Dysfunctions of hearing and balance are often irreversible in mammals owing to the inability of cells in the inner ear to proliferate and replace lost sensory receptors. To determine the molecular basis of this deficiency we have investigated the dynamics of growth and cellular proliferation in a murine vestibular organ, the utricle. Based on this analysis, we have created a theoretical model that captures the key features of the organ’s morphogenesis. Our experimental data and model demonstrate that an elastic force opposes growth of the utricular sensory epithelium during development, confines cellular proliferation to the organ’s periphery, and eventually arrests its growth. We find that an increase in cellular density and the subsequent degradation of the transcriptional cofactor Yap underlie this process. A reduction in mechanical constraints results in accumulation and nuclear translocation of Yap, which triggers proliferation and restores the utricle’s growth; interfering with Yap’s activity reverses this effect.https://doi.org/10.7554/eLife.25681.001
The sensory organs of the inner ear arise from patches of Sox2-positive cells specified in the prosensory domain of the otic vesicle (Kiernan et al., 2005; Hartman et al., 2010). Despite differences in function, the organs for hearing and balance have similar structures and are populated by the same type of mechanosensory receptor, the hair cell. Hair cells are intercalated with supporting cells that are necessary for the proper sensory functions (Haddon et al., 1999). Additionally, when hair cells are lost in nonmammalian species, supporting cells can proliferate and transdifferentiate into new sensory receptors, allowing for recovery of hearing and balance (Corwin and Cotanche, 1988; Ryals and Rubel, 1988; Harris et al., 2003; Taylor and Forge, 2005). Although supporting cells in the vestibular sensory organs of adult mammals retain a limited ability to regenerate hair cells through direct conversion (Ruben, 1967; Forge et al., 1993; Rubel et al., 1995; Kawamoto et al., 2009; Lin et al., 2011; Golub et al., 2012), they lose the ability to reenter the cell cycle after neonatal stages (Golub et al., 2012; Burns et al., 2012a; Wang et al., 2015; but see Warchol et al., 1993). This deficiency may be a primary reason why hearing and balance function fail to recover in mammals after hair cell damage.
In order to understand what blocks the proliferation of supporting cells during regeneration, it is important to uncover the mechanism that arrests cellular proliferation in the developing sensory epithelia of the inner ear. The signaling pathways that control the proliferation of supporting cells during development have been studied extensively. For example, proliferation is blocked by downregulation of Wnt signaling in the developing murine inner ear (Jacques et al., 2012; Chai et al., 2012). ErbB signaling is also involved, for EGF and heregulin enhance supporting-cell proliferation in vitro and inhibition of the EGFR pathway arrests mitotic activity in sensory epithelia (Zheng et al., 1999; Hume et al., 2003; Doetzlhofer et al., 2004; White et al., 2012). We have recently demonstrated that SoxC transcription factors play an important role and that reactivation of their expression in the adult utricle elicits supporting-cell proliferation and the production of hair cells (Gnedeva and Hudspeth, 2015). Maturational changes in cell-cell and cell-matrix adhesion have also been shown to correlate with the ability of supporting cells to proliferate and transdifferentiate into hair cells in the utricle (Davies et al., 2007; Burns et al., 2008; Collado et al., 2011). Finally, although it is not clear what triggers their expression, the upregulation of retinoblastoma protein and of cyclin-dependent kinase inhibitors such as p27Kip1 enforces a nonproliferative state in postmitotic supporting cells (Chen and Segil, 1999; Löwenheim et al., 1999; Yu et al., 2010). However, neither conditional ablation of p27Kip1 nor forced re-expression of cyclin D in adult sensory epithelia is sufficient to relieve the block on supporting-cell proliferation (Laine et al., 2010; Loponen et al., 2011). These observations suggest the existence of parallel repressive mechanisms.
In this work, we used a vestibular sensory organ—the utricle—as a model system to study the cell-cycle exit in the sensory epithelia of the inner ear. By combining theoretical and experimental approaches to recreate the known features of utricular organogenesis, we elucidated a previously unrecognized mechanism underlying the arrest of the organ’s growth.
To assess the dynamics of the sensory epithelial growth during development in the murine utricle, we monitored the organ's development from embryonic day 15.5 (E15.5) through postnatal day 14 (P14). We observed a rapid expansion of the utricle’s sensory area—the macula—as determined by Sox2 labeling, between E15.5 and E18.5 (Figure 1A,D). After E19.5 the rate of macular growth declined until the organ reached its final size at P2. As was shown previously (Burns et al., 2012b), the rate of areal growth as a function of time fits the von Bertalanffy growth equation in which is a rate constant, is the final area, and is the area at time . The solution to this equation is , in which is an integration constant (Figure 1D). This equality indicates that the growth rate decreases continuously from the outset to zero when the tissue has reached its target size . Although this relation implies that the rate is negatively regulated by an increase in the organ’s size, it offers no information about the underlying mechanism. We therefore examined several models for self-regulation of the utricle’s sensory epithelia growth, focusing on the two that proved most effective.
To reduce the mathematical complexity of the system we introduced a two-dimensional model that captures the main qualitative features of the developing utricle (Figure 1—figure supplement 1). In this representation, the sensory epithelium of the utricle is surrounded by an elastic boundary that comprises all the nonsensory tissues encompassing the macula. We assume that the boundary is a linear elastic material that can be described by its Young’s modulus, or elastic modulus. This material property is defined as the ratio of the stress, or force per unit area, to the strain, the ratio of the deformation over the initial length, and indicates how much the material resists deformations imposed by an external force. We then used a lattice-based approach to create a stochastic computational simulation of macular growth (Swat et al., 2012).
In our first representation—the elasticity-limited model—we employed a value of the elastic modulus E such that the tissue is restricted to its experimentally observed size solely by the force exerted by the elastic boundary (Figure 1B,E; Video 1; Figure 1—source data 1; Source code 1; Table 1). In this model, the growth of the utricle ceases when the pressure generated by stretching of the elastic boundary equals that created by the growth and division of supporting cells. In this case the final size of the utricle is inversely related to the square of the elastic modulus of the elastic boundary (Figure 1F; Figure 1—source data 2).
In our second representation—the morphogen-limited model—we retained a weaker elastic boundary and assumed that the supporting cells of the utricle secrete a growth-inhibiting molecule (Figure 1C,E; Video 2; Figure 1—source data 3; Source code 2; Table 1). We assume that the morphogen diffuses slowly from its cellular source. Once the concentration of the growth inhibitor reaches a certain threshold value at the location of a given cell, this cell stops proliferating. The delay between supporting-cell formation and production of the inhibitor produces a gradient with the highest concentration at the center of the utricular macula and a lower concentration at the periphery. Cells at the center therefore stop proliferating first and a front of cell-cycle arrest (white line in Figure 1C) expands until it reaches the edge of the sensory epithelium and fully stops its growth. In this case, the final size of the sensory epithelium largely depends on the secretion rate and diffusion coefficient of the morphogen and in its absence the tissue would grow to a much greater size than is observed experimentally (Figure 1G; Figure 1—source data 4).
Although the elasticity-limited model represents a better fit to the data, both models can explain the developmental growth curve of the utricle (Figure 1E; Figure 1—source datas 3,5). We therefore experimentally tested additional predictions of the two models to evaluate their validity.
To distinguish between the proposed models, we developed an ex vivo culture system that allows manipulation of the stiffness of the extracellular matrix surrounding the utricle while monitoring the organ's growth. For these experiments we used Atoh-nGFP transgenic mice (Lumpkin et al., 2003) in which the hair cells are labeled by GFP expression, to visualize the sensory epithelium. We dissected part of the vestibular apparatus—the utricle, two ampullae, and portions of the semicircular canals—at E17.5 and maintained it as a free-floating culture for 3 hr. Healing of the cuts introduced during dissection yielded a closed organotypic structure that we term a utricular bubble (Figure 2A). Such a culture reduces the leakage of any secreted molecules from the system, allowing us to test the predictions of the morphogen-limited model.
To change the force opposing the sensory epithelia growth, we embedded the utricular bubble in a collagen gel of calibrated stiffness (Figure 2B). In our mathematical model this manipulation equates to changing the stiffness of the elastic boundary surrounding the macula. Because the developing inner ear is normally encapsulated by cartilage, we increased the elastic modulus of the gels by cross-linking the collagen fibers with addition of chondrocytes (Figure 2—figure supplement 1). A gel with an elastic modulus of 640 ± 3 Pa, corresponding to the stiffness of supporting cells in the inner ear (Sugawara et al., 2004; Zetes et al., 2012), was used to approximate the stiffness of the elastic boundary that the utricle experiences in vivo. At the opposite extreme, a 40 ± 6 Pa gel with no chondrocytes created significantly less force on the growing utricle.
