Na+ influx triggers bleb formation on inner hair cells

Xiaorui Shi,1 Peter G. Gillespie,1,2 and Alfred L. Nuttall1,3

1Oregon Hearing Research Center, Department of Otolaryngology and Head and Neck Surgery, and 2Vollum Institute, Oregon Health and Science University, Portland, Oregon; and 3Kresge Hearing Research Institute, University of Michigan, Ann Arbor, Michigan

Submitted 26 October 2004 ; accepted in final form 26 January 2005


    ABSTRACT
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Large blebs form rapidly on apical membranes of sensory inner hair cells (IHCs) when the organ of Corti is freshly isolated from adult guinea pigs. Bleb formation had two distinguishable phases. Initially, we identified small particles labeled with fluorescent annexin V; these rapidly coalesced into larger aggregates. After particle aggregation, a single membrane bleb emerged from cuticular plate at the vestigial kinocilium location, eventually reaching ~10 µm maximum spherical diameter; blebs this size often detached from IHCs. Development of blebs was associated with elevated concentration of intracellular Na+; blocking Na+ influx through mechanotransduction and ATP channels in the apical pole of IHCs or by replacement of Na+ with N-methyl-D-glucamine prevented Na+ loading and bleb formation. Depletion of intracellular ATP, blocking cAMP synthesis, inhibition of vesicular transport with brefeldin A, or inhibition of phosphatidylinositol 3-kinase with 2-(4-morpholinyl)-8-phenyl-1(4H)-benzopyran-4-one (LY-294002) significantly reduced bleb formation in the presence of a Na+ load. Neither the mechanism of blebbing nor the size growth of the IHC blebs was associated with cellular apoptosis or necrosis. Bleb formation was not significantly reduced by disassembling microtubules or decreasing intracellular hydrostatic pressure. Moreover, no polymerized actin was observed in the lumen of blebs. We conclude that IHC bleb formation differs from classic blebbing mechanisms and that IHC blebs arise from imbalance of endocytosis and exocytosis in the apical plasma membrane, linked to Na+ loading that occurs in vitro.

annexins; endocytosis; exocytosis


HAIR CELLS, the primary detectors of sound in the inner ear, are morphologically polarized epithelial cells. Tight junctions demarcate hair cells into two major domains: an apical domain, bathed by the high-K+, low-Na+ and -Ca2+ endolymph environment, and a basolateral domain, bathed by a regular extracellular solution (38). Tight junctions also prevent plasma membrane proteins and lipids from diffusing from one domain to the other (13, 18). Transport of membrane components to the apical domain is thought to be mediated by vesicles traveling on a longitudinal microtubule network. This network terminates in a basketlike structure at the cuticular plate, an actin-rich meshwork that anchors the mechanically sensitive hair bundle; this microtubule network may contribute to the structural and mechanical integrity of the cuticular plate. All hair cells have a kinocilium, an axonemal structure, but this is lost during development of mammalian auditory hair cells (43). Near the vestigial kinocilium location, the cuticular plate is relatively depleted of actin filaments. The strength of the connection between the plasma membrane and the actin cortical layer could therefore be relatively weak at this location.

Plasma membrane blebs are usually defined as hemispherical protrusions at the cell surface. Blebs are found in association with cell injury and subsequent cellular necrosis or apoptosis (1, 5, 35, 46). Blebs also have been observed in physiologically healthy cells at particular stages of the cell cycle (3, 17, 27). As an example, when mesoderm cells migrate within the embryo, prominent blebs can be observed in situ (29). In addition, neurons differentiating in primary culture show circus movements, i.e., blebs circulating around the cell circumference (37).

Although the mechanisms behind bleb formation are not fully understood, at least three different types of blebs (types 1, 2, and 3) are formed by different mechanisms in polarized cells (26). Type 1 blebs (membrane-cortex dissociation blebs) are formed by dissociation of the plasma membrane from cortical actin. Although these blebs initially lack a cortical actin layer, one forms later, allowing for retraction of the bleb back to the cell. An F-actin structure called the restriction ring is observed at the base of the bleb (26). Type 1 blebs can be induced by stimulated disassembly of microtubules either experimentally (25, 28) or during the apoptotic process (51). Type 2 blebs (cortical actin disassembly blebs) are formed after destruction of the cell cortical actin layer in the presence of latrunculin A (26). Restriction rings form a transition zone between the intact cortical actin layer of the cell body and the compromised layer of the bleb. These blebs have no cytoplasmic actin and no intact cortical layer. Type 3 blebs (cell deformation blebs) have an intact cortical actin layer but no cytoplasmic actin layer. These blebs can be formed by cell deformation with micropipettes that increase intracellular pressure (39).

