deflation”). It would be good to have a video documenting this process and it will complement results shown in Video 1.

We have taken this out of the description as it distracts from the main findings.

2) Based on Video 4 and the summary diagram in Figure 7E, I got the impression that the release valve is at the tip of the ES. However, the light sheet data indicate that the lamella region is highly dynamic suggesting that the pressure relief at the ES could occur at any region of the ES. If correct, then a summary diagram that reflects the entire data is warranted.

Yes, this appears to be a common feature that different lamellae are responsible for different pressure relief cycles. While the initial diagram did not reflect this complexity, we now present this in Figure 7F as well as present the implications of this observation in the discussion, as mentioned above.

3) ES cells in Lmx1b mutant do not undergo lamella formation, which was attributed to the presence of tight junctions. Since tight junctions are normally found in epithelial cells, a more global assessment of tight junctions between wildtype and mutant ES using antibodies to tight-junction proteins or tight-junction proteins coupled to Gfp is warranted.

We have performed better assessment of the tight junctions and find them present in the wild-type and mutant ES. Live imaging of tight-junction proteins in the ES is not currently possible; we look forward to doing these experiments in the future.

4) Is Lmx1b expressed in the entire ES? How does the different cellular behaviors described in Figure 5 relate to the expression of lmx1b?

Lmx1bb appears to be expressed in a subset of the ES. A more thorough analysis of the cell types and molecular states of the ES cells is part of a larger ongoing effort and beyond the scope of this manuscript.

5) Video 4. Please clarify that the labeling of the endolymph at the beginning of the video is the result of a single injection to the otic vesicle rather than two separate injections to the vesicle and ES. It also seems like there could be two events of fluid release from the ES. If so, the text and panels in the figure should be revised accordingly.

We have clarified the video presentation and its discussion in the text.

Subsection “Loss of the epithelial barrier is sufficient for ES deflation”:

To assess whether ES inflations occur by transmission of endolymph pressure between the otic vesicle and the ES, we performed a single injection of a small volume of solution containing 3 kDa fluorescent dextran into the otic vesicle and followed its movement during inflation and deflation.

This has been clarified in the legend of Figure 2:

(B) Time points of individual sagittal slices of raw data from 3D time course (endolymph labeled yellow by single dye injection into otic vesicle, membrane citrine in cyan).

[Editors' note: the author responses to the re-review follow.]

Reviewer #1:

This revised manuscript is a huge improvement over the previous submission. The authors have gone beyond their call of duty in addressing the issues raised in the previous submission. The additional results of simultaneous labeling of perilymph and endolymph as well as the identification of the Lmx1b-positive cells in the endolymphatic sac provided clarity and added volume to this manuscript.

Further minor suggestions to improve this manuscript are listed as follow:

1) The new color scheme is very helpful. How about making the color scheme consistent in Figure 3I and J as well? Adding a white arrow in 3I and adding genotype information on Figure 3C, D and E will also help readers to get through this information-packed figure.

We are glad that the improved color scheme helps. In our imaging experiments we reused sources of contrast in different combinations. As such, there did not exist a color scheme that provides contrast for visualization in all combinations. We did continue with our color scheme of using green for membrane-citrine in Figure 3I and J. We made an exception regarding the color scheme in Figure 3 I and J because, what maximized contrast for other experiments, i.e. green for membrane-citrine and yellow for membrane-mCherry2, would have made it difficult to discern the two channels. One of the main reasons for presenting information with color is to provide contrast between different objects. There is not enough contrast between green and yellow—therefore we chose to maximize contrast with green and magenta. We hope that clear labeling helps to prevent any confusion.

We have added genotype information and white arrows to Figure 3I.

2) Figure 3L, Anti-collagen staining pattern in WT seems qualitatively different between 50 and 80 hpf. Is this a reproducible result? If so, the staining pattern of the lmx1b mutant resembles the WT in 50 hpf rather than 80 hpf, which suggests an arrest of development in the mutant. These points should be clarified in the text.