Although both our models predicted that reducing the stiffness of the elastic boundary would allow the sensory epithelium to expand to a larger size, the expected spatial distribution of proliferating cells differed between the models. Because the concentration of growth-inhibiting morphogen should not be affected by external force, the morphogen-limited model predicted that the cells at the center of the utricle would not proliferate, regardless of a decrease in the gel's stiffness. On the contrary, if cell growth is suppressed only by elastic force, as proposed by the elasticity-limited model, supporting-cell proliferation should occur even near the center of the utricle.
We observed that in a 640 Pa gel, the E17.5 utricle grew at a rate similar to that found during normal development, reaching the size of P2 utricle by 5 d in culture (Figure 2C,D; Figure 2—source data 1). In contrast, over the same period a substantial expansion of a utricle’s macular area occurred in a 40 Pa collagen gel, where the organ reached 180% of its normal adult size (Figure 2C,D; Figure 2—source data 1). Quantification of supporting-cell proliferation by EdU incorporation over 4 d in culture revealed a significant tenfold increase in 40 Pa gels as compared to 640 Pa gels (Figure 2E; Figure 2—source data 2). In agreement with the elasticity-limited model, we observed that whereas the proliferation was limited to the utricle’s periphery in 640 Pa gels, supporting cells throughout the sensory epithelium reentered the cell cycle in 40 Pa gels (Figure 2F).
To ensure that the presence of chondrocytes did not inhibit supporting-cell proliferation, we cultured utricles in 40 Pa gels with chondrocytes situated locally at one edge of the organ. When after 24 hr we assessed the number of EdU-positive cells, we found no effect of the cartilage on the proliferation of supporting cells (Figure 2—figure supplement 2).
Simulations of the elasticity-limited model suggest that elastic force arrests the growth of the utricle, but leaves the organ in an unstable state; if cells retain their capacity to proliferate, random fluctuations could evoke additional growth. Because this behavior is not observed in vivo, elastic force likely triggers a secondary mechanism that stops proliferation when the organ reaches its target size. Hippo signaling, which is known to arrest cellular proliferation in response to mechanical force (Aragona et al., 2013; Dupont et al., 2011; Robinson and Moberg, 2011), might provide such a mechanism. Activation of Hippo signaling results in phosphorylation and degradation of the transcriptional cofactor Yap, restricting it from the nucleus and curtailing cellular proliferation. To investigate the potential role of this pathway in the utricle’s growth control, we assessed the expression of Yap, Hippo signaling kinases, and downstream target genes during the utricle's development.
The growth of the murine utricle and supporting-cell proliferation decline dramatically within the first two days of postnatal life (Figure 1A,D; Burns et al., 2012b; Gnedeva and Hudspeth, 2015). During the following week, the organ matures and the supporting cells within it lose the ability to reenter the cell cycle after damage (Kawamoto et al., 2009; Golub et al., 2012; Burns et al., 2012a; Wang et al., 2015). Our RNA-sequencing data (Gnedeva and Hudspeth, 2015; GSE72293) indicate that the expression of genes encoding the key components of the pathway—Mst1 and Mst2, Lats1 and Lats2, Yap, and Taz—was similar in actively growing utricles at E17.5 and growth-arrested mature utricles at P9 (Table 2). However, consonant with the activation of Hippo signaling, the level of Yap protein decreased significantly from E17.5 to P14 (Figure 3A). The expression of genes encoding the downstream targets of nuclear Yap, such as Ankrd1, Ctgf, and Cyr61 (Aragona et al., 2013), was accordingly downregulated significantly by P2 (Figure 3B; Figure 3—source data 1). In addition, the expression of genes encoding known inhibitors of nuclear Yap translocation, such as E-cadherin, α-catenin, and gelsolin (Aragona et al., 2013; Robinson and Moberg, 2011), was upregulated postnatally as the organ matured and lost its capacity for proliferative regeneration (Figure 3C; Figure 3—source data 2).
To investigate the relationship between the subnuclear localization of Yap and supporting-cell proliferation in the developing utricle, we analyzed the inner ears of E17.5 Atoh-nGFP mice. A single EdU injection 4 hr prior to analysis was used to label proliferating supporting cells. We demonstrated cytoplasmic labeling of Yap in postmitotic supporting cells near the center of the utricular sensory epithelium (Figure 3D,E; Figure 3—source data 3). The proliferating cells at the organ’s periphery showed significantly stronger Yap labeling, indicating some nuclear translocation of the protein. Yap was not present in hair cells.
The supporting cells in the utricle of a neonatal mouse retains a limited capacity to reenter the cell cycle (Burns et al., 2012a; Wang et al., 2015). To assess the potential role of Yap during this process, we examined the protein's expression in P4 utricles after injury in vitro. Utricles were dissected and allowed to attach to the bottom of a Petri dish coated with Cell-Tack adhesive. Subsequently, a linear cut was made with a 30-gauge needle along one border of the sensory epithelium, and each utricle was allowed to recover for 48 hr in culture medium containing EdU (Figure 4A). As expected (Davies et al., 2007; Meyers and Corwin, 2007), Sox2-positive supporting cells reentered the cell cycle at the site of injury (Figure 4B,C). Accordingly, as demonstrated by significant increase in the intensity of Yap antibody labeling (Figure 4C; Figure 4—source data 1 and 2), we observed the accumulation of Yap protein in the proliferating sensory epithelium, where Sox2-positive supporting cells translocated Yap to the nuclei (Figure 4B).
We next sought to determine whether the ability of supporting cells to reenter the cell cycle after injury depends on nuclear Yap signaling. In most contexts, Yap must bind Tead transcription factors to stimulate cellular proliferation (Vassilev et al., 2001). We therefore overexpressed GFP fused to a Yap-Tead interfering peptide, YTIP (von Gise et al., 2012) in ex vivo cultures. By using the Ad-Easy cloning system (Chartier et al., 1996; He et al., 1998), we developed a serotype five adenoviral overexpression vector carrying GFP-YTIP. We tested the transfection efficiency and toxicity of the virus on E16.5 utricle explants in vitro and found no difference with respect to a control virus expressing only GFP (Figure 4—figure supplement 1). The number of proliferating cells in the sensory epithelia of the utricles infected with GFP-YTIP was greatly reduced, reaching only 7% that in GFP controls (Figure 4—figure supplement 1).
We tested the effect of GFP-YTIP overexpression in the injury assay. After the utricles had been dissected and affixed to the bottom of a Petri dish, the cultures were infected with control GFP virus or with GFP-YTIP virus and left for 24 hr to allow the accumulation of GFP and GFP-YTIP proteins (Figure 4D). On the following day, the injury assay was performed and each utricle was allowed to recover for 48 hr in culture medium containing EdU. As determined by Sox2 and EdU labeling, the number of GFP-negative proliferating supporting cells at the injury site in utricles infected with GFP-YTIP virus was not significantly different from that in control GFP cultures. However, the number of GFP-positive cells within the proliferating supporting-cell population was reduced significantly in GFP-YTIP cultures as compared to GFP controls (Figure 4E,F; Figure 4—source data 3).
The pattern of Yap expression in the utricle during development and after injury suggested that the protein regulates supporting-cell proliferation. To test whether nuclear Yap translocation is controlled by elastic force we assessed protein expression in collagen gels of varying stiffness. The elasticity-limited model predicts that reducing the stiffness of the elastic boundary would allow cells in the center of the organ to expand in volume and therefore to reenter the cell cycle. A significant increase in the area of the utricular macula was observed in 40 Pa gels as compared to 640 Pa gels after 2–3 d in culture (Figure 2D). In agreement with the model’s predictions, we observed a decrease in cellular density in 40 Pa gels as compared to 640 Pa gels (Figure 5D; Figure 5—source data 1). As shown by immunolabeling, Yap was localized to the cytoplasm in the supporting cells of utricular bubbles maintained for 3 d in both 640 Pa and 40 Pa gels (Figure 5B,E; Figure 5—source data 2). However, consistent with a role of Yap in the regulating the utricular growth, the protein translocated into the nuclei in over 50% of supporting cells only in 40 Pa gels (Figure 5C,F; Figure 5—source data 3). Whereas cell proliferation was limited to the organ’s periphery in 640 Pa gels by 4 d in culture, the supporting cells throurgout the utricular macula reentered the cell cycle in 40 Pa gels (Figure 2F; Figure 5—source data 3). Yap labeling at the same time revealed that the pattern of the protein's nuclear translocation in both conditions was consistent with the pattern of supporting-cell proliferation (Figure 5—figure supplement 1). The expression of genes encoding the downstream targets of nuclear Yap, Ankrd1, Ctgf, and Cyr61, was also upregulated significantly in 40 Pa gels as compared to 640 Pa gels (Figure 5G; Figure 5—source data 4).
