Stem/Progenitor Cells Derived from the Cochlear Sensory Epithelium Give Rise to Spheres with Distinct Morphologies and Features
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- Diensthuber, M., Oshima, K. & Heller, S. JARO (2009) 10: 173. doi:10.1007/s10162-009-0161-3
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Nonmammalian vertebrates regenerate lost sensory hair cells by means of asymmetric division of supporting cells. Inner ear or lateral line supporting cells in birds, amphibians, and fish consequently serve as bona fide stem cells resulting in high regenerative capacity of hair cell-bearing organs. Hair cell regeneration does not happen in the mammalian cochlea, but cells with proliferative capacity can be isolated from the neonatal cochlea. These cells have the ability to form clonal floating colonies, so-called spheres, when cultured in nonadherent conditions. We noticed that the sphere population derived from mouse cochlear sensory epithelium cells was heterogeneous, consisting of morphologically distinct sphere types, hereby classified as solid, transitional, and hollow. Cochlear sensory epithelium-derived stem/progenitor cells initially give rise to small solid spheres, which subsequently transition into hollow spheres, a change that is accompanied by epithelial differentiation of the majority of sphere cells. Only solid spheres, and to a lesser extent, transitional spheres, appeared to harbor self-renewing stem cells, whereas hollow spheres could not be consistently propagated. Solid spheres contained significantly more rapidly cycling Pax-2-expressing presumptive otic progenitor cells than hollow spheres. Islet-1, which becomes upregulated in nascent sensory patches, was also more abundant in solid than in hollow spheres. Likewise, hair cell-like cells, characterized by the expression of multiple hair cell markers, differentiated in significantly higher numbers in cell populations derived from solid spheres. We conclude that cochlear sensory epithelium cell populations initially give rise to small solid spheres that have self-renewing capacity before they subsequently convert into hollow spheres, a process that is accompanied by loss of stemness and reduced ability to spontaneously give rise to hair cell-like cells. Solid spheres might, therefore, represent the most suitable sphere type for cell-based assays or animal model transplantation studies aimed at development of cell replacement therapies.
Keywordsinner ear regeneration hair cell supporting cell organ of Corti stem cell
Hair cell-bearing sensory organs are used by virtually all metazoan life forms for the detection of mechanical stimuli (Holtmann and Thurm 2001; Watson and Mire 1999). Throughout the animal kingdom, these organs have maintained bona fide stem cell niches, which provide regenerative capacity to replace lost sensory hair cells throughout the life of the individual animal (Corwin 1981, 1985; Corwin and Cotanche 1988; Cruz et al. 1987; Lopez-Schier and Hudspeth 2006; Ryals and Rubel 1988). Mammals, however, a small but relevant subclass of chordates, appear to have lost the ability to maintain stem cells in their hearing organs. This loss is manifested by the permanence of auditory hair cell loss and its consequence: deafness. Nevertheless, the remnant of the once powerful regenerative ability can be detected in the mammalian adult vestibular sensory epithelia as well as in the neonatal cochlea. Particularly, it has been possible to isolate self-renewing progenitor cells from these organs and to use the progeny of these cells to generate hair cell-like cells in vitro and in vivo (Li et al. 2003; Oshima et al. 2007; Savary et al. 2007; Savary et al. 2008; Senn et al. 2007; White et al. 2006; Zhai et al. 2005; Zhang et al. 2007). These inner ear-derived stem/progenitor cells are probably well-suited for proof-of-principle experiments aimed to replace lost hair cells in the organ of Corti, if the hurdles of cell delivery and proper cell homing could be overcome.
A valuable technique for the isolation of stem/progenitor cells from the inner ear’s sensory epithelia is the sphere formation assay (Li et al. 2003; Malgrange et al. 2002), which is derived from the neurosphere assay used to isolate multipotent and self-renewing stem cells from the mammalian central nervous system (for review, see Reynolds and Rietze 2005). Sphere-forming otic stem/progenitor cells display a distinct capacity to divide in nonadherent culture conditions, which results in the formation of clonal floating colonies (spheres). These spheres can be propagated, the sphere-derived cells express marker genes of the developing ear and nervous system, and sphere cells are capable of differentiating into a variety of different cell types, including supporting and hair cell-like cells (Li et al. 2003; Oshima et al. 2007; Savary et al. 2008; Senn et al. 2007; Zhai et al. 2005; Zhang et al. 2007). Sphere-forming otic progenitor cells are not only an excellent tool for in vitro developmental studies but they are also an important cell source for transplantation studies into damaged inner ears of animal models with the long-term objective of developing cell-based replacement therapies.
