A molecular basis for water motion detection by the mechanosensory lateral line of zebrafish

Chou, Shih-Wei; Chen, Zongwei; Zhu, Shaoyuan; Davis, Robin W.; Hu, Jiaqi; Liu, Li; Fernando, Carol A.; Kindig, Kayla; Brown, William C.; Stepanyan, Ruben; McDermott, Brian M.

doi:10.1038/s41467-017-01604-2

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Published: 21 December 2017

A molecular basis for water motion detection by the mechanosensory lateral line of zebrafish

Shih-Wei Chou^1,2^na1,
Zongwei Chen^1,2^na1,
Shaoyuan Zhu^1,2,
Robin W. Davis^1,2,
Jiaqi Hu^1,2,
Li Liu^1,2,
Carol A. Fernando¹,
Kayla Kindig^1,2,
William C. Brown¹,
Ruben Stepanyan^1,3 &
…
Brian M. McDermott Jr.^1,2,3,4

Nature Communications volume 8, Article number: 2234 (2017) Cite this article

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Abstract

Detection of water motion by the lateral line relies on mechanotransduction complexes at stereocilia tips. This sensory system is comprised of neuromasts, patches of hair cells with stereociliary bundles arranged with morphological mirror symmetry that are mechanically responsive to two opposing directions. Here, we find that transmembrane channel-like 2b (Tmc2b) is differentially required for mechanotransduction in the zebrafish lateral line. Despite similarities in neuromast hair cell morphology, three classes of these cells can be distinguished by their Tmc2b reliance. We map mechanosensitivity along the lateral line using imaging and electrophysiology to determine that a hair cell’s Tmc2b dependence is governed by neuromast topological position and hair bundle orientation. Overall, water flow is detected by molecular machinery that can vary between hair cells of different neuromasts. Moreover, hair cells within the same neuromast can break morphologic symmetry of the sensory organ at the stereocilia tips.

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Introduction

Perception of water motion in fish is initiated by the deflection of mechanosensitive hair bundles that are topographically fixed on the animal’s surface in the lateral line system^1,2,3. For this sense, mechanotransduction occurs at the tips of cellular protrusions, the stereocilia, which have increasing heights and are interconnected by tip links⁴ to comprise the hair cell’s bundle. Mechanically gated ion channels at the tips of the shorter stereocilia are opened when a bundle is deflected in the direction of the tallest stereocilia, permitting cation entry^5,6.

The lateral line system detects two types of water movement through receptive neuromasts on the animal’s surface: water velocity^7,8 and minuscule water vibrations^7,9. Though the anterior lateral line (ALL) bespeckles the larval head in a complex pattern, the posterior lateral line (PLL) generally runs evenly along the trunk and tail¹⁰. Each neuromast houses a sheet of hair cells organized with planar cell polarity (PCP) in a mirror symmetrical arrangement of hair bundles², half in each direction. Hair cells with similarly facing hair bundles are innervated by a dedicated afferent neuron, which carries that group’s information centrally^11,12. Each PLL neuromast’s axis of mirror symmetry is oriented either parallel or perpendicular to the anterior–posterior (A–P) body axis¹³. Understanding the molecular basis of water motion detection requires identification of molecules at the tips of stereocilia that are necessary for mechanotransduction and relating them to the intricate topology of the lateral line. An intriguing class of proteins that may play a role in this relationship are the members of the transmembrane channel-like (Tmc) protein family, known for their intimate association with the mechanotransduction process^{14,15,16,17,18,19,20,21}.

Results and Discussion

A lateral line with two general axes of best sensitivity

Establishing the molecular basis of lateral line function first requires clarifying the anatomical relationships that may exist between neuromasts of the zebrafish head and those of the trunk and tail. Most neuromasts of the head, trunk, and tail lie on approximate single planes or sheets on either lateral side of the fish (Fig. 1a, top); these were the first to be characterized, as they are the most abundant. The topologically complex larval ALL may have neuromasts with organizations similar to cranial neuromasts of other adult fish species, with axes of best sensitivity oriented to multiple directions²². Alternatively, this lateral line may have its neuromasts oriented along just two axes.

Addressing this question required multiple steps. First, we functionally tested whether the axis of best sensitivity within a single neuromast was the axis in which the bevel-shaped hair bundles face opposite directions. Specifically, we asked if stimuli provided along this axis had similar or attenuated responses to those of stimuli administered from an orthogonal direction (Fig. 1b, c). For this experiment, we evaluated the quality of mechanotransduction in neuromast organs by measuring microphonic potentials evoked by sinusoidal stimuli²³. In normal neuromasts of wild-type larvae, a 2f (frequency) response is generally observed because each phase of stimulation activates approximately half of the hair cells, whereas the rest are inhibited. This telltale signature results from the presence of two groups of hair cells with hair bundles facing opposite directions responding to opposing stimuli^2,24 (Fig. 1b, c). The responses from stimuli that we provided along the axis with opposite facing hair bundles were larger than responses from stimuli perpendicular to this axis (Fig. 1d, e), indicating that the axis in which the hair bundles are opposing each other is the axis of best sensitivity for each larval neuromast.

Next, by labeling hair bundle actin of larvae with phalloidin, we showed that most anterior neuromasts’ axes of mirror symmetry are parallel or perpendicular to the A–P axis, similar to the PLL (Fig. 1a, top, f; Supplementary Fig. 1). Using the same method, we determined the axes of best sensitivity for a minor subset of neuromasts that are positioned superior or inferior to the eye (Fig. 1a, bottom). Interestingly, the axes of best sensitivity for the neuromasts above the eye were tangential to the perimeter of the eye (Fig. 1a, bottom left). For three neuromasts below the eye, the axes of best sensitivity were in A–P orientations for two, and one had an axis that was positioned obliquely to the A–P axis (Fig. 1a, bottom right).