We report here that phosphatidylserine (PS)-rich particles and blebs, both directly detected by fluorescently labeled annexin V, appeared at the apical membrane of inner hair cells (IHCs) and more rarely at outer hair cells (OHCs) of freshly isolated organ of Corti from adult guinea pigs. These particles aggregated at the edge of the cuticular plates of IHCs, principally at the vestigial kinocilium area. Aggregation increased over time, after which a single spherical bleb emerged from the cuticular plate at the kinocilium location. In this study, we use the term "IHC bleb" to describe what may actually be a giant vesicle or lipid bubble and not a classic pathological hemispherical bleb. To understand the mechanism of IHC blebbing, we applied reagents that affect other classes of blebs, including apoptosis inhibitors, agents that deplete ATP, and inhibitors of cAMP synthesis. We also found that IHC bleb formation is not a strong function of intracellular turgor pressure and it is not an apoptotic or necrotic event. Rather, blebs form as a disruption of the dynamics of apical membrane recycling triggered by Na+; moreover, blebbing is ATP- and cAMP dependent via a biochemical pathway regulated by phosphatidylinositol 3-kinase (PtdIns 3-kinase) activity.


    MATERIALS AND METHODS
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 MATERIALS AND METHODS
 RESULTS
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Table 1 lists reagents used in these experiments and their sources.


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Table 1. Reagents used

 
Tissue preparation. Temporal bones were dissected from adult albino guinea pigs (both sexes, 250–350 g) killed by decapitation after anesthesia. Anesthesia was accomplished by intramuscular injection of an anesthetic mixture (1 ml/kg) of 59 mg/ml ketamine, 2.4 mg/ml xylazine, and 1.2 mg/ml acepromazine. Cochleas were opened rapidly and dissected in a petri dish filled with a standard artificial perilymph composed of (in mM) 125 NaCl, 3.5 KCl, 5 glucose, 10 HEPES, 1.3 CaCl2, 1.5 MgCl2, and 0.15 NaH2PO4·H2O. The osmolarity of the solution was adjusted to 310 mosM with NaCl, and the pH was buffered to 7.4 with NaOH. All experiments were performed at room temperature. A segment of the organ of Corti from the third turn of the cochlea was carefully removed and used as the experimental tissue. The tissue was then transferred to a 4-ml perfusion bath and pinned to the silicon rubber (Sylgard 184, Dow Corning) layer in the bottom of the bath. The bath was then connected with a perfusion system. All procedures in this study were reviewed and approved by the Institutional Animal Care and Use Committee at Oregon Health & Science University.

Visualization of blebs with annexin V. Tissues were incubated with 1 µg/ml annexin V conjugated with either FITC or Alexa Fluor 568 (Molecular Probes, Eugene, OR) in the medium for 10 min. This reagent labeled apical membranes of IHCs and OHCs. The tissue was then washed with annexin V-free solution. Tissue labeling was visualized by fluorescence microscopy using 488-nm excitation and 520-nm emission filters for FITC or 568-nm excitation and 615-nm emission filters for Alexa Fluor 568. For each experiment, ~50 IHCs were visualized in one optical field with a x40, 0.8 numerical aperture (NA) objective lens. For many of the analyses below, cells from three optical fields, one each from a different animal's organ of Corti, were pooled.

Double labeling with annexin V and phalloidin. To detect F-actin, tissues were fixed with 4% paraformaldehyde for 2 h, washed with PBS for 30 min, and finally permeabilized in 0.5% Triton X-100 for 1 h. Tissues were incubated with rhodamine-phalloidin (Molecular Probes) at 4 U/ml for 1 h and then rinsed repeatedly with PBS for 10 min. Tissues were then mounted on slides, and fluorescence was visualized by a Bio-Rad MRC 1024 confocal laser microscope system with a x40, 1.0 NA objective lens. Phalloidin was imaged with 568-nm excitation and 615-nm emission filters.

Fluorescent imaging of intracellular Na+. To detect intracellular Na+ in IHCs, tissues were incubated with 10 µM sodium green (Molecular Probes) for 15 min and then washed with fresh standard perilymph solution. Sodium fluorescence signals were imaged with 488-nm excitation and 520-nm emission filters.

Cell necrosis (viability) test. After incubation of tissues for 60 min in the standard perilymph solution, 1 µg/ml propidium iodide (PI, 1.5 µM; Molecular Probes) was added to the solution for 10 min. After washout with perilymph, images were taken with 568-nm excitation and 615-nm emission filters.