Cochlear cell-derived spheres have been isolated and characterized by several laboratories and we noticed a considerable diversity in the reported morphology of spheres derived from inner ear sensory epithelium preparations (compare Li et al. 2003; Oshima et al. 2007; Savary et al. 2007; Savary et al. 2008; Senn et al. 2007; Zhai et al. 2005; Zhang et al. 2007). This observation raised the question about the origin and the propagation/differentiation potential of the different sphere types that grow from the neonatal organ of Corti. In this study, we provide an in-depth characterization of the different sphere morphologies that form in low-density nonadherent cultures of cochlear sensory epithelium cells. We found a distinct and stereotypic pattern of sphere growth and maturation. Cochlear sensory epithelium-derived progenitor cells initially give rise to compact solid/round spheres that gradually convert into irregular and partially hollow spheres, and ultimately, they form large hollow spheres. We found that the different sphere types express different markers and that they display different propagation and differentiation potential. Our results also suggest that the different sphere types are not derived from different progenitor cell types, but rather, they are the product of a single proliferating progenitor cell that initially grows into a solid compact sphere, which subsequently converts into the other sphere types.
Animals and cochlear dissection
Cell dissociation and sphere generation
Two cochlear epithelial sheets were transferred into a 100-μL drop of 0.125% trypsin/ethylenediaminetetraacetic acid (EDTA) (Invitrogen, Carlsbad, CA, USA) in phosphate-buffered saline (PBS; pH 7.2) and incubated for 5 min at 37°C. The enzymatic digest was blocked by adding 100 μL of 10 mg/mL soybean trypsin inhibitor and 1 mg/mL DNAse I solution (Worthington Biochemical, Lakewood, NY, USA) in Dulbecco’s modified Eagle’s medium (DMEM)/high glucose and F12 media (mixed 1:1, Invitrogen). The tissue was triturated carefully 30–50 times with plastic pipette tips (epTIPS Filter 20–300 μL; Eppendorf, Hamburg, Germany) and the resulting cell suspension was microscopically inspected to assess tissue dissociation. The cell suspension was diluted with 2 mL of sphere culture medium consisting of DMEM/F12 mixed 1:1 (DMEM-F12) supplemented with N2 and B27 (media and supplements were from Invitrogen), epidermal growth factor (20 ng/mL), basic fibroblast growth factor (10 ng/mL), insulin-like growth factor-1 (50 ng/mL), heparan sulfate (50 ng/mL) (growth factors and heparan sulfate were obtained from R&D systems, Minneapolis, MN, USA and Sigma, St. Louis, MO, USA), and ampicillin (50 μg/mL). Detailed descriptions of the sphere formation assay can be found in previous publications (Li et al. 2003; Oshima et al. 2009). Cell aggregates were removed by passing of the cells through a 70-μm cell strainer (BD Labware, San Jose, CA, USA) directly into plastic Petri dishes (suspension culture plate; Greiner Bio-One, Monroe, NC, USA). Ten microliters of the cell suspension were used for the determination of viable cell numbers with trypan blue dye exclusion and a hemocytometer (Neubauer improved). Routinely, we plated the cells obtained from two cochlear epithelial sheets into one 3.5-cm diameter Petri dish. For propagation, we collected 50 spheres and dissociated the cells mechanically after treatment with 0.125% trypsin/EDTA at 37°C for 5 min. Cells were replated in sphere culture medium for 3–5 days to obtain the next generation. This method is feasible for the propagation of inner ear-derived spheres for up to ten generations (Li et al. 2003; Oshima et al. 2007; Savary et al. 2008; Senn et al. 2007). Specific analyses were conducted with solid spheres harvested after 3–4 days in vitro (DIV), with transitional spheres collected after 4–6 DIV, and with hollow spheres picked after 6–7 DIV, unless indicated differently.