Taken together, these data suggest that the majority of neuromasts have axes of best sensitivity that set up a Cartesian coordinate system to organize water motion information along the whole body. Moreover, the consolidation of many potential neuromast sensitivities to just two axes may allow for economical use of a limited number of larval neuromasts, which could facilitate an additive response from a population of neuromasts with hair cells that face the same direction. However, this anatomical arrangement creates a potential conundrum. A water flow stimulus from the front, caused by normal swimming, may require a different response than one originating from the rear, possibly caused by a predator. To detect this difference, neuromasts sensitive in the A–P plane may require breaking morphologic symmetry to differentially encode directional information. We reasoned that a direct mechanism to modify neuromast responses to flow from different directions would be to alter the mechanotransduction apparatus itself.

Posterior neuromasts require Tmc2b for mechanotransduction

Tmc proteins are hypothesized elements of the mechanotransduction apparatus of the lateral line based on established roles of orthologous proteins in mammalian hearing^{14,16,17,18,19,20,21} and expression patterns and dominant negative studies in fish¹⁵. According to RNA in situ hybridization studies of zebrafish larva, tmc2b is robustly expressed in hair cells of the lateral line, but the mRNAs of the paralogs tmc2a and tmc1 are present at much lower levels and cannot be detected by this method. However, their presence is nonetheless observed by more sensitive molecular biological techniques¹⁵. Because of the higher level of expression and the known function of mouse orthologous proteins, we explicitly tested the notion that Tmc2b is necessary for mechanotransduction in the lateral line. First, by transgenesis, we showed that full-length Tmc2b fused to GFP localizes to the tips of neuromast stereocilia (Fig. 2a, b), fulfilling a key requirement of being a component of the mechanotransduction apparatus in zebrafish hair cells. Second, to test whether Tmc2b is necessary for hair cell mechanotransduction in the lateral line system, we performed genome editing with transcription activator-like effector nucleases (TALENs), synthetic restriction enzymes generated by fusing a TAL effector DNA-binding domain to a DNA-cleavage domain (Fig. 2c, d)²⁵. Targeting with the TAL effector pair resulted in a mutant with a 7-bp deletion in exon 4, which was bred to homozygosity (Fig. 2e, f). The tmc2b mutation is predicted to result in a frame shift that yields a premature opal stop codon (UGA), creating a truncated protein of 148 aa, as opposed to the wild-type protein of 940 aa. The mutation is expected to stop all putative transmembrane domains from being produced to create a null (Fig. 2e, g).

Initially, we appraised the quality of mechanotransduction in neuromast organs by measuring stimulus-evoked microphonic potentials²³. In posterior neuromasts with A–P orientations in larvae at 6-days postfertilization (dpf), delivery of a sinusoidal stimulus evoked either no extracellular receptor potentials in tmc2b ^−/− larvae (9 of 15) (Fig. 3a, bottom trace), or a greatly attenuated response (6 of 15) (Fig. 3a, third trace), relative to controls (Fig. 3a first and second trace) (mean potential of tmc2b ^−/− responders ± SEM = 5.2 ± 1.1 µV (n = 6), mean potential of tmc2b ^+/− = 8.4 ± 0.34 µV (n = 15), and mean potential of tmc2b ^+/+ (WT) = 8.0 ± 0.81 µV (n = 5)). Interestingly, the attenuated responses were often asymmetrical, with the 2f response replaced with a 1f response (Fig. 3a, third trace), which was stronger when the hair bundles were deflected anteriorly. In mutants, no defects in hair bundle morphology (Supplementary Fig. 2a–d) or hearing (Supplementary Fig. 2f, g) were observed. However, posterior neuromasts of mutants had a mild reduction in hair cell numbers (approximately one-third decrease, 6 dpf; Supplementary Fig. 2c–e; Supplementary Fig. 3 ). These studies demonstrate that Tmc2b is necessary for normal mechanotransduction in posterior neuromast organs with A–P orientations.

Posterior neuromast hair cells differentially require Tmc2b

To test whether mutation of tmc2b reduces mechanotransduction in some hair cells more than others in posterior neuromasts with A–P orientations, we visualized the uptake of cationic styryl dye FM1-43FX. This fluorescent molecule transits through the hair cell’s mechanotransduction channel, serving as a qualitative indicator of channel function when fixed²⁶ (Fig. 3b, c). Tmc2b ^−/− mutants at 6-dpf showed exceptionally uneven FM1-43FX loading between cells in the same neuromast (Fig. 3c), indicating that the molecular components that are directly or indirectly involved in mechanotransduction can differ between hair cells.

We next examined whether residual mechanotransduction channel function was uniform or heterogeneous among permissive hair cells in posterior neuromasts with A–P orientations in mutants using a multistep process. Quantitative 4-(4-diethylaminostyryl)-1-methylpyridinium iodide (4-Di-2-ASP) hair cell uptake assays were used because this fluorophore traverses mechanotransduction channels^27,28. First, we determined the percentages of hair cells that loaded 4-Di-2-ASP in posterior neuromasts in 6-dpf larvae. 94 ± 1.2% (n = 55) of control hair cells loaded fluorophore; in contrast, 35 ± 1.8% (n = 48) of tmc2b ^−/− hair cells loaded (Fig. 3d), confirming Tmc2b is absolutely required for a subpopulation of hair cells within a single neuromast. For comparison, we determined the percentage of hair cells that took up fluorophore in posterior neuromasts from cdh23 ^aj64a/aj64a mutants, which lack functional tip links²⁹, and found that 97 ± 1.6% (n = 27) of hair cells took up 4-Di-2-ASP, but at a much reduced level (Fig. 3d, f; Supplementary Fig. 4b). Interestingly, this indicates mechanotransduction is uniformly dependent on Cdh23 but heterogeneously dependent on Tmc2b. We also asked whether posterior neuromasts with D–V orientations had similar or different percentages of hair cells with Tmc2b reliance as A–P-oriented neuromasts, and found that both classes of neuromasts were similarly impacted (Fig. 3e).

Second, the quality of mechanotransduction was roughly ascertained by determining the relative amount of 4-Di-2-ASP that could enter hair cells by measuring the fluorescence intensity of the brightest cell for each neuromast through live imaging. In tmc2b ^−/− animals, the cells eligible to take up the dye did so at a much reduced efficiency, ~fourfold less than controls, when the fluorescence signals were compared (Fig. 3f). This indicates a class of hair cell that is partially dependent on Tmc2b. The decreased dye uptake correlated with reduced microphonic potentials in tmc2b ^−/− mutants; all tmc2b ^−/− mutants (15/15) examined by both assays demonstrated reduced dye uptake and microphonic potentials, but all controls (10/10) had normal uptake and microphonic potentials.