We clarified these points in the text, in subsection “Lamellar barriers appear to form an ES relief valve”:

3) Figure 3 legend, 3L, eLife probably will require a more specific n number than 6-32.

We now indicate the specific n number for each stain presented in Figure 3L in the figure legend.

4) Subsection “Apical and basal myosin during ES inflation cycles”, Do the authors mean video 13 instead of video 11?

Yes, this has been corrected. Thanks!

5) Discussion section, it may be misleading to say Lmx1a mutant exhibits ES defects like the fish lmx1bb mutant because Lmx1a mouse mutants have no endolymphatic duct or sac. One could say they exhibit similar enlarged ear defects.

We have clarified the text. The work in Koo et al., indicates that the endolymphatic duct is not discernable by paint fill in end-point analyses. It is possible that a distended lumen would cause a nascent duct and sac to lapse into wall of the otocyst but this will await further investigation.

6) Introduction, extra brackets.

Reviewer #3:

The authors have made great efforts to answer the reviewer’s points. They have performed all the experiments requested to address the data that was not clear and that could help to discard other possible scenarios. The experiments have indeed helped to improve substantially the paper, together with major rewriting of the text and incorporation of more videos. The Results section flows very smoothly now, and it is to thank that they have also extended the discussion with the three different possibilities of how the cycles of inflation and deflation might be regulated. The role of Lmx1b, as well as of the apical cell remodeling is also more clear in this version.

The paper combines a large set of different techniques, that altogether make a beautiful and also interesting piece of work exploring how hydrostatic pressure might be regulated in the inner ear. This work could be extended to other organs in which fluid homeostasis has to be tightly regulated. The possible role of Lmx1bb in controlling remodeling of apical junctions and lamellae might be relevant in other cellular contexts.

I only suggest that the authors mention in Materials and methods section whether the analysis in lmx1bb crispr mutants was done in F1 embryos.

We have clarified the text in subsection “In situ and fluorescent immunostains”. We would like to point out that F0 crispants were only used for the myosin-EGFP analysis to avoid a ~3-month delay due to crossing. The other experiments in the paper were done with the established germ-line allele lmx1bbjj410.

https://doi.org/10.7554/eLife.37131.031

Article and author information

Author details

  1. Ian A Swinburne

    Department of Systems Biology, Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Funding acquisition, Investigation, Writing—original draft, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4162-0508
  2. Kishore R Mosaliganti

    Department of Systems Biology, Harvard Medical School, Boston, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  3. Srigokul Upadhyayula

    1. Department of Pediatrics, Harvard Medical School, Boston, United States
    2. Program in Cellular and Molecular Medicine, Boston Children’s Hospital, Boston, United States
    3. Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6911-0270
  4. Tsung-Li Liu

    Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, United States
    Contribution
    Resources, Investigation
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    No competing interests declared
  5. David G C Hildebrand

    Department of Molecular and Cellular Biology, Harvard University, Cambridge, United States
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    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2079-3517
  6. Tony Y -C Tsai

    Department of Systems Biology, Harvard Medical School, Boston, United States
    Contribution
    Investigation
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    No competing interests declared
  7. Anzhi Chen

    Department of Systems Biology, Harvard Medical School, Boston, United States
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    No competing interests declared
  8. Ebaa Al-Obeidi

    Department of Systems Biology, Harvard Medical School, Boston, United States
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    No competing interests declared
  9. Anna K Fass

    Department of Systems Biology, Harvard Medical School, Boston, United States
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    No competing interests declared
  10. Samir Malhotra

    Department of Systems Biology, Harvard Medical School, Boston, United States
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    No competing interests declared
  11. Florian Engert

    Department of Molecular and Cellular Biology, Harvard University, Cambridge, United States
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  12. Jeff W Lichtman