To study cell differentiation, spheres were individually collected and transferred into plastic four-well tissue culture plates (Greiner 35/10 mm four-well tissue culture dishes) using an inverted microscope and a micromanipulator-guided system (TransferMan NK 2 Micromanipulator; CellTram Oil Microinjector; Eppendorf North America, Westbury, NY, USA) equipped with pulled glass microcapillaries. Tissue culture plates were coated with fibronectin (20 μL/mL; Sigma) for 2 h and washed with sterile PBS directly before use. Sphere cells were analyzed immediately after attaching to determine uptake of 5-bromo-2′-deoxyuridine (BrdU) during sphere formation, total cell number (after visualizing nuclei with 4′,6-diamidino-2-phenylindole [DAPI; Invitrogen]), as well as to characterize the expression of markers. For differentiation, we maintained the attached sphere-derived cells in a humidified incubator in a 5% CO2 atmosphere at 37°C in differentiation medium consisting of DMEM-F12 supplemented with N2, B27, and ampicillin. Eighty percent of the medium was replaced every 3 days. The differentiated cells were analyzed by immunocytochemistry 14 days after plating. To ensure that hair cell-like cells were newly generated in vitro during the differentiation period, we used transgenic Math-1/nGFP mice for the differentiation assay. Sphere-derived cell populations were observed for nGFP fluorescence after attaching (6 h after plating) and only cultures devoid of nGFP-positive cells were used to study hair cell differentiation in vitro.
The following primary antibodies were used: polyclonal guinea pig antibodies to myosin VIIa (1:2,000; Oshima et al. 2007), rabbit antibodies to parvalbumin 3 (1:3,000; Heller et al. 2002), rabbit antibodies to Pax-2 (1:200; Covance, PRB-276B), monoclonal mouse antibodies to pan-cytokeratin (1:150; Sigma, C2562), monoclonal mouse antibodies to islet-1 (1:100; clone 40.3A4, cell culture supernatant; Developmental Hybridoma Bank, University of Iowa), monoclonal mouse antibody to BrdU (1:500; Sigma, B2531), and monoclonal rat antibody to uvomorulin/E-cadherin (1:2,000; Sigma, clone DECMA-1; U3254). Fluorescein isothiocyanate-, tetramethylrhodamine isothiocyanate-, or cyanine 5 (Cy5)-conjugated goat antirabbit, antimouse, antirat, and donkey–antiguinea pig secondary antibodies were used to detect primary antibodies. All secondary antibodies were multilabeling grade and purchased from Jackson Immunoresearch Laboratories (West Grove, PA, USA).
Immunocytochemistry and 5-bromo-2′-deoxyuridine labeling
The cultured cells were fixed with 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA, USA) in PBS (pH 7.2) for 15 min at room temperature. Nonspecific binding sites were blocked for 1 h at room temperature with 1% bovine serum albumin (BSA; w/v) and 5% (v/v) heat-inactivated goat serum in 0.1% Triton X-100 in PBS (PBT-1). The cells were then incubated overnight at 4°C with primary antibodies diluted in PBT-1. The following day, unbound antibodies were removed by three 15-min PBT-1 washes and one 15-min wash with PBT-2 (same as PBT-1 but without serum and 0.1% BSA). Fluorophore-conjugated secondary antibodies were used at a dilution of 1:200 in PBT-2. A 2-h incubation period at room temperature in the secondary antibody mixture preceded three washes for 15 min each in PBT-2. DAPI was used to visualize cell nuclei. BrdU (Sigma) was added at 9.8 μM final concentration to the suspension culture during sphere formation to determine the incidence of S-phase entry. For BrdU antibody-labeling, cultures were treated with 2 N HCl for 20 min after fixation. All primary antibodies were positively tested on native tissues (cryosections) as well as in control experiments, where they were omitted, which resulted in the absence of specific labeling in all cases.
For the detection of apoptotic cells, sphere-derived cells were incubated with a fluorescent conjugate of annexin V (Annexin V–Cy5 Apoptosis Detection Kit, BioVision, CA, USA) for 5 min immediately after attachment to fibronectin-coated culture dishes and washed twice with annexin V binding buffer. After fixation with 4% paraformaldehyde, the cells were processed for coimmunolabeling with antibodies as described above.
Microscopy and image processing
The immunolabeled specimens were visualized with an epifluorescence microscope (Zeiss Axio Imager), photographed with a digital camera (AxioCam HR), and acquired with AxioVision 4.0 software running on a personal computer (Fujitsu-Siemens) running Windows XP. Fluorophores were color-coded using Adobe Photoshop (Version CS3, on a Macintosh computer running Mac OS X). This software was also used to adjust brightness, contrast, and dynamic range of some images. For the presentation of the time-lapse image series of sphere transition (Fig. 6), the spheres were rotated to match their orientation in all images.