Third, with high resolution, we examined whether mechanotransduction properties are uniform in functional hair cells or if there is a range of mechanotransduction function in neuromasts of tmc2b ^−/− animals. 4-Di-2-ASP presence was evaluated in all of the individual hair cells within a neuromast organ using quantitative confocal microscopy (Supplementary Fig. 4a). In tmc2b ^+/− fish, every L1 hair cell loaded the fluorophore, displaying a range of intensities that were all above background. In contrast, in a tmc2b ^−/− neuromast, most hair cells’ intensities were not above background, and the two cells that did transduce did so at a reduced level (Supplementary Fig. 4a). To inspect each neuromast for hair cells that took up the fluorophore, even at minuscule levels, we increased the optical power of the imaging by increasing the gain. For the same neuromast imaged in Supplementary Fig. 4a, we observed a several-fold increase in fluorescence levels of the two cells that did transduce (Supplementary Fig. 4b). Overall, these findings demonstrate that there is a subpopulation of hair cells that are entirely dependent on a single Tmc family member, which strikingly contrasts with the requirement in the mammalian ear, where there is redundancy between TMC1 and TMC2^16,18,19. Moreover, within the population of hair cells that still transduce, there is variability in transduction, yet all show some Tmc2b reliance, suggesting potential heterogeneity in the mechanotransduction channel, or its ancillary proteins, between hair cells within a single neuromast.

To determine if there was upregulation of tmc2b paralogs tmc1 or tmc2a, we assessed relative expression levels of cognate mRNAs in wild-type and tmc2b ^−/− mutant zebrafish. For each of these genes, expression in the mutants decreased slightly relative to controls as determined by RT-PCR experiments (Supplementary Fig. 5). This is an expected result that correlates with the decrease in hair cell numbers in the mutants because fewer hair cells would naturally lead to fewer total transcripts. These findings indicate that these genes are not upregulated in response to genetic inactivation of tmc2b.

Spatial position controls head neuromast mechanotransduction

Because the geometry of neuromast position is distinct between neuromasts of the head and those of the trunk and tail, complex vs. linear (Fig. 1a, top)^10,30, we examined whether patterns of mechanotransduction function varied in the ALL of tmc2b ^−/− animals at 6 dpf using 4-Di-2-ASP. MI1 neuromasts demonstrated a similar pattern of partial dependence as posterior neuromasts, with most hair cells absolutely requiring Tmc2b, and a smaller subset, 22%, still able to load fluorophore (Fig. 4a–c,f). These cells took up fluorophore at a reduced efficiency, ~fivefold less than controls (Fig. 4g). Similar patterns of fluorophore entry were observed for most neuromasts of the head (Fig. 4f, g). In sharp contrast, neuromast IO4 was minimally impacted by the tmc2b mutation (Fig. 4d–g). The number of hair cells that took up 4-Di-2-ASP was similar to control values and the fluorescence intensity was mildly reduced (normalized mean fluorescence intensity ratio of whole neuromasts from tmc2b ^−/− larvae is 0.72 ± 0.1 (n = 15)), indicating that hair cells of this neuromast are minimally reliant on, or independent of, Tmc2b for mechanotransduction (also see microphonic potential values for IO4 below). Additionally, the number of IO4 hair cells are the same in tmc2b ^−/− mutants and wild-type animals, showing that this gene is not necessary to sustain this assemblage of hair cells (Supplementary Fig. 3c).

Hair bundle orientation influences mechanotransduction

To further explore the molecular and cellular logic of heterogeneity in mechanotransduction of the PLL, we co-labeled tmc2b ^+/− and tmc2b ^−/− larvae with fluorophore-coupled phalloidin and FM1-43FX to reveal the potential relationship between hair bundle orientation and residual mechanotransduction channel function. In controls, hair cells of posterior neuromasts L1 and LII.1, with A–P- and D–V-facing hair bundles, respectively, took up FM1-43FX similarly (Fig. 5a, c, g, h). In agreement with 4-Di-2-ASP loading assays displayed in Fig. 4, only a subset of hair cells loaded with FM1-43FX in posterior neuromasts with either an A–P or a D–V orientation in tmc2b ^−/− larvae (Fig. 5b, d). Interestingly, we observed that the number of posterior-facing cells with entry of FM1-43FX was significantly higher than anterior-facing cells (Fig. 5g; Supplementary Table 1). A hair cell with posterior-facing hair bundles is defined as one in which the longest stereocilia are more proximal to the head than the shorter protrusions; these hair bundles transduce when deflected anteriorly. From these data, we conclude that in the PLL, hair bundle orientation is in lockstep with molecular components that control mechanotransduction apparatus function, offering the possibility that PCP formation during development modulates proteins that impact transducer action.

Tmc2b reliance is constant for each neuromast identity

The fluorescent molecule uptake experiments described above were performed on larvae at the same time point, 6-dpf. Therefore, one potential explanation for limited fluorescent molecule uptake is that the neuromast’s Tmc2b reliance is transitioning between levels of dependence, and we observed a time point in that shift. Alternately, the distribution of Tmc2b reliance may be constant. This distinction could be important, as the latter would imply that water flow is being detected by different sets of proteins that directly or indirectly influence transduction at the tips of stereocilia rather than a stepwise process for the developmental regulation of protein expression. To distinguish if this reliance is transient and developmentally related, we repeated the uptake assay on A–P oriented posterior neuromasts of older fish, 14 dpf. The percentage and orientation of hair cells that had partially active mechanotransduction channels were nearly the same as at 6 dpf, indicating that Tmc2b reliance is constant within this time period and not being modulated (Supplementary Fig. 6a; Supplementary Table 1). In addition, examination of D–V-oriented neuromasts in the tmc2b ^−/− mutant’s PLL revealed that hair cells with residual mechanotransduction (dye uptake) are predominantly those that face ventrally, and this dependence is preserved at 6- and 14-dpf (Fig. 5h; Supplementary Fig. 6b; Supplementary Table 1). These results indicate that during early larval stages, Tmc2b reliance is constant with respect to each neuromast position and the particular orientations of hair cells within.