Microscopy and image processing. We used a microscope equipped with differential interference contrast (DIC) and fluorescence optics (Nikon Eclipse E 600); images were acquired with a charge-coupled device camera (Photometrics Sensys, KAF 0400 G-2) controlled by Metamorph software (Universal Imaging, West Chester, PA). Time-lapse images were captured to record dynamics of particle movement and bleb formation. Bleb sizes at various time points were measured with a micrometer. General morphology of blebs was confirmed by DIC microscopy without application of annexin V.

Statistical analysis. Data are presented as means and SD. Significant differences between data sets were assessed with a paired Student's t-test. Differences were considered significant at P < 0.05.


    RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
IHC bleb formation on hair cells incubated in standard artificial perilymph. When dissected organs of Corti were placed in standard artificial perilymph containing annexin V-FITC, we immediately observed externally exposed PS in the apical membrane of IHCs and OHCs, including the stereocilia. In the cuticular plate region of IHCs, the annexin V staining appeared particulate (Fig. 1A). We call these structures "particles," although they are likely to be small vesicular structures. Labeled particles were frequently located in the midlateral region of the cuticular plate, concentrated at or near the location of the vestigial kinocilium (Fig. 1A, arrowhead). Particles in the early stage of bleb formation largely moved randomly (see supplemental movie 1, available at http://ajpcell.physiology.org/cgi/content/full/00522.2004/DC1). Aggregated particles were often present in the lumens of small-diameter blebs. The size of blebs on IHCs increased over time, whereas the total number of particles at or in the bleb lumen decreased. At 60 min, the average bleb diameter was 7.06 µm (SD 1.19), whereas by 120 min, blebs were approximately uniform in diameter at 9.9 µm (SD 0.91) (Fig. 1F). During the following 1 h, little further increase in bleb size was observed. By 180 min, some of blebs began to bud and to slowly move away from the cell (Fig. 1, D and E). At this point, we sometimes observed a connection strand between the bleb and the apical IHC surface (Fig. 1D; see supplemental movie 2).



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Fig. 1. Time course of bleb development. A–E: bleb formation from inner hair cells (IHCs) but less or none from outer hair cells (OHCs) visualized with annexin V-FITC labeling. A: arrowheads, aggregation complexes; long arrow, the row of IHCs; OHCs1, OHCs2, OHCs3, 3 rows of OHCs. B and C: arrowheads, blebs on IHCs. Blebs moved away from the cell and/or collapsed and disappeared. D: a connection strand could sometimes be noticed between the bleb and the apical IHC's surface after 180-min incubation (arrow). E: another image showing that blebs leave from IHCs (arrowheads). F: plot of bleb surface area over 180 min of incubation (n1 = 39, n2 = 39 pooled from 2 specimens of 2 animals of each group). G: % of IHCs with blebs over 120-min incubation. H–K: bleb formation at different time points (10, 20, 30, and 120 min; n = 150) with differential interference contrast (DIC) imaging without annexin V-FITC labeling (arrowheads = blebs). There was no statistical difference in sizes of blebs with annexin V-FITC in the medium and without annexin V-FITC in the bath (n = 150 IHC blebs pooled from 3 specimens of 3 animals; P = 0.235). Bars, 10 µm.

 
The bleb surface area increased linearly between 0 and 120 min at a rate of ~0.04 µm2/s. After 120 min, the rate of growth of the bleb slowed, although additional membrane may have been extruded from the hair cell to form the connection strands.

Bleb formation also could be seen under DIC (Fig. 1, H–K). When blebs were observed with DIC, annexin V-FITC applied in the bath had no significant effect on bleb frequency or size (Fig. 1F). In addition, we observed that IHC bleb formation occurred equally in the different turns of guinea pig cochlea.

Polymerized actin is not present in lumens of IHC blebs. To determine whether F-actin is present in blebs, we incubated tissues in perilymph solution and then fixed and labeled them with phalloidin. We were unable to detect F-actin in the early aggregated particles (Fig. 2A), in small blebs (Fig. 2B), or in full-size blebs (Fig. 2C). No obvious restriction ring was visualized in either small blebs or full-size blebs, which is consistent with the relative lack of F-actin in the regions of the cuticular plate (at or near the vestigial kinocilium and at the cuticular plate edge) where blebs form (24).