If the extensive growth and cellular proliferation observed in 40 Pa collagen gels are triggered by the nuclear translocation of Yap, perturbation of the protein's nuclear function should abrogate the effect of low stiffness. We therefore tested the effect of GFP-YTIP on the utricle’s ability to grow in a low-stiffness gel. Bubble cultures were established in a 40 Pa collagen gel and left overnight to allow the attachment of mesenchymal stroma cells (Figure 6A). On the following day either GFP-YTIP or control GFP virus was injected into each utricular bubble and the cultures were left for an additional 24 hr to allow the accumulation of GFP or GFP-YTIP protein. EdU was then added to the medium and the utricular bubbles were cultured for an additional 48 hr. As determined by Sox2 labeling, the areas of the utricular maculae of GFP controls increased significantly as compared to GFP-YTIP cultures (Figure 6B,C; Figure 6—source data 1). In accord with our previous results, the areas of the sensory epithelia in GFP virus-infected bubble cultures were similar to those observed in uninfected cultures after 4–5 d (Figure 2D), whereas GFP-YTIP cultures stalled at a size typical of control cultures at 2–3 d. The increase in macular area was reflected by a significant increase in the number of EdU-positive supporting cells (Figure 6B,D; Figure 6—source data 2). The supporting-cell density in GFP-YTIP virus-infected utricles was significantly higher than that in GFP virus-infected control cultures (Figure 6E; Figure 6—source data 3). These results strongly suggest that reduction in elastic force triggers supporting-cell proliferation through nuclear Yap signaling.
To determine whether the elasticity-limited model can explain the rate and pattern of supporting-cell proliferation in vivo, we analyzed supporting-cell numbers during utricular development. The macular area of the organ expanded dramatically between E15.5 and E17.5 (Figure 1); the number of supporting cells concurrently quadrupled (Figure 7A; Figure 7—source data 1). Over the same period, we observed a significant increase in supporting-cell density (Figure 7B). The rate of growth decreased thereafter and cell numbers plateaued by P2, when supporting-cell proliferation decreases dramatically (Burns et al., 2012b; Gnedeva and Hudspeth, 2015). However, we did not detect a significant increase in cellular density from E18.5 to P2 (Figure 7B; Figure 7—source data 2).
To visualize the pattern of supporting-cell proliferation we performed an EdU pulse-chase experiment. Pregnant mice were injected at E17.5 and the inner ears of their progeny were analyzed 12 hr later. Consistent with previous reports (Burns et al., 2012b; Gnedeva and Hudspeth, 2015), recently divided supporting cells were localized at the periphery of the utricle’s sensory epithelium, where small patches of EdU-positive cells were observed (Figure 7C). To quantify this phenomenon, we measured the average density of EdU-positive cells and the average outline of the macular sensory epithelium in multiple utricles at E18.5 (Figure 7D). This analysis demonstrated a clear maximum in the EdU intensity near the border of the macula (Figure 7D,E; Figure 7—source data 3). Using our elasticity-limited model to simulate the EdU pulse-chase experiment, we found a remarkably similar pattern of supporting-cell proliferation (Figure 7G). We explored the computational simulations to explain this pattern.
In our model, cells act as linear springs: the smaller a cell is compared to its equilibrium volume, the larger its internal pressure. The elasticity-limited model predicts that the elastic force produced by macular expansion creates a gradient of cellular pressure, with higher values at the center of the sensory epithelium and lower ones at the periphery where proliferation occurs (Figure 7H). Because cells with higher internal pressure have smaller volumes, the model predicts that the pressure gradient creates a corresponding gradient in cellular density. Although the average density does not increase after E17.5 (Figure 7B), we found that the density at the center of the macula significantly exceeded that at the periphery at E18.5 (Figure 7F; Figure 7—source data 4). As predicted by the model and demonstrated by our experimental data, the gradient in cellular pressure and density disappears concurrently with the cessation of macular growth at P2 (Figure 7F,H). In accord with the role of Yap in supporting-cell proliferation, an increase in cellular density is known to activate Hippo signaling and the subsequent degradation of Yap (Aragona et al., 2013; Dupont et al., 2011; Robinson and Moberg, 2011).
Our computational simulations also permitted evaluation of the role of stem cells in the developing utricle. Numerical simulations of a model invoking the original population of stem cells as the only source of proliferation yielded a pattern of cell divisions inconsistent with the experimental observations (Video 3; Source code 3). In fact, LGR5-positive cells, proposed to be the putative stem-cell population in the utricle (Wang et al., 2015), are restricted to the striolar region by E15.5 and are therefore unlikely to contribute to growth at the organ’s periphery.
In conjunction with published data (Burns et al., 2012b; Gnedeva and Hudspeth, 2015), our experimental results indicate that the sensory epithelium of the utricle grows primarily through the proliferation of supporting cells at its periphery. The rate of proliferation is self-regulated and decreases as the organ approaches its target size. To explore the mechanism underlying this process we implemented a combination of theoretical and experimental approaches.
Our main conclusion is that growth of the sensory epithelium is restricted by the elastic force produced by the surrounding nonsensory tissues. This force increases as the utricular macula expands in size, physically impeding cellular growth and proliferation. A similar mechanism has been shown to exist in the context of malignant growth, in which an external pressure can restrict the size of a tumor by physically opposing its growth and inhibiting cellular proliferation (Helmlinger et al., 1997; Cheng et al., 2009; Montel et al., 2011). In keeping with this idea, we demonstrate that reduction of the elastic force in utricular bubble cultures decreased the cellular density and allowed supporting cells to reenter the cell cycle and the sensory epithelium to nearly double in size. To our knowledge such an expansion has not been observed heretofore in any organ of the inner ear.
The elasticity-limited model not only explains the self-regulatory nature of macular growth but also suggests the involvement of the Hippo signaling pathway, which links mechanical force to cell-cycle arrest (Dupont et al., 2011; Robinson and Moberg, 2011; Aragona et al., 2013). By assessing the localization of Yap and supporting-cell density during normal development and in culture, we found that it is likely to control supporting-cell proliferation. Our simulations and data demonstrate that an elastic force, compressing the tissue, creates a cell-density gradient in the utricular macula, with a higher density at the center of the organ and a lower density at the periphery (Figure 8). High cellular density triggers the loss of nuclear Yap, causing supporting cells to exit the cell cycle. In support of this idea, removing the elastic constraints in organotypic utricular cultures decreased the supporting-cell density and resulted in nuclear Yap translocation and cell-cycle reentry throughout the sensory epithelium. Because all the sensory organs in the inner ear are subjected to mechanical constraints similar to those in the utricle, it is likely that Hippo signaling also controls cell-cycle exit in the developing organ of Corti. In fact, a mechanism of size control whereby mechanical force restricts the size of a tissue through activation of the Hippo signaling pathway has been conserved across a range of organs and species. Hippo signaling plays a major role in size control of the wing disk in Drosophila (Hariharan, 2015), determines the final size of the liver in mammals (Dong et al., 2007; Camargo et al., 2007), and regulates limb-bud regeneration in Xenopus (Hayashi et al., 2014).
Our experiments implicate a DNA-binding partner of nuclear Yap in the developing inner ear. When translocated to the nucleus, Yap can bind a variety of transcription factors to stimulate downstream gene expression (Yagi et al., 1999; Vassilev et al., 2001; Ferrigno et al., 2002). By specifically blocking the Yap-Tead interaction, we are able to arrest supporting-cell proliferation after injury and to reverse the effect of low stiffness on the expansion of utricular maculae in bubble cultures. As we demonstrated earlier (Gnedeva and Hudspeth, 2015), activation of SoxC transcription factors can restore proliferation even in utricles of young adult mice. SoxC proteins directly upregulate the expression of Tead2 (Bhattaram et al., 2010), which is highly expressed in the developing sensory epithelium and whose expression declines dramatically as supporting-cell proliferation ceases (Gnedeva and Hudspeth, 2015). In conjunction with the present results, this finding suggests that Hippo-initiated loss of the Yap-Tead transcription-factor complex is ultimately responsible for arresting the utricle’s growth and limiting supporting-cell proliferation.