Representative spheres of each distinct morphologic phenotype were individually selected for scanning electron microscopy (SEM) using a micromanipulator-guided system (Eppendorf). Cells were fixed in 2% glutaraldehyde/4% paraformaldehyde in 0.1 M sodium cacodylate buffer (pH 7.3) for 2–4 h followed by postfixation in 1% aqueous osmium tetroxide for 1 h. Spheres were then dehydrated in a graded ethanol series (50–70–80–90–100% ethanol, 20 min each), infiltrated for 30 min with 50% hexamethyldisilazane (HMDS) in ethanol, and chemically dried in 100% HMDS (30 min). Spheres were subsequently mounted onto 12-mm aluminum stubs with double-sided carbon-conductive tape and sputter-coated with a 100-Å layer of Au/Pd with a Denton Desk II Vacuum unit. Specimens were viewed with a Hitachi S-3400N variable pressure SEM operated under high vacuum at 5–10 kV at a working distance of 7–10 mm. Images (2,560 × 1,920 pixels) were captured with a CCD camera in TIF format. All chemicals were supplied by Electron Microscopy Sciences (EMS, Hatfield, PA, USA).
Data analysis and statistics
Presented are the mean values and standard deviations (SDs) of n independent samples, unless indicated differently. Independent samples are defined as being obtained from individual animals. P values were obtained with unpaired Student’s t tests. Differences were considered significant at a level of P < 0.05 and are indicated in graphs and figure legends. Graphs were created using Citrin 1.2 software (Gigawiz).
Different sphere morphologies arise from neonatal cochlear sensory epithelium cell suspensions
The different sphere types emerge at different time points
Dividing cells give rise to all three sphere types
Based on the BrdU incorporation experiments, we conclude that all three sphere types are formed from neonatal cochlear sensory epithelium cells that have high proliferative capacity. It has been previously demonstrated that utricular and cochlear sensory epithelium-derived spheres arise from individual cells (Li et al. 2003; Senn et al. 2007), which supports the conclusion that all three sphere types arise from proliferating individual cells.
Hollow spheres emerge from solid spheres
Hollow spheres consist of epithelial cells that lose the ability for self-renewal
Increased supporting cell specialization and maturation has been associated with upregulation or subcellular redistribution of E-cadherin (Whitlon 1993). Likewise, decreased capacity for sphere formation (or loss of stemness) in maturing neonatal auditory sensory epithelia has been hypothesized to be a function of increased cytomorphological specialization of cochlear supporting cells (Oshima et al. 2007).
Proliferative spheres contain newly generated otic progenitors
To investigate whether cell death is responsible for the decline of Pax-2-positive cells during sphere transition, we used annexin V labeling to detect apoptotic cells (Martin et al. 1995). Overall, we found a low number of apoptotic cells, ranging from 0.68 ± 0.39% in solid spheres to 0.55 ± 0.26% in transitional spheres and 0.26% ± 0.12% in hollow spheres. Using the annexin V assay, we detected no apoptotic cell (out of several hundreds in total) that was positive for Pax-2 (Fig. 9D). We conclude that apoptosis does not play a significant role in the observed reduction of Pax-2-expressing cells.
A distinguishing feature of dividing otic progenitor cells is Pax-2 expression (Li et al. 2004a), and we consequently sought to determine whether Pax-2-positive cells in spheres were (1) generated from dividing cells and (2) whether the Pax-2-expressing cells were actively cycling. We found that 100% of the Pax-2-positive cells that were detectable in 3-day-old solid spheres were generated from cells that went through the S-phase during sphere formation, which is not surprising, because nearly all cells in the spheres were BrdU-positive (Fig. 5). When we added BrdU to 66-h-old sphere cultures and incubated for an additional 6 h in the presence of BrdU, we found that a subset (31.8 ± 7.5%) of Pax-2-positive cells had incorporated BrdU. Conversely, only 19 ± 4.5% of the Pax-2-negative cell population had incorporated BrdU during the 6-h pulse (n = 6, with 25 spheres analyzed per experiment). These results show that solid spheres contain actively cycling Pax-2-expressing cells and that these cells appear to cycle faster than the other sphere cells.