Tmc2b dependence is non-uniform and asymmetric on head

We next tested whether the asymmetric molecular biases of the trunk and tail neuromasts were recapitulated in the head neuromasts that lie approximately on single planes or sheets on each of the lateral sides of the fish (Fig. 1a, Supplementary Fig. 1) of tmc2b ^−/− mutants. Co-labeling 6-dpf larvae with phalloidin and FM1-43FX demonstrated that hair cells that took up dye within each neuromast were strongly biased for a single direction in all but one neuromast (Fig. 5g, h; Supplementary Table 1). In agreement with 4-Di-2-ASP uptake assays performed on IO4 (Fig. 4d, e), FM1-43FX accumulated in all hair cells irrespective of hair bundle orientation (Fig. 5e, f, h), demonstrating that PCP was not a significant factor in Tmc2b regulation in this particular neuromast. The populations of hair cells that loaded FM1-43FX generally faced posteriorly for A–P-oriented neuromasts of the head (Fig. 5g; Supplementary Table 1). For MI1 and M2, which have different placodal origins and accordingly are innervated by different ganglionic afferents, the uptake pattern was opposite for MI1 (ventral facing) and M2 (dorsal facing) (Fig. 5h; Supplementary Fig. 7; Supplementary Table 1 ). This indicates that Tmc2b can be a dominant factor for either direction, dorsal or ventral. These patterns were maintained in 14-dpf zebrafish (Supplementary Fig. 6; Supplementary Table 1). Our studies indicate that each ALL mechanotransduction apparatus is tailored for each neuromast during development, based on the position in the body plan and orientation of the hair cells.

Overall, the molecular differences within the entire complement of neuromasts suggest that not all neuromasts are functionally equivalent. This agrees with a recent finding demonstrating that distinct neuromasts contribute differently to behaviors³¹. The exact parameters impacted by differential Tmc2b expression is an intriguing subject for further investigation.

Mechanosensory map of Tmc2b reliance

To visualize the complex anatomical and molecular associations of the lateral line as they relate to Tmc2b, we generated an integrated map of Tmc2b reliance (Fig. 6). Through this mechanosensory map, which displays neuromast position, hair cell orientation, and quality of mechanotransduction, a molecular logic for water flow detection begins to emerge. For all observed A–P-oriented neuromasts, partially functional mechanotransduction channels were predominantly from posterior-facing hair cells in tmc2b ^−/− mutants (Fig. 6). The breaks in these neuromasts’ symmetries indicate that detection of water flow from the front of the fish is heavily dependent on Tmc2b. But, encoding water flow coming from the rear relies on a different molecular mechanism that directly or indirectly involves Tmc2b and another factor.

The map (Fig. 6) also demonstrates that Tmc2b dependence varies with specific topological positions of neuromasts. M2 and MI1 had opposite residual sensitivities, indicating that Tmc2b can be a dominant factor for the detection of water flow coming from the top or bottom of the fish (Fig. 6, Supplementary Fig. 7). IO4 is nearly independent of Tmc2b (Fig. 6, Fig. 4d–g, Fig. 5e, f, Supplementary Fig. 2h). Loss of Tmc2b in IO4 does not affect microphonic potentials, which are the result of small cation entry, but entry of the bulky cationic fluorophore is mildly impacted (4-Di-2-ASP⁺ atomic mass = 394 u) (Fig. 6, Fig. 4g, Supplementary Fig. 2h). These findings indicate that IO4 hair cells are minimally dependent on Tmc2b, and this modest reliance is not impacted by hair bundle orientation; however, IO4 may depend on another yet unidentified protein.

Characterization of neuromasts above the eye demonstrated that these neuromasts, whose axes of best sensitivity are tangential to the perimeter of the eye, have variable fluorescent molecule uptake patterns. In the tmc2b ^−/− mutant, SO1 and SO3 are similar to IO4, with all hair cells able to take up fluorescent molecules similarly. However, SO2 has limited uptake from hair cells that mainly face a single direction. This surprising result indicates that despite being derived from the same placode, not all neuromasts have the same molecular components for mechanotransduction (Fig. 6). Similarly, IO1-4, all derived from a common placode, have variable Tmc2b reliance (Fig. 6). Overall, the map demonstrates that neuromasts employ different molecular mechanisms that are required for mechanotransduction to encode water motion. These mechanisms are based on neuromast topological position and hair bundle orientation.

Tmc2a and Tmc2b coordinate to permit mechanotransduction

We next asked what is the identity of the other element that may coordinate with Tmc2b to allow ion entry. To answer this question, we turned to a promising paralogous candidate gene, tmc2a, known to be expressed at low levels in the lateral line¹⁵. Tmc2a and Tmc2b have a 69% amino acid sequence similarity. To determine if tmc2a and tmc2b genetically interact to enable mechanotransduction, we successfully lesioned both of these genes, which reside on Chromosome 5, separated by ~39.9 centiMorgans. This was accomplished by the simultaneous injection of customized single-guide RNAs (sgRNAs) directed towards exon 6 of tmc2a and exon 7 of tmc2b along with Cas9 (CRISPR-associated protein 9) mRNA (Fig. 7). Fish with a 2-bp deletion in exon 6 of tmc2a and a 5-bp deletion in exon 7 of tmc2b, which lead to premature amber (UAG) and opal (UGA) stop codons upstream of regions that encode most transmembrane domains in the respective genes, were characterized. Mechanotransduction quality in double-knockout zebrafish homozygous (tmc2a ^−/− /tmc2b ^−/−) for the mutations and in controls were assessed using fluorescent molecule uptake and electrophysiological measurements. Stimulus-evoked microphonic potentials were not detectable in neuromasts with A–P orientations in the PLL of tmc2a ^−/− /tmc2b ^−/− larvae (Fig. 8a, b) at 6 dpf, yet robust signatures were apparent in controls. This finding was buttressed in an FM1-43FX uptake experiment, where no significant fluorescence was detected in posterior D–V-oriented LII.1 or head D–V-oriented IO4 neuromasts in tmc2a ^−/− /tmc2b ^−/− animals (Fig. 8c, d). Enumeration of hair cells in tmc2a ^−/− /tmc2b ^−/− mutant larvae revealed a small reduction (~25%) in the numbers of hair cells of posterior neuromast relative to controls (Supplementary Fig. 8). Overall, these findings indicate that the residual activity in the tmc2b ^−/− mutant is dependent on Tmc2a.