During the last decade there has been a renewed interest in the study of the role of mechanical forces and their interplay with molecular signaling during development (Hernández-Hernández et al., 2014; Hamada, 2015; Pasakarnis et al., 2016; Dreher et al., 2016). Our work constitutes a new example of such interactions and raises the possibility that the elusive mechanism that triggers supporting-cell proliferation after the loss of hair cells in nonmammalian species is mechanical in nature. Both the extrusion of dying hair cells from the sensory epithelium and the subsequent transdifferentiation might affect the mechanical force sensed by the residual supporting cells, causing them to re-enter the cell cycle. Although further investigation of the role of Hippo signaling in the inner ear is required, biochemical manipulation of this pathway might aid in the recovery of hearing and balance after the loss of hair cells.
Experiments were conducted in accordance with the policies of The Rockefeller University’s Institutional Animal Care and Use Committee and the Keck School of Medicine of the University of Southern California. Atoh1-nGFP mice were a kind gift from Dr. Jane Johnson. Swiss Webster mice with timed pregnancies were obtained from Charles River Laboratories.
Embryos were extracted from euthanized mice and placed into ice-cold Hank’s balanced salt solution (HBSS, Life Technologies). Internal ears were dissected as described (Gnedeva and Hudspeth, 2015).
For organotypic cultures the ampullae of the anterior and horizontal semicircular canals and the nonsensory epithelium surrounding each utricle were left intact. The preparations were maintained for 3 hr at 37˚C in complete growth medium comprising DMEM/F12 supplemented with 33 mM D-glucose, 19 mM NaHCO3, 15 mM HEPES, 1 mM glutamine, 1 mM nicotinamide, 29 nM sodium selenite, 20 mg/L epidermal growth factor, 20 mg/L fibroblast growth factor, 10 mg/L insulin, and 5.5 mg/L transferrin (Sigma-Aldrich). Healing of the cut edges introduced during dissection allowed each preparation to reseal, creating an ellipsoidal structure termed a utricular bubble.
Collagen I was extracted from mouse-tail tendons (Rajan et al., 2006) and its concentration was adjusted to 2.0 mg/mL in 6 mM trichloroacetic acid (Sigma-Aldrich). 450 μL of collagen solution was mixed with 50 μL of 10X phosphate-buffered saline solution with phenol red pH indicator and neutralized by the addition of 11.9 mM NaOH and 1.3 mM NaHCO3 to initiate polymerization. Utricular bubbles were placed into the collagen solution at room temperature and incubated for 20 min at 37˚C to allow polymerization. The resultant cultures were then maintained at 37˚C in complete growth medium equilibrated with 5% CO2.
Chondrocytes were isolated from the cartilage surrounding the inner ear as described previously (Gosset et al., 2008).
Utricles were dissected in ice-cold HBSS and fixed in 4% formaldehyde for 1 hr at room temperature. Whole inner ears were fixed for 18 hr at 4˚C, treated with 0.88 M sucrose for 18 hr at 4˚C, embedded in Tissue-Tek O.C.T. (Sakura), and frozen in liquid-nitrogen vapor. Wholemounted sensory epithelia or 10 μm frozen sections were then blocked with 3% normal donkey serum (Sigma-Aldrich) in 500 mM NaCl, 0.3% Triton X-100 (Sigma-Aldrich), and 20 mM tris(hydroxymethyl)aminomethane (Bio-Rad) at pH 7.5. The primary antisera—goat anti-Sox2 (Santa Cruz), rabbit anti-Myo7A (Proteus Bioscience), rabbit anti-GFP (Torrey Pines Biolabs), mouse anti-Yap (Santa Cruz), and rabbit anti-Yap (Cell Signaling)—were reconstituted in blocking solution and applied overnight at 4˚C. For labeling with E-cadherin antibodies from clone DECMA-1 (Millipore) no Triton X-100 was used in the blocking solution.
Samples were washed with phosphate-buffered saline solution supplemented with 0.1% Tween 20 (Sigma-Aldrich), after which Alexa Fluor-labeled secondary antisera (Life Technologies) were applied in the same solution for 1 hr at room temperature.
Phalloidin conjugated to Alexa 633 was used to label filamentous actin and nuclei were stained with 3 μM DAPI.
EdU pulse-chase experiments were initiated by single intraperitoneal injections of 50 μg EdU (Life Technologies) per gram of body mass. Animals were sacrificed at the indicated times and the cells in the utricular sensory epithelia were analyzed by Click-iT EdU labeling (Life Technologies).
The AdEasy Adenoviral Vector System (Chartier et al., 1996; He et al., 1998) was used to create adenoviral vectors containing the full-length coding sequence of green-fluorescent protein fused to the Yap-Tead interfering peptide (GFP-YTIP; Addgene plasmid 42238) under the control of a cytomegalovirus promoter. Viral particles were amplified in HEK cells and purified by CsCl-gradient centrifugation followed by dialysis (Viral Vector Core Facility, Sanford-Burnham Medical Research Institute). Each utricle was dissected at E16.5-P4 and infected in 200 μL of culture medium with 10 μL of virus at a titer of 1010 PFU/mL. Alternatively, 1 μL of the virus was injected into the utricle in 3D organotypic cultures. Ad-GFP virus (Vector Biolabs) at the same titer was used as a control. One day later 3 mL of culture medium was supplemented with 10 μM EdU to label mitotic cells.
The sample preparation and analysis for RNA sequencing have been described in detail (Gnedeva and Hudspeth, 2015). For qPCR, utricles at each developmental stage were isolated by microdissections and treated with 0.5% Dispase I (Sigma) for 15 min at 37˚C to isolate the sensory epithelia. For each sample, total RNA from 7 to 12 utricular maculae was isolated by a standard protocol (RNeasy Micro Kit, Qiagen) and used to create a cDNA library. The qPCR primers were designed with PrimerQuest (Integrated DNA Technologies). Relative gene-expression levels were obtained by normalization to the expression of Gapdh in each sample. qPCR analyses were performed on an Applied Biosystems 7900HT Sequence Detection System with FastStart Universal SYBR Green Master mix (Roche Applied Science).
The standard Western blotting protocol (BioRad) was used with the following specifications. The utricles containing sensory epithelia, transitional epithelia, and underlying mesenchyme were isolated by microdissection and lysed in 50 μL RIPA lysis buffer for 30 min at 4 ˚C and sonicated thrice at low power for 10 s each with the sample kept on ice between the sonications. The total protein concentration in each sample was determined by the BCA assay (Thermo Fisher). A NuPAGE 12% Bis-Tris Protein Gel (Thermo Fisher) was used to resolve the proteins in 5 μg of each sample. The proteins were transferred to a nitrocellulose membrane (BioRad) and blocked for 1 hr at room tempirature in a 5% solution of skim-milk powder (Sigma-Aldrich) in tris buffer (BioRad) with 0.1% Tween 20 (Sigma-Aldrich). After the primary antibodies—rabbit anti-Yap (Cell Signaling) and rabbit anti-H3 (Millipore)—had been reconstituted at 1:10000 in tris buffer blocking solution containing 0.1% Tween 20% and 5% normal sheep serum (Sigma-Aldrich), the membrane was incubated over night at 4 ˚C. After 5 30 min washes at room tempirature in TBST, the anti-rabbit HRP secondary antibody (Millipore) was applied in TBST for 1 hr at room temperature. Horseradish-peroxidase activity was detected with the Amersham ECL Western Blotting System (GE Healthcare Life Sciences).
Confocal imaging was conducted with an Olympus IX81 microscope equipped with a Fluoview FV1000 laser-scanning system (Olympus America). To determine the areas of utricular maculae and to enumerate Sox2-positive supporting cells, we imaged wholemounted utricles as Z-stacks and selected representative areas of slices for maximal-intensity Z-projections. To avoid counting Sox2-positive hair cells, Myo7A staining was performed to eliminate the optical sections through the hair cell layer. The statistical significance of comparisons between cell counts was determined by two-tailed Student’s t-tests. The error bars in figures represent standard errors. P-values less than 0.05 are represented by a single star, those less than 0.01 by two stars, and those less than 0.001 by three stars.
For each developmental time point, Sox2-positive cells in the high-resolution image of a maximal-intensity Z-projections of a wholemounted utricle were enumerated automatically with the image-analysis software CellProfiler (Carpenter et al., 2006). We also used this software to automate the areal measurements of utricular sensory epithelia. The algorithms employed for these tasks were tested against manually quantified images and the error of the algorithms was estimated to be consistently lower than 2%.
The intensity of fluorescent staining was measured as mean gray value using ImageJ, which yields a unitless relative value. In a single-channel image, the intensity of a black pixel is 0 and that of a saturated pixel is 255. For RGB images, the maximal intensity of a pixel in each channel is 85 (255/3).