Differentiation of mature inner ear cell types
We hypothesize that lost sensory hair cells are replaced in most metazoans throughout life and that the replacement mechanisms are generally stem cell-based. In vertebrates, replacement has been described for hair cells that are lost in response to trauma, as seen in birds (Corwin and Cotanche 1988; Cotanche 1987; Ryals and Rubel 1988), due to natural turnover, for example, in the lateral line of zebrafish (Lopez-Schier and Hudspeth 2006), or due to continual growth, such as in the inner ear of sharks (Corwin 1981). In regenerating hair cell-bearing organs, supporting cells evidently serve as bona fide self-renewing somatic stem cells. In the basilar papilla of birds, for example, supporting cells have the ability to divide asymmetrically, thereby generating new hair cells as well as identical copies of themselves. Likewise, symmetrically dividing supporting cells that replenish the pool of supporting cells, which replace lost hair cells by phenotypic conversion, are also fulfilling the definition for somatic stem cells because they self-renew. In birds, this regenerative ability is maintained throughout the life of the animal, resulting in maintenance of auditory thresholds, even in very old specimens (Langemann et al. 1999). In mammals, however, sensory hair cell regeneration is severely hampered. As a result of this loss of regenerative capacity, cochlear hair cells do not regenerate and vestibular hair cells regenerate only at a very low level (Forge et al. 1993; Warchol et al. 1993). Using a sphere-generation assay, it has been shown that the adult vestibular system harbors few, but clearly demonstrable, self-renewing stem cells, which could account for the weak regenerative capacity of the adult vestibular sensory epithelia (Li et al. 2003). Although the adult mammalian cochlea does not appear to contain cells with stem cell features, it was surprising that neonatal cochlear sensory epithelial cells have the ability to generate self-renewing floating clonal colonies (spheres), which give rise to hair cell-like cells after a differentiation period in adherent culture conditions (Oshima et al. 2007). In mice, this proliferative ability of some supporting cells sharply declines during the first two neonatal weeks, and cochlear cells with proliferative capacity cannot be isolated from animals 3 weeks or older (Oshima et al. 2007; White et al. 2006).
Several laboratories have described sphere formation from cultured neonatal cochlear or vestibular cells, and we noticed a substantial variation of sphere morphology among recent reports, including our own (Li et al. 2003; Oshima et al. 2007; Savary et al. 2007, 2008; Senn et al. 2007; Zhai et al. 2005; Zhang et al. 2007). These morphological differences raised the question of whether the different sphere types arise from different proliferative inner ear cells, which could be an indication of different types of stem/progenitor cells. In this study, we report that the different sphere morphologies are the product of sequential morphological changes from one sphere type into another, which supports the conclusion that sphere morphology is not indicative of distinct populations of stem/progenitor cells in the cochlear sensory epithelia. By categorizing three distinct sphere morphologies and by analyzing their distinct features, we show that neonatal cochlear stem/progenitor cells initially proliferate into small solid spheres. These spheres are detectable 48 h after culturing cochlear sensory epithelium cells in nonadherent culture dishes. Small solid spheres subsequently undergo a transition into hollow spheres, which is characterized by occurrence of a small hollow space that continuously enlarges until the spheres become completely hollow. This morphological conversion is accompanied by maturation of the sphere cells into a single epithelial cell layer where E-cadherin is prominently upregulated and distributed to adherens junctions. In comparison, solid spheres contain cells that do not display mature epithelial phenotypes. Dissociation of hollow spheres and culture of the resulting sphere cells did not consistently result in sustained growth of new spheres, which is an indication that cells with stem cell features are scarce in hollow spheres. Solid spheres, on the other hand, were propagatable, as shown previously (Oshima et al. 2007; Senn et al. 2007). We consequently conclude that stemness, or the robust capacity to generate a subsequent sphere generation, is lost during the transition of small solid spheres into hollow spheres.
It is interesting to note that solid spheres also occur after 7 DIV or even 12 DIV. We hypothesize that perhaps not all cells with sphere-forming capacity immediately proliferate at the onset of the culture period. Some cells could initially remain in a less proliferative state and these single cells might initiate sphere formation at different time points of the culture period. A similar model describing the relationship between sphere size/morphology and maturation level of the clone-forming cell has been proposed for neurospheres (see Suslov et al. 2002). Similarly, such a delay could be indicative of differences in the phenotypes of cochlear cells with sphere-forming capacity. Either scenario could explain the simultaneous presence of small, solid spheres and large hollow spheres after 7 or 12 DIV.