    Department of Molecular and Cellular Biology, Harvard University, Cambridge, United States
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    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0208-3212
  13. Tomas Kirchhausen

    1. Department of Pediatrics, Harvard Medical School, Boston, United States
    2. Program in Cellular and Molecular Medicine, Boston Children’s Hospital, Boston, United States
    3. Department of Cell Biology, Harvard Medical School, Boston, United States
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    Resources, Supervision, Funding acquisition, Writing—review and editing
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    No competing interests declared
  14. Eric Betzig

    Janelia Research Campus, Howard Hughes Medical Institute, Ashburn, United States
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    No competing interests declared
  15. Sean G Megason

    Department of Systems Biology, Harvard Medical School, Boston, United States
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    Conceptualization, Supervision, Funding acquisition, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    megason@hms.harvard.edu
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    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9330-2934

Funding

National Institutes of Health (Fellowship 5F32HL097599)

  • Ian A Swinburne

Hearing Health Foundation (Emerging Research Grant)

  • Ian A Swinburne

Novartis (Fellowship in Systems Biology)

  • Ian A Swinburne

National Institutes of Health (K25 HD071969)

  • Kishore R Mosaliganti

National Institutes of Health (T32 MH20017)

  • David G C Hildebrand

National Institutes of Health (T32 HL007901)

  • David G C Hildebrand

National Science Foundation (IIA EAPSI Award (1317014))

  • David G C Hildebrand

National Institutes of Health (DP1 NS082121)

  • Florian Engert

National Institutes of Health (RC2 NS069407)

  • Florian Engert

National Institute of Neurological Disorders and Stroke

  • Florian Engert

National Institutes of Health (R01 GM075252)

  • Tomas Kirchausen

Janelia Visitor Program

  • Tomas Kirchausen

Howard Hughes Medical Institute

  • Tomas Kirchausen

Biogen

  • Tomas Kirchausen

National Institutes of Health (R01 DC010791)

  • Sean G Megason

National Institutes of Health (R01 DC015478)

  • Sean G Megason

National Institute of Deafness and Other Communication Disorders

  • Sean G Megason

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

Acknowledgements

We thank Dante D’India for fish care. We thank Brian Link and Erez Raz for reagents. We thank Louise Trakimas, Elizabeth Benecchi, and Margaret Coughlin for assistance with electron microscopy at the Harvard Medical School EM facility. Lisa Goodrich, Becky Ward, Amelia Green, Akankshi Munjal, Sandra Swinburne, and the Megason lab for comments. Kerry Sobieski, Betzig lab members, and members of the Janelia Research Campus aquatics facility for helping with travel, shipping, and assisting with experiments at Janelia Research Campus. IAS was supported by NIH fellowship 5F32HL097599, a Hearing Health Foundation Emerging Research Grant, and a Novartis Fellowship in Systems Biology. KRM was supported by NIH grant K25 HD071969. DGCH was supported by NIH grants T32 MH20017 and T32 HL007901 and NSF IIA EAPSI award 1317014. TK acknowledges support from the Janelia Visitor Program, HHMI, Biogen and NIH grant R01 GM075252. SU is a Fellow at the Image and Data Analysis Core at Harvard Medical School. This work was supported by R01 DC010791 and R01 DC015478 from the National Institute of Deafness and Other Communication Disorders (SGM) and by NIH grants DP1 NS082121, RC2 NS069407 from the NINDS (FE).

Ethics

Animal experimentation: This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The Harvard Medical Area Standing Committee on Animals approved zebrafish work under protocol number 04487

Reviewing Editor

  1. Gary L Westbrook, Vollum Institute, United States

Publication history

  1. Received: March 29, 2018
  2. Accepted: May 9, 2018
  3. Version of Record published: June 19, 2018 (version 1)
  4. Version of Record updated: August 15, 2018 (version 2)

Copyright

© 2018, Swinburne 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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