To produce the EdU-intensity plot in Figure 7D we used E18.5 utricles labeled for the detection of EdU and Sox2. We then took maximal Z-projections of five EdU-positive images, applied a Gaussian-blur filter to smooth the punctae observed in individual cells, and centered, aligned, and averaged the images. By thresholding and averaging over Sox2-positive channel we calculated the mean outline of the sensory epithelium. To produce the average EdU intensity plots in Figure 7E, we performed a spline interpolation of the calculated mean outlines to remove sharp variations. From each image we then copied strips of 400 pixels perpendicular to, and centered on, the smoothed outline. Finally, we calculated the average over all the strips to obtain a plot of the mean EdU density against the distance from the utricle’s perimeter.
For the quantifications of cellular densities, EdU labeling, and viral infections at the periphery versus center of the utricle and in experiments in vitro, cells were enumerated in areas of 2500–10,000 μm2.
We estimated the elastic modulus of each collagen gel with a piezoelectric bimorph system. An individual gel was mounted between a glass slide and a glass coverslip whose diameter exceeded that of the gel. The mounted gel’s original cross-sectional area (A0) and height (L0) were measured. The free end of a piezoelectric bimorph cantilever (PZT-5H, Vernitron Piezoelectric Division, Bedford, OH) mounted on a micromanipulator (MP-285, Sutter Instrument, Novato, CA) was then brought into contact with the glass coverslip. To deliver forces to the gel, the bimorph’s base was displaced vertically by the micromanipulator under the control of custom software (Source code 4) written in LabVIEW (version 10.0, National Instruments, Austin, TX). The micromanipulator was lowered in 20–50 sucessive steps of constant amplitude, each with a duration of 1 s and with separations of 2 s. The increments for different gels ranged from 0.625 μm to 5 μm. Each displacement induced flexion of the bimorph, resulting in an electrical potential owing to the piezoelectric effect. This signal was amplified and bandpass filtered at 0.1–10 Hz (Grass P55, Astro-Med, Inc., West Warwick, RI). The filtered signal was recorded at sampling intervals of 500 μs by a computer running a custom program in LabVIEW.
We calculated the voltage change for each force step as the difference between the voltage prior to the pulse and the maximal voltage after the pulse’s onset. We next determined from each response both the force delivered by the piezoelectric bimorph and the vertical displacement of the coverslip. For a bimorph cantilever whose free end contacted a glass coverslip, the force generated at the bimorph’s free end was , in which = 2.62 mN/V is the bimorph's sensitivity, = 6.5 mm is its width, and = 22 mm is its length (Corey and Hudspeth, 1980). The stress applied to the gel was then . The displacement of the bimorph’s free end was , in which = 645 N·m−1 is the bimorph's stiffness. The difference between the displacement of the bimorph cantilever’s base and that of its free end yielded the change in height () of the gel and in turn the strain ε experienced by the gel, .
We estimated the elastic modulus for each gel by fitting the stress-strain relations to the relation , in which is an estimate of the population intercept. All fits possessed coefficients of determination exceeding 0.98. Errors are standard errors of the elastic moduli of multiple gels of a particular composition.
We conducted numerical simulations using CompuCell 3D, a program that allows rapid creation of Glazier–Graner–Hogeweg (GGH) Monte Carlo models (Swat et al., 2012). We used a two-dimensional hexagonal lattice and a standard form for the interaction energy of the system:
The first term on the right describes the contact energy between neighboring cells owing to adhesive interactions. If a cell of type occupies the lattice site and a cell of type occupies the lattice site , then is the boundary energy per unit of contact length. The delta function in the summation indicates that the contact energy between pixels of the same cell is zero.
The second term on the right describes a constraint on the cellular volume, which in two dimensions corresponds to the number of pixels that a cell occupies. A cell whose volume deviates from the target value produces an increase in the effective energy proportional to , the two-dimensional elastic modulus of the cell. This modulus corresponds to the stiffness of a single cell and is distinct from the modulus E of the elastic boundary.
The third sum on the right imposes a constraint on the cell’s surface area, which in two dimensions corresponds to the cellular perimeter. Deviations of the surface area from the target value increase the effective energy proportionally to , the elastic modulus of the cell’s surface.
Cell dynamics consists of a series of index-copy attempts using a modified Metropolis algorithm. Before each attempt the CompuCell 3D selects a pair of target and source sites, and If different cells occupy these sites the algorithm then sets with probability given by the Boltzman acceptance function:
in which is the change in effective energy, calculated from Equation 1 and is a parameter describing the amplitude of cell-membrane fluctuations. A Monte Carlo step is defined as index-copy attempts and sets the computational time unit of the model and, in biologically relevant situations, the step is proportional to experimental time (Swat et al., 2012). We converted this measure into actual time by multiplying it by and fitting this scaling factor to experimental data. The value of depends on the average value of . We confirmed by numerical simulations that changing the value of modifies the values of and other parameters required to fit the experimental data, but it does not change the result of our simulations in any significant way. We likewise converted areas in pixels to actual areas by multiplying by a factor and fitting the results to the experimental data. For further details on CompuCell 3d or the used Metropolis algorithm we refer the reader to Swat et al. (2012).
We studied the dynamics of cellular proliferation and compared two models to explain the observed pattern of growth. can represent a value either for supporting cells (SC) or for the surrounding elastic boundary (EB). The contact-energy term of Equation 1 is therefore described by two types of contact energies, and . To simplify our notation we define and .
We assumed that the elastic boundary applies hydrostatic compression over the growing sensory epithelium: the stress created by the boundary can change the volume but not the shape of the sensory epithelium. If the elastic boundary is initially unstressed, expansion of the utricle to a surface area stretches the elastic boundary by and increases its elastic energy (Landau and Lifshitz, 1970) by an amount . In our two-dimensional model this surface area corresponds to the perimeter of the utricle, whereas in three dimensions the relevant value would be the perimeter multiplied by the thickness of the tissue.
The first term of Equation 1 can be rewritten as
in which the first sum is evaluated over all the pixels at the boundary of the tissue, for only those contribute to the sum. The second sum is effected for all the pixels in the bulk. The initial sum evaluates as . Multiplying this term by reveals that the energy of the elastic boundary is proportional to . Identifying with the elastic modulus of the boundary, Equation 1 becomes
For computational efficiency the sums in the first two terms on the right-hand side of Equation 3 are evaluated over only a small neighborhood of pixels. This simplification and the discrete nature of the simulations creates artifacts that include sharp corners and anisotropic compression of the tissue when the force produced by the elastic boundary is large.
Cells in our model have a refractory period after division until they can divide again, which represents the time required for a cell to transverce the cell cycle. After this refractory time a cell divides into two cells when . Each cell accordingly grows under the influence of the volume-energy term until it has passed the refractory period and its volume is exceeds the division volume , whereafter it splits into two cells that resume growth until they again meet the conditions for cell division. The refractory time for each cell is picked from a Gaussian distribution with mean and standard deviation .
In our simulations of the elasticity-limited model the rate of cell proliferation decreases to almost zero as the tissue reaches its final size (Figure 1E). We observe, though, that the rate of cell divisions is not exactly zero and some residual growth occurs on long timescales. Because of the stochastic nature of our model, random fluctuations eventually push a cell beyond its division volume and initiate division. Although the probability for this to happen decreases with the size of the utricle, it is never zero. This behavior poises the utricle in an unstable situation in which, without the presence of an additional mechanism, it grows indefinitely at a very slow pace. We suspect that this instability arises in the utricle and that the Hippo pathway is responsible for completely arresting growth once the utricle has reached its final size. In our model we simulate the action of this pathway in a heuristic way: cells at high density cannot easily increase their volumes and therefore remain unable to divide for a long time. We use this as a measure of density: if a cell is unable to divide after a time , we render that cell quiescent in our simulation, so that it cannot divide even if it reaches its division volume.
For the elasticity-limited model we adjusted the value of such that the tissue growth ceased when the number of cells reaches the experimentally observed value. For the morphogen-limited model we considere a much lower value of such that in the absence of morphogen the sensory epithelium would grow to occupy the entire simulation domain.
The dynamics of the morphogen concentration is simulated by integrating the diffusion equation
in which is the diffusion constant of the morphogen. Each cell secretes morphogen molecules at a rate from its center of mass . When the value of at position reaches a certain threshold the cell stops dividing, even if it has reached a volume of . The integration is conducted with finite differences in space and a forward Euler’s scheme in time. During numerical integration of Equation 5, time is advanced by every Monte Carlo step. CompuCell 3D internally rescales the diffusion equation and adjusts the required number of integration steps per Monte Carlo step to ensure the numerical stability of the method (Swat et al., 2012).To simulate EdU pulse chase experiments in the elasticity-limited model (Figure 6D) we added a variable to each cell. When the utricle reaches a stage equivalent to E17.5 we activate EdU labeling by setting in each newly divided cell. The time EdU labeling is turned on is adjusted so that the number of labeled cells matches the experiments. After labeling is turned off the concentration of EdU dilutes by half with every cell division and we assume that it becomes undetectable after four divisions, that is, for .