Previous studies have revealed that organ of Corti-derived solid spheres harbor two to three stem cells that, upon mechanical dissociation of primary spheres, have the ability to generate two to three second-generation spheres (Oshima et al. 2007; Senn et al. 2007). In this study, we report that conversion of solid spheres into hollow spheres is accompanied by a significant reduction of cells that express the embryonic otic progenitor marker Pax-2. In solid spheres, a considerable number of these Pax-2-expressing cells was actively cycling, which indicates that these cells might serve as progenitors for mature inner ear cell types including hair cell-like cells. Pax-2-expressing cells are rarely detectable in hollow spheres, and differentiation of hollow spheres did result in significantly reduced numbers of hair cell marker-expressing cells, when compared with solid spheres. This observation supports our hypothesis that hair cell marker-expressing cells might differentiate from Pax-2-expressing sphere cells. It remains to be determined, however, whether the Pax-2-expressing sphere cells are able to generate new spheres after solid sphere dissociation, which would indicate that Pax-2 is a marker for the sphere-forming cells. We speculate that this is probably not the case and that the sphere-forming and self-renewing stem cells in solid spheres are distinctively different from the Pax-2-expressing cells. We base this speculation on known facts of otic development where Pax-2-expressing otic progenitors are presumably committed toward the otic lineage (Groves and Bronner-Fraser 2000). Likewise, it is difficult to envision that Pax-2-expressing progenitors display pluripotency, which we previously demonstrated by grafting spheres into gastrulating chicken embryos, albeit these experiments were done with spheres generated from vestibular sensory epithelia (Li et al. 2003). We consequently hypothesize that solid spheres contain cycling Pax-2-positive otic progenitors as well as an unidentified population of two to three stem cells. A potential candidate population for these stem cells could be cells that express the ATP-binding cassette transporter Abcg2 and Musashi1. These neural stem cell markers are expressed by a subset of sphere-forming cells that can be isolated from neonatal cochlear supporting cells via Hoechst dye exclusion and fluorescence-activated cell sorting (Savary et al. 2007).
The otic progenitor cell marker islet-1, on the other hand, is much more abundant in spheres than Pax-2. This is interesting because, during otic development, islet-1 becomes upregulated in nascent sensory epithelia when Pax-2 expression disappears (Huang et al. 2008; Li et al. 2004b). Indeed, only a small subset of Pax-2-positive sphere cells coexpressed islet-1. One explanation for this result is that a large proportion of sphere cells, particularly in solid spheres, which harbor the highest number of islet-1-positive cells, could be phenotypically similar to nascent sensory epithelia. If this would be the case, it is puzzling that only a few cells upregulate hair cell markers, which could mean that sphere-derived populations lack certain hair cell-inducing signals or that a large number of the islet-1-expressing cells are incapable of differentiating into hair cell-like cells. It is unlikely that the islet-1-positive sphere cells are neural progenitors because we only occasionally detect cells with neuronal morphology that express neuron-specific markers after differentiation of cochlear sensory epithelium-derived spheres (data not shown).
In summary, we define three morphologically distinct sphere types that form in nonadherent cultures of neonatal cochlear sensory epithelium. Only the population of small solid spheres that occurs predominantly during the first days of nonadherent culture displays the classic features of stem cell-derived spheres, such as the robust ability for self-renewal. Two other sphere types, classified as transitional and buoyant hollow spheres, are derived from solid spheres. The transformation of solid spheres into hollow spheres is associated with the appearance of mature epithelial markers and cytomorphology as well as with the loss of stemness, which was manifested in a decreased propagation ability of the hollow sphere type. Our results provide an explanation for the different sphere types that can be observed in cultures of neonatal cochlear sensory epithelial cells; they also suggest that the solid sphere type is more suitable than the transitional and hollow sphere types for grafting experiments aimed at hair cell regeneration.
We thank the members of our research group, particularly Anthony Peng, for the valuable discussions and for the suggestions on the manuscript. We are grateful to Dr. Lydia-Marie Joubert of the Stanford EM core facility for the expert assistance. We gratefully acknowledge Drs. Thomas Lenarz and Stephen G. Lisberger for their roles in comentoring M.D. This work was supported in part by a Feodor Lynen research fellowship from the Alexander-von-Humboldt Foundation to M.D., a Stanford Dean’s fellowship award to M.D., as well as grant DC006167 from the National Institutes of Health to S.H.