Hypothetical models to explain the relationship between neuromast topological position, hair bundle orientation, and Tmc2a and Tmc2b dependencies are displayed in Fig. 9 and Supplementary Fig. 9. These models show how varied Tmc2a and Tmc2b protein expression may give rise to the range of observed phenotypes. Figure 9 models the differences between L1 and IO4. Whereas, Fig. 9a and Supplementary Fig. 9 explain the subtle variations found in L1.

Taken together, our studies suggest that the basis of water motion detection is the coordination of specific mechanotransduction components with discrete neuromast positions and cellular orientations throughout the lateral line. First, we demonstrate that most ALL and PLL neuromasts of the larval zebrafish have maximum sensitivities along two axes, despite having a complicated arrangement of neuromasts on the head (Fig. 1, Supplementary Fig. 1). Second, there are three classes of hair cells with regard to Tmc2b reliance in the lateral line system: absolute, partial, and independent. This suggests that, along with the fact that hair cells with opposite bundle orientations are innervated by different afferents, there is an additional layer of regulation—at the molecular level—in mechanotransduction (Figs. 3, 4). Third, our findings demonstrate that most neuromasts are heavily dependent on Tmc2b, but IO4 is not, suggesting that the developmental factors that result in neuromast placement can govern mechanotransduction channel apparatus function (Fig. 4). Therefore, a developmental plan may predetermine distinct sets of mechanotransduction apparatus components for the neuromast to function properly at its specific topological position. Fourth, hair cells with partial reliance on Tmc2b for mechanotransduction within a single neuromast generally face one direction, indicating that the factors that regulate PCP may also modulate mechanotransduction apparatus composition (Fig. 5). This is functionally evident in the loss of the 2f response in stimulus-evoked microphonic potentials from posterior neuromasts as well as in fluorescent molecule uptake studies. Fifth, interestingly, not all neuromasts that are derived from the same placode demonstrate the same directional dependence on Tmc2b (Fig. 6), suggesting complex modes of Tmc2b regulation even within the same branch of the lateral line.

We speculate that the use of multiple different channel subunits or accessory proteins to change mechanotransduction channel quality may have important consequences for the fish’s ability to detect the direction of water flow. One axis of best sensitivity has hair cells that face anteriorly and posteriorly, but water flowing in either direction may require different responses than a system composed of perfectly symmetrical subunits could provide. The lateral line seems to resolve this conundrum by breaking neuromasts’ morphological mirror symmetries at the molecular level to generate functional asymmetries. Our data suggest that asymmetries are achieved by using at least two sets of components, which contain Tmc2b or both Tmc2a and Tmc2b, to directly or indirectly influence the mechanotransduction channel apparatus, allowing changes in mechanosensory properties for one direction vs. the other. The mechanotransduction parameters affected by differential Tmc2b dependence are an intriguing consideration for the future, as is whether Tmc2b is a member of the mechanotransduction channel itself or if it plays more of an accessory role in the neuromast.

The cochlea is known to have subtle changes in the properties of mechanotransduction channel conductance along the tonotopic gradient, which are considered important for frequency selectivity^32,33,34. Mice lacking functional TMC1 largely lose the tonotopic gradient in conductance of outer hair cells¹⁹. Our findings in the neuromast demonstrate that mechanotransduction channel properties can be modulated by a single mechanotransduction apparatus component, Tmc2b. And, its absence results in three different outcomes, no, partial, and normal transduction, depending on hair cell location and orientation. This suggests that the neuromast and the cochlea may have evolutionarily preserved mechanisms of mechanotransduction conductance modulation—the mixing and matching of mechanotransduction apparatus components. But the neuromast also has another class of hair cell, perhaps an archaic class, with a simpler requirement for mechanotransduction: absolute dependence on Tmc2b. This hair cell class in this respect is similar to Drosophila mechanosensitive neurons, which rely on just one TMC protein for mechanotransduction-based larval locomotion³⁵ and food texture sensation³⁶.

Methods

Zebrafish

Wild-type Tübingen (Tü), cdh23 ^aj64a/aj64a mutant²⁹, and Tg(pvalb3b:β-actin-mCherry) ³⁷ zebrafish were used in this study. The CWRU IACUC approved all animal use.

Molecular biology

PCR was used to amplify tmc2b cDNA from zebrafish macular cDNA (Superscript III, Invitrogen) using primers Tmc2b-TA-5′-Cterm (5′-GGGATGGAAATGGATGCAGAATTTGATAGT-3′) and Tmc2b-TA-3′-Cterm (5′-ATCTTCGTGGTGGTGGTCCTCC-3′). The amplicon was ligated into pCR4 TOPO vector (Invitrogen) and digested with AgeI and PmeI before being shuttled, with T4 DNA ligase (Promega), into pMT/pV3b/mGFP vector³⁸, which had been digested with XmaI and PmeI, resulting in pMT/pV3b/tmc2b-mGFP.

Imaging Tmc2b localization

Tg(pvalb3b:β-actin-mCherry) zebrafish injected with DNA plasmid, pMT/pV3b/tmc2b-mGFP, at 150 ng/μl and Tol2 RNA at 25 ng/μl³⁹ were anesthetized in 650 mM 3-aminobenzoic acid ethyl ester methanesulfonate (Sigma-Aldrich) in fish water at 4 dpf. Immobilized fish were imaged by confocal microscopy with a 40× objective (Leica SP8; Leica). 488 nm and 594 nm wavelengths were used to excite mGFP and mCherry, respectively.

Image analyses

Enumeration of the numbers of hair cells that loaded with 4-Di-2-ASP were determined by individual hair cell fluorescent signal on a confocal microscope and compared to the total number of hair cells as determined from bright-field imaging (Fig. 3d, e, Fig. 4f, and Fig. 6).