Internal cellular pressure is calculated from the model as (Swat et al., 2012). Cells with high internal pressure have smaller volumes and therefore higher density. These cells in high density areas are therefore less likely to reach and divide.
The utricular macula and non-sensory epithelium form an approximately ellipsoidal chamber filled with endolymphatic fluid and surrounded by mesenchymal stroma and cartilage. The sensory epithelium is located at the bottom of this ellipsoid and is surrounded by band of non-sensory tissue called the transitional epithelium (Figure 1—figure supplement 1).
The compressive stress produced by the elastic matrix is transmitted to the epithelial layer as a combination of normal and tangential stresses. Given that the surface upon which the utricular macula lies is relatively flat, we assumed that the normal stress makes little contribution in changing its shape, and therefore could be neglected. This assumption simplified the model's geometry and allowed a two-dimensional representation of the system. The tangential stress is transmitted with the plane of the epithelium. In our two-dimensional representation this stress acted on the boundary of the utricular macula (Figure 1—figure supplement 1). We assumed that all the deformations produced in the different tissues are elastic and linearly proportional to the stresses.
Our two-dimensional representation of the utricle comprises a patch of sensory epithelium surrounded by a band of transitional epithelium of Young’s modulus and embedded in an elastic matrix of Young's modulus (Figure 1—figure supplement 1). We must first demonstrate that these two materials can be described by one band of tissue with an effective Young’s modulus and that the final size of the utricle depends on .
In polar coordinates the band of transitional epithelium has respective inner and outer radii and . The elastic matrix has an internal radius of ; for the sake of simplicity, we assume that it extends to infinity. We designate the displacements of the transitional epithelium and those of the elastic matrix and respectively. Each of these displacements follows , in which is an arbitrary integration constant (Landau and Lifshitz, 1970). Hereafter = 1, 2 represents the variables for the transitional epithelium or elastic matrix respectively. Because the problem is radially symmetrical, only the radial components of the divergence of are nonzero and so we may omit the vector arrow. In polar coordinates the equation reads
The general solution of this equation is , in which and are integration constants determined by the boundary conditions. The components of the strain tensor are then , , and . The radial symmetry also implies that the non-diagonal components of the strain tensor are zero.
Using the stress-strain relation for a homogeneous deformation we can write the radial component of the stress tensor as
in which is the Poisson ratio that we assume to be the same for the transitional epithelium and the elastic matrix.
To calculate the different integration constants, we next introduce the boundary conditions of the system. For the elastic matrix, we assume that deformations decay to zero at infinity, that is, , which implies that = 0. Because there can be no gaps in the boundary between the two regions, the displacements on both sides of the boundary are equal and . For the system to be in mechanical equilibrium the normal stresses on both sides of the boundary have to be equal, . Finally, we assume that the sensory epithelium generates a hydrostatic pressure while expanding, so that . Using these boundary conditions, we can now solve for the three remaining constants to obtain:
Inserting these expressions into the equations for and yields solutions for the system’s deformations and stresses. Eliding this step, we instead show how the combined stiffnesses of the transitional epithelium and elastic matrix can be combined into a single effective term in our simulations.
Following the same steps as before we can calculate the deformation of the effective elastic band of internal radius , external radius and Young’s modulus under an internal pressure :
We next require that the expansion of the sensory epithelium under the effect of this elastic band equals that under the conbined effect of the transitional epithelium and elastic matrix. The change in radius the sensory epithelium is given by the deformation of the inner radius of the transitional epithelium deforms, that is, , this relation implies that . We can therefore calculate that the effective Young’s modulus is:
In our bubble-culture experiments we replace the cartilage surrounding the utricle by a collagen matrix. In the mathematical representation of the system this condition is equivalent to a change the value of . To more easily see the dependence of on we can consider a thin band of transitional epithelium and expand Equation 12 around to obtain:
from which we can see that the effect of reducing in our experiments can be approximated by a reduction of in our simulations.
Finally, we can calculate how the radius of the utricle depends on the elastic modulus . The net rate of cell division—the rate of division minus the rate of apoptosis—depends on pressure (Ranft, 2012). The homeostatic pressure is the value at which the rates of division and apoptosis are equal and the net division rate is accordingly zero. The disk of tissue grows as long as the pressure exerted by the elastic band is smaller than the homeostatic pressure. Therefore, from Equation 11 we can see that the total change in radius once the utricle has reached equilibrium is:
The change in radius is proportional to , and therefore the change in area of the utricle is proportional to . The equilibrium areas of simulated utricles for different values of the Young’s modulus agree with this expectation (Figure 1F).
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Tanya T WhitfieldReviewing Editor; University of Sheffield, United Kingdom
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.
Thank you for submitting your article "Mechanical force restricts the growth of the murine utricle" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by Tanya Whitfield as the Reviewing Editor and Andrew King as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Alexander Fletcher (Reviewer #1) and Jennifer Stone (Reviewer #3).
The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
Gnedeva et al. address the very interesting question of the mechanisms responsible for growth arrest during development, in particular the role that mechanical constraint plays in this process. This question is especially pertinent concerning the mammalian inner ear, since growth arrest might be linked to the inability of mammals to produce new sensory hair cells in adults, making hearing and balance dysfunctions often irreversible.
This work combines analysis of the murine utricle in vivo and in a novel ex vivo culture system with computational modelling to address the mechanisms underlying the abrupt growth arrest of this tissue observed late in embryonic development. The authors conclude that, as the utricle grows, it experiences a resistive force from the surrounding elastic environment. This force creates a cell-density gradient across the tissue that is interpreted through the subcellular localization of Yap, and corresponding gradient of supporting-cell proliferation. The authors demonstrate that this hypothesis can explain the observed tissue growth dynamics when the stiffness of the surrounding collagen gel is reduced. These results reflect a tissue-level coordination of proliferation based on mechanical conditions at the boundary, and are not consistent with a model where growth is driven by stem cells located at the centre of the utricle.
1) Please incorporate the extra details requested by reviewer 1 into the model.
2) Further experimental work is needed to support the conclusions concerning the regeneration post-injury in both soft and stiff gels.
a) The role of Yap should be shown more clearly both during regeneration upon injury of the utricle and in the doubling of the utricle grown in a softer gel. Please see comments from reviewer 2.
b) The authors should determine if dividing cells are indeed Sox2+ (sensory epithelial) cells and if they give rise to hair cells, in order to determine whether their observations are relevant to hair cell production. This could be done with pulse/fix experiments for Sox2 and pulse/chase experiments for identification of post-M cells. Please see comments from reviewer 3.
3) Reviewer 3 has made a number of suggestions for additional literature that could be cited to acknowledge previous relevant work. Please take this into account.
4) It is essential that all methods are described in sufficient detail to allow others to repeat the study. Please see the comments from reviewer 3.
5) All three reviewers have suggested numerous small changes or corrections that would help to clarify or improve the manuscript. Please attend to these where possible.
The full reviews are appended below for further information.
Overall, I found the manuscript to be clearly written, well motivated, and an important contribution to our understanding of the role of mechanics in tissue growth and size control. As such, I do not have any explicit requests for additional work, though I do have a number of queries and concerns that I would like to see addressed or clarified in the manuscript.
There are a few details missing from the computational model that prevent a full critique of the results. First, in subsection “Numerical simulation of utricular growth and cellular differentiation” the authors refer to "the surrounding elastic boundary", but based on Figure 2B it seems that there is also an elastic collagen gel underlying the cells. Thus, for a stiffer gel, cell movement in the centre of the tissue (not just at the periphery) should be affected; could this affect the relative timescales of mechanical relaxation and cell growth, and thus potentially affect the observed growth dynamics? Second, the authors assume that the link between mechanical force and cell-cycle arrest occurs specifically via the pressure (compression) on each cell, but it was not immediately clear to me that this must be the way that mechanosensing occurs in this system; has this been demonstrated experimentally?
A further assumption is that the growing utricle is essentially two-dimensional (Results section); yet the authors state in subsection “Dissection and culturing of utricles” that the utricular bubble is "a spherical structure". It was not quite clear to me how good the planar approximation is for this tissue, and whether tissue curvature could complicate the mechanical response to a stiff surrounding medium.