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Fig. 2. Blebs do not contain F-actin. Fluorescence confocal microscope images of fixed tissues labeled with annexin V-FITC (green) and phalloidin (red) after different-length incubations in the artificial perilymph solution. A: an aggregation complex but no phalloidin staining at 20-min incubation (arrow). B: small blebs with strong annexin V-FITC-labeled particles inside the blebs have no phalloidin signal at 30 min (arrow). C: full-size blebs are devoid of annexin V-FITC-labeled internal contents and still have no cytoplasmic polymerized actin after 120-min incubation (arrow). Bar, 5 µm.

 
IHC bleb formation is not a sign of early apoptosis and necrosis. Membrane changes leading to PS exposure occur rapidly in apoptotic and necrotic cells (4). To test whether annexin V-FITC-positive labeling blebs are an early apoptotic event, we applied 50 µM benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (z-VAD-fmk), a broad-spectrum caspase inhibitor (7, 20). The number of IHCs with blebs was similar to that in untreated cells (Fig. 3A; n = 150).



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Fig. 3. Blebs are not the result of apoptosis and necrosis. A: no significant reduction of the bleb numbers was observed after benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (z-VAD-fmk) treatment (n = 150; P = 0.94 at 30 min, P = 0.7 at 60 min, P = 0.5 at 120 min); annexin V-FITC-labeled blebbed IHCs. Annexin V-FITC-labeled blebbed IHCs (arrows, B) do not have nuclear propidium iodide (PI) labeling (arrows, C). Bars, 10 µm.

 
Necrotic cells are known to have blebs; these cells admit the DNA-binding dye PI through their permeable plasma membranes (9). When we labeled tissues with 1 µg/ml PI and annexin V-FITC, nuclear staining was absent in IHCs, even those that had blebs at 60 min (Fig. 3, B and C). This result indicated that the plasma membranes of blebbing cells retained their integrity and that bleb formation was not associated with early apoptosis or necrosis.

Hypertonic solution slows IHC bleb formation but does not prevent bleb initiation. An increase of intracellular hydrostatic pressure (turgor pressure) could induce bleb formation (12, 17). To determine whether or not the IHC blebs are due to excessive turgor pressure, tissues were exposed to hypertonic perilymph solution at 335 and 368 mosM. A hypertonic solution of 335 mosM had no significant effect on bleb formation (Fig. 4A; n = 150). IHC somas were shrunken when exposed to a saline solution of 368 mosM (data not shown). Although 368 mosM solutions reduced the numbers of blebs formed at the 30-min stage (Fig. 4A), after 60 min of incubation the percentage of blebbed cells was not significantly different from the percentage in tissues exposed to isotonic solution (Fig. 4, A and B). When exposed to a higher-osmolarity solution (394 mosM), some blebs collapsed immediately, yet more collapsed after 10 min.



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Fig. 4. Effects of hypertonic solutions on IHC blebs. A: bleb formation rates. There was no significant difference in the percentage of IHCs with blebs when cells were exposed to the 335 mosM hypertonic solution (P = 0.64 at 30 min, P = 0.80 at 60 min, P = 0.87 at 120 min). When cells were exposed to the 368 mosM hypertonic solution at early stage, there was a significant difference in % of IHCs with blebs (P = 0.02 at 30 min), but by 60 and 120 min there was no significant difference (P = 0.87 at 120 min). B: blebs remained spherical at 368 mosM perilymph solution. C: blebs immediately became disrupted when exposed to the 394 mosM perilymph solution. D: most of the blebs collapsed after 10-min perfusion with 394 mosM solution. Bar, 10 µm.

 
Calcium levels do not affect bleb formation. Because apical surfaces of hair cells are normally exposed to a solution of very low Ca2+ (~50 µM), bleb formation might be triggered by the high Ca2+ concentration present in our perilymph solution. When we decreased the extracellular Ca2+ concentration from 1.3 mM to 50 µM, however, the percentage of IHCs with blebs was unchanged (data not shown; n = 150). After an incubation of 120 min, over 80% of IHCs had blebs. Moreover, bleb formation was unaffected by treatment with the cell-permeant Ca2+ chelator BAPTA-AM at 30 or 50 µM (data not shown).

Bleb formation is triggered by Na+ influx. By labeling with annexin V and examining fluorescence from sodium green dye, we found that formation of blebs was highly correlated with the degree of intracellular Na+ loading. In Fig. 5, cells 1, 2, 6, and 9 are examples of cells that had blebs (Fig. 5A) and high Na+ fluorescent signals (Fig. 5, B and C); adjacent cells had low Na+ signals and no blebs.