For the PLL (Fig. 3f), raw images gathered using a microscope (Olympus BX51WI) were subjected to fluorescence measurements using ImageJ. A ROI, a circle 1 μm in diameter, was used to obtain measurements from the brightest cell in each neuromast (I _cell) and an area without cells (I _background) in the same image. Fluorescence intensity of 4-Di-2-ASP-loading (I _loading) for each neuromast was normalized using the formula, I _loading = I _cell−I _background. To avoid pixel saturation, raw images were taken with three different filter sets that allowed different amounts of light to enter each sample. The relation between incoming light and emitted fluorescence recorded by camera was approximately linear; therefore, I _loading measured from images taken with a darker filter (6% of light passes), lighter filter (25% of light passes), and no filter (100% of light passes) were corrected by dividing each measurement by a factor of 1, 4.17, or 16.7, respectively, to obtain I _corrected ^40,41. Values of I _corrected were plotted and subjected to statistical tests.

For the ALL (Fig. 4g), the mean intensities of 4-Di-2-ASP fluorescence were measured from the maximum projection of a z-stack image that was collected on a confocal microscope (Leica TCS SP8, Leica) for each neuromast. ROIs were drawn for individual neuromast manually, and the outline of the whole neuromast was defined by the bright-field image taken simultaneously with a fluorescent channel. Intensity values are using a 0–225 range scale set by Leica software without further processing, and no neuromast images with saturated pixels were used in fluorescence quantitation. I _whole = I _neuromast–I _background.

To quantitate 4-Di-2-ASP fluorescence intensity profiles for multiple individual hair cells within a single neuromast (Supplementary Fig. 4 and Fig. 4c, e), the basolateral region for each hair cell soma was used. The outlines of individual hair cells (ROI) were defined manually by the bright-field image taken simultaneously with the confocal z-stack fluorescent image series (Leica TCS SP8, Leica). The 4-Di-2-ASP fluorescence intensity of each hair cell was the mean pixel value measured within each ROI after subtracting the background pixel intensity value from a 6 × 6 μm region devoid of hair cells (LAS X; Leica).

For Fig. 5g, h and Supplementary Fig. 6, FM1-43FX loading %=(number of A-, P-, D-, or V-facing hair cells that loaded/total number of hair cells with the same orientation)×100.

Mutation of the tmc2b locus with TALENs

To identify a suitable region of exon 4 of the tmc2b gene (ENSDARG00000030311) for targeting by transcription activator-like (TAL) effector nucleases (TALENs), the ZiFiT Targeter computational program^42,43 was used. Specificity of the tmc2b TALEN target site was confirmed by the NCBI BLAST tool using the zebrafish genomic DNA database (https://blast.ncbi.nlm.nih.gov/Blast.cgi). TALEN expression vectors were constructed with the fast ligation-based automatable solid-phase high-throughput (FLASH) assembly protocol^25,44,45. Biotinylated α units, extension units, termination units, and the expression vectors used in this study are listed in order for tmc2b-Left-TALEN (TAL373, TAL145-TAL72-TAL203, TAL274, JDS70) and tmc2b-Right-TALEN (TAL374, TAL10-TAL32-TAL140, TAL302, JDS71). Designed vectors were confirmed by DNA sequencing using primers oSQT1 (5′-AGTAACAGCGGTAGAGGCAG-3′), oSQT3 (5′-ATTGGGCTACGATGGACTCC-3′), and oJS2980 (5′-TTAATTCAATATATTCATGAGGCAC-3′). Sequence-confirmed expression vectors carrying either tmc2b-Left-TALEN or tmc2b-Right-TALEN were linearized with PmeI and were used as templates for in vitro transcription reactions (Ambion mMessage mMACHINE Ultra T7 kit; Life Technologies Inc.). RNAs were purified by lithium chloride precipitation, and the quality and quantity of each was verified (Agilent 2100 Bioanalyzer, Agilent Technologies; Nanodrop 2000, Thermo Scientific). RNAs that encode each half site of tmc2b-TALEN at 150 ng per μl were mixed with phenol red and 1× Danieau’s solution (58 mM NaCl, 0.7 mM KCl, 0.4 mM MgSO₄, 0.6 mM Ca(NO₃)₂, and 5.0 mM HEPES pH 7.6) and then coinjected into zebrafish embryos at the one-cell stage to create gene-specific mutations. Embryos injected with tmc2b-TALEN RNAs were collected individually and subjected to mutation detection at the tmc2b TALEN target site using high resolution melting analysis (HRMA)⁴⁶ with primers tmc2b HRM F (5′-GACGACGAGTCTATGTCTGAAGG-3′) and tmc2b HRM R (5′-CAGGTTCACAGCACAAACAGCAAG-3′). The mutations in tmc2b carried by founder fish and F1 fish were identified using sequencing primers tmc2b SEQ F (5′- CAGCATTCGATGAAATTCTTCATG-3′) and tmc2b SEQ R (5′- GAAGGGTAAGGGCAATGCAGTA-3′). tmc2b TALEN mutant fish, with a 7-bp deletion (Fig. 2e), used for assays in this work, were genotyped by two sets of primers, tmc2b WT SEQ F (5′-CAGCATTCGATGAAATTCTTCATG-3′) and tmc2b WT SEQ R (5′- GAGTCTCCTCTTCATGCGCCAG-3′) for the wild-type allele and tmc2b mut SEQ F (5′-CTTAGGAACAAACCCTGAAGA-3′) and tmc2b mut SEQ R (5′ - CTACCCTGTAAAGGCAATC-3′) for the mutant allele.