In addition, in subsection “Dynamics of utricular development” the authors refer to "processes such as cell division, differentiation, molecular signaling, and physical force", but do not refer to apoptosis or extrusion from the utricle. Since they do refer to apoptosis in subsection “Growth of a utricle embedded in an elastic matrix”, it would be helpful to know whether any cell death is observed during the experimental period in the ex vivo culture system. In other tissues, apoptosis and live-cell extrusion have been demonstrated to locally relieve stress, thus contributing to mechanical homeostasis in the tissue (e.g. Eisenhoffer et al., doi:10.1016/j.tcb.2012.11.006). It would therefore be of interest to know whether apoptosis occurs in the utricle, since this could affect the authors' mechanical argument.
I cannot comment with authority on the experimental protocols, but was unsure whether one might expect any matrix or collagen remodelling; is it reasonable to assume no mechanical feedback from the utricle back on the surrounding medium over the timescale of interest? Also, the authors use the phrase "mechanical force" in the Title and Abstract, but this is quite a general term, and could be made more precise – the authors' conclusion is that it is in particular the resistive, elastic force from the surrounding medium that drives growth arrest.
Overall this is a nice study and the paper is clearly written and pleasant to read. Yet I think some points need to be addressed or clarified.
1) Figure 3: The authors have shown in Figure 1 that the utricule area increases rapidly between E15.5 and P2 after which growth slows down and stops. Why then looking at Yap protein level only at P14. If a decrease in Yap activity accounts for this growth arrest, one would expect a decrease in Yap protein and activity as early as P2. Same applies to the expression of Yap target genes and genes encoding inhibitors of Yap translocations.
2) Figure 4: The authors show an increase of proliferation and of Yap protein during regeneration upon injury of the utricule. These are only correlative observations and no proof that the observed proliferation depends on Yap, nor that Yap plays a role in regeneration.a) The authors should show, using the same canonical Hippo target genes they have used in Figure 3B that Yap activity is indeed increased during regenerationb) The authors should perform this experiment while blocking Yap function using for example the blocking peptide to show that regeneration is then blocked
3) Figure 5: Some controls and a more in-depth analysis are required to convincingly show that Yap activity is solely responsible for increase proliferation and a doubling of the utricule in a less stiff gel. Also here,a) The authors should show, using the same canonical Hippo target genes they have used in Figure 3B that Yap activity is indeed increased in a softer gel as compared to a stiff gelb) The authors should use an alternative method, next to the blocking peptide, at best a yap1 mutant, to show that utricule area and proliferation also do not increase in a soft gel.
4) Figure 5 text: The last paragraph of subsection “Yap is necessary for supporting-cell proliferation”
"Unexpectedly, the supporting-cell density in GFP-YTIP-infected utricles was significantly lower than that in GFP-infected control cultures (Figure 5G). As a result, although the area doubled, the total number of supporting cells in GFP-infected cultures increased only 20% in comparison with GFP-YTIP cultures (Figure 5H). " This is very unclear. "Lower" should be "higher". The fact that "the total number of supporting cells in GFP-infected cultures increased only 20% in comparison with GFP-YTIP cultures" is to my understanding the cause and not the consequence of the higher density.
"These data suggest that Yap activity is also required for cell spreading. Additional experiments are necessary to validate this prediction and to uncover the underlying mechanism." I think this statement is fully in the scope of this paper and the authors should show this increased spreading by measuring the cell area and the distance between individual cells.
5) Figure 5—figure supplement 1: The authors should clarify in which panel of B, one looks at an apical or at a basal view. More importantly, hair cells in the utricule in 40Pa gel appear much bigger than in a 640Pa gel. The authors do not mention this observation. Given that Yap is not expressed in hair cells, how do the authors explain this?
6) Figure 6: It is not clear to me how the cell density can increase so much between E15.5 and E17.5 in vivo (Figure 6B) if both the supporting cell number (Figure 6A) and the utricule area (Figure 1A) increase at about the same speed. In this case, I would expect the cell density not to change much.
7) a) It is not clear whether the authors interpretation is that both the decrease in cell density (or increased spreading) and increase in proliferation are driven by increased Yap activity or whether the decrease in stiffness results in decrease in cell density (yap independently) and the decrease in cell density, in turn, drives yap-dependent proliferation. This should be clarified in a precisely justified manner by the authors.b) The initial question the authors stated in the Abstract and in the Introduction is concerning the link between inner ear growth arrest and inability to regenerate hair cells. Although the authors show that support for cell proliferation can be enhanced by lowering the elastic mechanical forces of the surrounding tissue, they do not show the consequences on hair cell regeneration. It would be very interesting that the authors damage the hair cells of ex-vivo cultured utricules in both 40Pa and 640Pa gels to compare their capacity to regenerate hair cells.
This is a beautiful paper examining something quite novel and significant – the effect of organ stiffness and epithelial cell density on the developmental growth of the sensory epithelium for a balance organ, the utricle. Overall, the paper is well composed, and the data that are presented are compelling and easy to understand. The role of stiffness on inner ear organ development has been understudied, and this is the first report to my knowledge of the role of Hippo signaling in controlling addition of new cells to a growing organ in the inner ear. Unfortunately, the paper needs quite a bit of editing to make it suitable for publication, as several statements are vague or misleading, and some aspects of the data are hard to interpret because they are incomplete or presented in an unclear or in one case incorrect manner. Further, many methods were not described.
The lack of line numbers and labels on the figure made it particularly challenging to review this proposal.
Below, I summarize the major concerns for each section.
Please take into consideration that several labs (Forge, Raphael, Stone labs) have shown there is some spontaneous replacement of vestibular hair cells in adult rodents, probably due to supporting cell to hair cell conversion, and perhaps even in humans (Taylor et al., 2015). Plus, Warchol et al., (1993) showed that cultured adult utricles from guinea pigs and humans have considerable cell division in the macula (presumably of supporting cells) and Rubel et al., (1995) showed cell division leading to supporting cell replacement occurs in adult rodent vestibular epithelia.
Add "the" in between "of" and "utricle's development".
The statement that hair cells almost never contact one another is misleading, especially since a recent paper (Pujol et al., 2014) showed that in the adult mouse utricle, a large proportion of the type II hair cell population is in contact via basolateral processes.
In the fourth sentence the statements are untrue or misleading and should be modified. They suggest that supporting cell division does not proceed into the postnatal period and vestibular hair cells are not replaced after birth. However, Ruben (1967) showed that significant numbers of new vestibular epithelial cells are made postnatally; this paper should be cited. Further, Burns et al., (2012) showed continued macular cell proliferation into the neonatal period and production of several hair cells from the products of those dividing cells. The wording suggests that the authors have not read several papers showing that there is considerable regeneration of vestibular hair cells in adult rodents, probably by direct transdifferentiation. By these accounts, percentages of regenerated hair cells range from 20% to 70% of normal levels (in some regions). Several references should be cited, including Forge et al., 1993; Forge et al., 1998; Kawamoto et al., 2009; Lin et al., 2011; Golub et al., 2012; and Slowik et al., 2013. Further, these findings should be taken into consideration and addressed in the Discussion in the context of this paper's findings during development.
Second paragraph, third sentence. Since White et al., (2012) saw the EGF effect in chicken auditory organs, I do not believe this sentence helps. Several groups have examined EGF signaling in adult mouse utricles (e.g., Oesterle's group and Gao's group). Although White et al., assessed effects of EGF on mouse cochlear supporting cells, they were dissociated cells, not intact epithelia.
I would consider mentioning here work from the Pirvola and the Burns groups on the failure of cell cycle effectors such as cyclin D to promote division in mouse utricles. This seems quite germane to this discussion.
Results section. Why assume supporting cells secrete a growth-inhibiting molecule? Why not hair cells, or why couldn't supporting cells secrete a growth-promoting molecule? Would the observations still rule out a morphogen model if the morphogen were growth-promoting?
Figure 7. I disagree on several points here; what is stated does not match the figure. First, cytoplasmic Yap labeling looks strong in the images. I think it is clear from the images in Figure 3E that no nuclear Yap is seen in the central region while some nuclear Yap is seen in the periphery. It is clear however, that some cytoplasmic Yap is retained in the periphery, which makes it a bit hard to discern.
Subsection “Yap signaling during development and regeneration of the murine utr”. Need to add Kawamoto et al., 2009 to Golub et al., 2012 to reference list.
Subsection “Yap signaling during development and regeneration of the murine utr”, Figure 4. This is an odd experiment to perform, as opposed to targeted hair cell ablation, which could be accomplished with neomycin in vitro using published methods. One concern is that the authors are not actually examining cell divisions related to sensory epithelial cell repair. Were the EdU+ cells Sox2+? This would indicate if the cells dividing were indeed supporting cells or some other cell, such as from the stroma or the transitional epithelium. Did the authors investigate the fate of the post-M cells in that region? This would indicate divisions are leading to hair cells. I see this as a major problem with interpreting the data.
Subsection “Yap is necessary for supporting-cell proliferation”. No – the density was higher! Did you mean macular area? Overall, this section is confusing. Here is what the data in Figure 5D-H say to me: In YTIP-treated epithelia, macular area was halved (E) and dividing cell numbers were reduced to ~20% of normal (H) while supporting cell density doubled (G). This is not surprising to me, but I don't understand why this suggests that cell spreading was reduced. Further, it is strange to me is that epithelial cell numbers were not dramatically reduced, given the EdU data.
Subsection “Mechanical force confines cellular proliferation to the utricle’s periphery”. Utricular growth does not cease at P2. Burns et al., 2012 (Figure 1C) show macular area increases significantly out to P8.
Discussion section This is a short Discussion. The authors should consider addressing the following as well:
Does Yap play a similar role in limiting growth of any other sensory or neural tissues (retina, olfactory epithelium, other)? It would be a good addition to discuss this here, briefly.
Is the favoured model consistent with all forces controlling cell division being restricted to the sensory epithelium (i.e., related to epithelial cell density)? Might some forces be derived from outside the macula? If so, what could they be?
What significance is the boundary of transitional epithelial (TE) cells with respect to limiting supporting cell division and growth? What is Yap expression like in the TE?
Why might supporting cell division eventually cease? What changes in the stiffness of cell density might allow this to occur?
Do you feel you have ruled out the possibility that both types of forces – morphogen and elasticity – might be actively controlling utricular size?
During damage, how might extrusion or degradation of hair cells alter Hippo signaling?
Video 3: This seems like new data; why isn't it included in the Results section?
No methods were provided for the following: Western blots, RNAseq, virus experiments! For the Western and RNAseq, were sensory epithelia only used or were whole utricles used?
Subsection “Mechanical force confines cellular proliferation to the utricle’s periphery”:
– What% of the macula was sampled?
– Were cells from each zone (striola, lateral extrastriola, medial extrastriola) sampled?
– In utricles, type II hair cells are also Sox2Sox2+; how did you assure you were not counting Sox2Sox2+ hair cells? (This is particularly an issue with manual counting methods).
– What was the degree of error with automated areal measurements? (This should be known, since the algorithms were compared to manual counts).
Subsection “Mechanical force confines cellular proliferation to the utricle’s periphery”. There is no Figure 4D. What does this mean: "calculated the mean outline?" Is this area?
Subsection “Mechanical force confines cellular proliferation to the utricle’s periphery”. There is no Figure 4E. Since I cannot link this information to any figure, I cannot interpret or assess this paragraph.
Figure 2, here, and for several places throughout the manuscript: N's should be provided for each group (e.g., N=3 for 40 Pa and N=3 for 640 Pa)
– Figure 2D source data file is very confusing. I opened the files in Excel. First, it states supporting cell area instead of epithelial area. Second, it is not clear which group (40, PA 640 PA or in vivo) is described in each column, since there seem to be column titles missing for each column. Column B has no heading at all.
– Figure 2E source data. Again, I cannot see the column titles, so I am not sure they are there. I recommend carefully checking all source data to make sure they are complete (have all headings) and match the terms used in the figures and text.
Figure 3D, E, are you confident the cells with the higher Yap intensity are in the sensory epithelium, or might they be transitional epithelial cells? What are the units for fluorescence intensity?
– It is impossible to tell that hair cells do not express Yap. The reader would be more convinced if fluorescence were shown to not overlap with cytoplasmic hair cell marker labelling. Another possibility is to reference Figure 5—figure supplement 1, since it is quite clear there that hair cell cytoplasm isn't labelled.
Figure 4 legend, regarding the statement "The same cells enter the cell cycle". This is true for some cells, but lots of dividing cells seem NOT to have translocated nuclear Yap. This could be misleading or suggest a bias. These images should be shown at higher magnification, to better enable readers to see the Yap and EdU labelling in the nuclei.
Figure 5, did the authors examine the cell-autonomous nature of the effect of GFP-YTIP? Do they know if there was a significant change in the GFP+ versus GFP- cells? This would be very important to understand, as it could expand our understanding of how Yap functions.
– I think it's important to note that the supporting cell density decreased with low stiffness (Figure 5C). How about effects on total supporting cell numbers?
Figure 6F, I do not understand what is being shown here. What is the dotted line? I do not see any shaded area around the line. What does x axis represent?
– The authors should clearly state which cells were counted by including "supporting" or "macular" in the y-axis labels whenever possible.
– What is significance of asterisks in the graphs? This should be explained in the legend.
[Editors' note: further revisions were requested prior to acceptance, as described below.]
Thank you for resubmitting your work entitled "Elastic force restricts the growth of the murine utricle" for further consideration at eLife. Your revised article has been favourably evaluated by Andrew King (Senior editor), a Reviewing editor (Tanya Whitfield), and three reviewers.
The manuscript has been substantially improved and reviewers 1 and 2 are happy that their queries and concerns have been addressed. Reviewer 3 is very positive overall but has some remaining suggestions concerning wording or comparison to published studies. We are therefore returning the manuscript to give you the option of addressing or rebutting these concerns, which should be quick to do. The detailed comments are included below.
This excellent paper has been significantly enhanced by the added experiments and the careful editing of the text. I have only a few additional suggestions before publication.
Abstract. I recommend rephrasing the first sentence to "limited ability of cells in the inner ear to proliferate", as inner ear proliferation would suggest duplications of the organ.
Introduction. I am in favor of the wording now, with two small gripes.
The Warchol et al., 1993 Science paper refutes the last statement that "supporting cells [...] lose the ability to reenter the cell cycle after neonatal stages", since they showed that cultured utricles from adult humans and guinea pigs attained numerous tritiated thymidine-labeled supporting cells over time. I realize this is a stand-out paper, but I wonder if you could add to the reference list, "but see Warchol et al., 1993". This applies to text in other places in the manuscript (e.g., subsection “Yap signaling during development and regeneration of the murine utricle”).
I recommend switching the last sentence, which reads "making hearing and balance loss" to "and may be a primary reason why hearing and balance function fail to recover in mammals after hair cell damage". In my opinion, it is too bold to suggest that the lack of proliferation is THE cause for the lack of functional recovery.
Subsection” Yap signaling during development and regeneration of the murine utricle”. Please remove Kawamoto et al. and Golub et al., from this list; only experiments on adult mice were performed in these studies.
Subsection “The effect of elastic force on the pattern of supporting-cell proliferation”. I originally disagreed with the statement that utricular growth ceases at P2, and I still find information confusing, since no changes were made to the text.
It's important to distinguish between cell division and organ growth. Although cell division may cease at P2, the organ could continue to grow beyond that point. Indeed, Burns et al., 2012b found evidence to support that the utricle does grow in size and hair cell numbers increase after P2. I recommend removing the reference to Burns et al., 2012b here, or providing a clearer explanation of the disparate conclusions in the two papers, as you did in your response to reviewers.
I will also note that Figure 1 of Burns et al., 2012a and Figure 4 of Burns et al., 2012b show BrdU incorporation in utricular macula cells after P2. So, I am also confused by the declarations throughout that supporting cell proliferation ceases within the first 2 days of postnatal life.
These may seem like petty gripes, but it's important to be accurate, and I feel some findings are stated as absolutes that are misleading.https://doi.org/10.7554/eLife.25681.050
- A J Hudspeth
- Ksenia Gnedeva
- A J Hudspeth
- Ksenia Gnedeva
- Adrian Jacobo
- Joshua D Salvi
- Joshua D Salvi
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
The authors thank members of their research group and Dr. N Segil for comments on the manuscript. KG was supported by National Institute on Deafness and Other Communication Disorders; AJ and AP were supported by FM Kirby Foundation; JS was supported by National Institute on Deafness and Other Communication Disorders and by National Institute of General Medical Sciences. All authors were also supported by Howard Hughes Medical Institute, of which AJH is an Investigator.
Animal experimentation: Experiments were conducted in accordance with the policies of The Rockefeller University's Institutional Animal Care and Use Committee (IACUC Protocol 15832) and the Keck School of Medicine of the University of Southern California (IACUC Protocol 20108).
- Tanya T Whitfield, Reviewing Editor, University of Sheffield, United Kingdom
© 2017, Gnedeva 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.