Measurement of neuromast microphonic potentials

Zebrafish larvae were anesthetized using ethyl 3-aminobenzoate methanesulfonic acid (Sigma-Aldrich) dissolved in a standard bath solution (120 mM NaCl, 2 mM KCl, 10 mM HEPES, 2 mM CaCl₂, 0.7 mM NaH₂PO₄ and adjusted to pH 7.3). The larvae were secured in a recording chamber using dental floss tie downs⁴⁷ and placed under the microscope for observation. We visually monitored blood flow and heart rate to assess the viability of each zebrafish larva. Zebrafish were visualized on an upright microscope (BX51WI; Olympus) equipped with a swing nosepiece and 4 × 0.1 and 100 × 1 NA objectives. We recorded microphonic potentials using a PC-505B amplifier (Warner Instruments) and a PCI-6221 digitizer (National Instruments). Images were observed with a Grasshopper3 CMOS camera (Point Grey) and captured with manufacturer provided software. We recorded from neuromasts of larvae ranging from 5 to 6 dpf. Kinocilia tufts were deflected with a fluid jet^23,48 delivered via a glass pipette with a diameter of ∼7 µm and controlled by HSPC-1 (ALA Scientific Instruments). The fluid jet pipette was placed ∼50 µm away from the neuromast and used to deliver sinusoidal stimuli of 50 Hz, generated by jClamp software (Scisoft, courtesy of Joseph Santos-Sacchi, Yale University, New Haven, CT, USA). The microphonic potentials were recorded at room temperature (~22 °C). We used a borosilicate glass pipette with a resistance of 3–6 MΩ when filled with standard bath solution and placed near the apical edges of the lateral line neuromasts. Placement of fluid jet and recording pipettes were controlled by a set of micromanipulators (MPC-325; Sutter Instrument). Microphonic potentials were recorded with a jClamp in current-clamp mode, amplified by 20 (SIM983, Stanford Research), and low-pass filtered at 200 Hz. All records represent an average of at least 500 trials.

Measurement of inner ear microphonic potentials

Recording of microphonic potentials from the inner ear in response to vibratory stimuli was conducted on 8–9 dpf larvae. Anesthetized larvae were adhered to the recording chamber by low melting point agarose (IBI Scientific, Peosta, IA). The recording pipette (prepared as described above), loaded with the standard bath solution, was inserted into the otic capsule to reside in close proximity to both maculae. The vibrations were delivered via a glass stylus with a tip diameter of ∼7 µm using a piezo actuator (PA 4/12, Piezosystem Jena) driven by a power amplifier (ENV 800, Piezosystem Jena). The command 200 Hz sinusoidal stimulus was generated by jClamp. Microphonic potentials were recorded as described above and low pass filtered at 1000 Hz.

Scanning electron microscopy

Zebrafish larvae (5–6 dpf) were fixed in 2.5% glutaraldehyde containing 0.1 M cacodylate buffer (EMS, Hatfield, PA) supplemented with 2 mM CaCl₂ for 2 h at room temperature. Then, the samples were dehydrated in a graded series of ethanol, critical-point dried with liquid CO₂ (CPD 030, BAL-TEC AG, Balzers), and sputter-coated with ~10.0 nm palladium (Desk IV, Denton Vacuum). Samples were imaged with a field-emission SEM (Helios Nanolab 650, FEI).

Real-time polymerase chain reaction

Total RNA from four 6-dpf wild-type or tmc2b ^−/− mutant zebrafish were separately extracted (RNeasy Mini Kit; Qiagen). 400 ng total RNA from each sample was reverse transcribed to produce cDNA (SuperScript III First Strand Synthesis System for RT-PCR Kit; Invitrogen Life Technologies). The relative quantity of each mRNA was obtained with the comparative threshold cycle (C _t) method. PCR amplification was performed with a 20 μl reaction solution (10 ul of 2× RT² SYBR Green ROX Fast Mastermix (Qiagen), 0.4 μl of forward primer (10 μM), 0.4 μl of reverse primer (10 μM), 5.2 μl nuclease-free water, and 4 μl of diluted template cDNA (400 ng/μl)) in a PCR machine (StepOnePlus real-time PCR system; Applied Biosystems). All reactions were performed in quadruplicate. The PCR conditions were as follows: 95 °C for 10 min then forty amplification cycles (95 °C for 15 s, 60 °C for 1 min). As an internal control, primers for beta-actin were used to amplify in parallel reactions. In all cases, a reverse transcriptase negative control was included. Primer sequences: b-actin RT-PCR F2, 5′-GACCCAGACATCAGGGAGTGATGG-3′; b-actin RT-PCR R2, 5′-AGGTGTGATGCCAGATCTTCTCCAGT-3′; Tmc1 RT-PCR F3, 5′-CAGGTCTCAGAAGTTTGCACTG-3′; Tmc1 RT-PCR R3, 5′- GATCACATCGAACAGCATCG-3′; Tmc2a RT-PCR F3, 5′-ATGGCATGAACTTGGTCCTC-3′; and TMC2a RT-PCR R3, 5′-CAGTTTTCCGAGGAATGGAC-3′. Relative amounts of each mRNA were determined by the C _t strategy, with the mean wild-type control value set at 1.0. C _t values were obtained for each gene mRNA, with C _t defined as the threshold cycle of the PCR at which the amplified product was first detected. ΔC _t is the difference in C _t values between targets and the β-actin internal control. ΔΔC _t represents the difference between ΔC_ts from wild-type and tmc2b ^−/− zebrafish. The fold difference in a target's expression between wild-type and tmc2b ^−/− mutant zebrafish was calculated as 2^−ΔΔCt.

Mutation of tmc2a and tmc2b with CRISPR/Cas9

Because tmc2a (ENSDARG00000033104) and tmc2b (ENSDARG00000030311) both reside on chromosome 5, generation of double knockout zebrafish was achieved by co-injection of one-cell stage zebrafish embryos with Cas9 RNA and two sgRNAs that target each of the respective genes. CRISPR target sites were selected using the CHOPCHOP web tool (http://chopchop.cbu.uib.no). sgRNAs were synthesized using T7 RNA polymerase (Ambion MEGAscript)⁴⁹. tmc2a was targeted with an sgRNA complementary to exon 6 (5′-CCTCATGGTGGGCATTGGGACCT-3′). tmc2b was targeted with an sgRNA complementary to exon 7 (5′-CCAGAGAGGAGCAGGATTCGGCC-3′). Successfully targeted loci were identified by HRMA and DNA sequencing. HRMA primers: tmc2a CRISPR HRMA Forward (5′-CAGGGTTACTGCAAGTACTCAG-3′), tmc2a CRISPR HRMA Reverse (5′-GCAATGAATGAAAGAGGGACTC-3′), tmc2b CRISPR HRMA Forward (5′-TGGGCATCCCTTATGGCTC-3′), and tmc2b CRISPR HRMA Reverse (5′-ATGAGGGGAAACTTACAGCTTG-3′). Sequencing primer pairs for genotyping: tmc2a CRISPR seq Forward (5′-GAGACCGGAAATCTCGTGCC-3′), tmc2a CRISPR seq Reverse (5′-CCTTTTGTCACTGAGGGACAAC-3′), tmc2b CRISPR seq Froward (5′-CCTTGTTCTCTTCGGCTTCATG-3′), and tmc2b CRISPR seq Reverse (5′-TGACAGAGGTAACCGGAACTGC-3′).

4-Di-2-ASP uptake assay

At 6 or 14 dpf, zebrafish were bathed with 20 μM 4-Di-2-ASP for 30 s and washed three times at 5 min intervals with rinsing solution (0.612 mM of Ethyl 3-aminobenzoate methanesulfonic acid and 0.01 mg/ml BSA in fish water) prior to imaging. In more detail, for fluorescent molecule uptake, the fish were put into baskets and bathed with labeling solution with manual agitation that resulted in fish movements forward and backward, up and down, and side to side, randomly for 30 s. Data was collected with an upright microscope (BX51WI; Olympus) equipped with a 100× objective with a NA of 1 or a confocal microscope with a 40× objective with a NA of 1.3. On the confocal microscope, a white light laser was used for excitation at 470 nm, and fluorescent signals were collected between wavelengths of 550–600 nm. For comparisons with different gains, laser settings under both low and high conditions were kept the same between genotypes.

FM1-43FX uptake assays

Zebrafish at 6 or 14 dpf were incubated with 5 μM FM1-43FX (Invitrogen), a fixable analog of FM1-43, for 30 s, then washed three times at 5 min intervals with the rinsing solution before overnight fixation with 4% paraformaldehyde (PFA) at 4 °C. More specifically, for FM1-43FX uptake, the fish were put into baskets and bathed with labeling solution with manual agitation that resulted in fish movements forward and backward, up and down, and side to side, randomly for 30 s. Permeabilization was performed with 3% Triton-X100 (Sigma) overnight at room temperature. Blocking with 5% goat serum was carried out for 6 h at room temperature. Following this, samples were rinsed with phosphate-buffered saline (PBS) four times for 5 min. To visualize hair cells, parvalbumin 3 antiserum at 1:600 dilution was used as a primary antibody and Alexa Fluor 633 goat-anti-rabbit IgG (Invitrogen) at 1:200 dilution was used as the secondary antibody. To identify hair-bundle orientation, Alexa Fluor 633 phalloidin at 1:50 dilution was used to label actin filaments. Processed zebrafish samples were mounted and kept in antifade mounting medium (VECTASHIELD; Vector Laboratories). Images were acquired on a confocal microscope with excitation wavelengths of 488 nm for FM1-43FX and 633 nm for Alexa Fluor 633.

Statistics and software

All statistical analyses were performed using GraphPad Prism 7. Data are reported as mean ± SEM. Comparisons between groups were tested by Mann–Whitney test, Student’s t-Test, Kruskal–Wallis test with Dunn’s multiple comparison test, or ANOVA with Holm-Sidak post hoc testing depending on data distribution. All sample n values were greater than 5, unless noted. Topographical representations of proteins were generated with software (TMHMM 2.0; http://www.cbs.dtu.dk/~krogh/TMHMM/).

Data availability

Data supporting the findings of this study are available upon request from the corresponding author.

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Acknowledgements

We are grateful to D. Wang and K. Abbasi for assistance with electron microscopy, A.J. Hudspeth for parvalbumin 3 antiserum, and T. Nicolson for the cdh23 mutant strain. We also thank L. Zon, A. Wynshaw-Boris, and the members of our laboratory who provided discussion and comments on this manuscript. We acknowledge use of the Leica SP8 confocal microscope in the Light Microscopy Imaging Facility at Case Western Reserve University made available through the Office of Research Infrastructure (NIH-ORIP) Shared Instrumentation Grant S10OD016164. This research was supported by NIH grants DC009437 (B.M.M.), DC015016 (R.S.), and the Center for Clinical Research and Technology at University Hospitals Cleveland Medical Center (B.M.M.).

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Shih-Wei Chou and Zongwei Chen contributed equally to this work.

Authors and Affiliations

Department of Otolaryngology–Head and Neck Surgery, Case Western Reserve University School of Medicine, Cleveland, OH, 44106, USA
Shih-Wei Chou, Zongwei Chen, Shaoyuan Zhu, Robin W. Davis, Jiaqi Hu, Li Liu, Carol A. Fernando, Kayla Kindig, William C. Brown, Ruben Stepanyan & Brian M. McDermott Jr.
Department of Biology, Case Western Reserve University, Cleveland, OH, 44106, USA
Shih-Wei Chou, Zongwei Chen, Shaoyuan Zhu, Robin W. Davis, Jiaqi Hu, Li Liu, Kayla Kindig & Brian M. McDermott Jr.
Department of Neurosciences, Case Western Reserve University School of Medicine, Cleveland, OH, 44106, USA
Ruben Stepanyan & Brian M. McDermott Jr.
Department of Genetics and Genome Sciences, Case Western Reserve University School of Medicine, Cleveland, OH, 44106, USA
Brian M. McDermott Jr.

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Shih-Wei Chou
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Contributions

Gene editing was carried out by S.-W.C., J.H., C.A.F., L.L. and R.W.D. Imaging was performed by S.-W.C., Z.C. and S.Z. Electrophysiology experiments were implemented by R.W.D., S.-W.C, K.K. and W.C.B. Experimental plans were developed by B.M.M., R.S., S.-W.C. and Z.C. Paper was written by B.M.M. S.Z. and R.W.D. contributed equally to this work. All authors commented and contributed to the final version of the manuscript.

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Correspondence to Brian M. McDermott Jr..

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Chou, SW., Chen, Z., Zhu, S. et al. A molecular basis for water motion detection by the mechanosensory lateral line of zebrafish. Nat Commun 8, 2234 (2017). https://doi.org/10.1038/s41467-017-01604-2

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Received: 27 January 2017
Accepted: 29 September 2017
Published: 21 December 2017
DOI: https://doi.org/10.1038/s41467-017-01604-2

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