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Fig. 5. Correlation of bleb formation with Na+ loading (arrows point out example IHCs with blebs). A–C: partially blebbed cells labeled with annexin V and sodium green (B, superficial focal plane; C, deep focal plane). Blebbing cells have high Na+ fluorescent signals. D and E: no IHC blebs form (D) in the presence of N-methyl-D-glucamine (NMDG), and Na+ fluorescent signals are low (E). F and G: in contrast, blebs were rapidly formed (F) after the NMDG solution was replaced with regular perilymph solution; high Na+ fluorescent signals were measured in IHCs that have blebs (G). H and I: Na+ loading (I) was significantly prevented when tissues were incubated with mechanoelectric transduction channel and P2X2 channel blockers; correspondingly, bleb formation was diminished (H). J and K: sodium green fluorescence increased after washout of amiloride and pyroxidal phosphate-6-azophenyl-2',4'-disulfonic acid (PPADS) (K) while bleb formation occurred (K). L and M: % of IHCs with blebs (L) and Na+ fluorescence signals (M) during and after (J) PPADS and amiloride treatment. Bars, 10 µm.

 
We performed two experiments to further test whether Na+ triggers IHC bleb formation. First, when we incubated tissues for 60 min with a solution containing N-methyl-D-glucamine (NMDG)·Cl replacing NaCl, intracellular loading of Na+ was prevented (Fig. 5E) and no blebs formed (Fig. 5D). When the NMDG·Cl solution was replaced with standard perilymph solution containing NaCl, almost all IHCs immediately began to form blebs, which grew in size over the next 60 min (Fig. 5F). At the same time, each blebbing IHC exhibited a strong sodium green signal (Fig. 6G). Second, when we bathed tissues in standard perilymph solution that contained amiloride (to block the IHC mechanoelectrical transduction channel) and pyridoxal phosphate-6-azophenyl-2',4'-disulfonic acid (PPADS; to block P2X2 channels known to be present on IHC stereocilia), both bleb formation (Fig. 5H) and Na+ loading of IHCs (Fig. 5I) were inhibited. After washout of amiloride and PPADS, the majority of IHCs quickly formed blebs (Fig. 5, J and L). Sodium green fluorescence signals also rapidly increased, indicating that Na+ entry was restored (Fig. 5, K and M).



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Fig. 6. Effects of ATP depletion and cAMP synthesis inhibition on bleb formation. A: no blebs formed during 60-min incubation with 2-deoxy-D-glucose. B: blebs formed and increased in size after washout of 2-deoxy-D-glucose for 15 min. C: IHCs with blebs after 60 min after washout of 2-deoxy-D-glucose. D: SQ-22536 prevents IHC bleb formation. E: blebs occurred and increased in size after washout of SQ-22536. F: IHCs formed blebs immediately after washout of SQ-22536. G: concentration dependence of bleb formation on SQ-22536. Bars, 50 µm.

 
Bleb formation depends on intracellular ATP and cAMP levels. Two experiments were performed to test the roles of intracellular ATP and cAMP on bleb formation. First, to deplete intracellular ATP, we applied 2-deoxy-D-glucose at 20 mM in the artificial perilymph solution. No blebs formed on IHCs (Fig. 6A). When the 2-deoxy-D-glucose solution was replaced with fresh perilymph solution for 15 min, particle aggregation increased, and blebs were initiated and then grew vigorously (Fig. 6B). Over 80% of IHCs had blebs within a 60-min incubation (Fig. 6C). Second, because Na+ loading could elevate cAMP levels (36), we treated tissues with 250 µM SQ-22536, an adenylyl cyclase inhibitor. No blebs occurred after 60-min incubation (Fig. 6D), but immediately after washout of SQ-22536 most IHCs (88%) began to form blebs (Fig. 6F). After 60 min of washout, most blebs grew to the typical 10-µm diameter; some smaller blebs were also arranged on the cuticular plate of some IHCs (Fig. 6E). Inhibition of IHC bleb formation demonstrated a SQ-22536 concentration dependence (Fig. 6G).

Inhibition of PtdIns 3-kinase decreases bleb formation. LY-294002 inhibits PtdIns 3-kinase (49). In some cell types, endocytosis and exocytosis are mediated by PtdIns 3-kinase, and LY-294002 hence blocks membrane recycling (6, 40). Although LY-294002 treatment did not block IHC fluorescent Na+ loading (Fig. 7C), the inhibitor reduced the number of IHCs with blebs about twofold (Fig. 7, A and B; n = 150, P = 0.003). After LY-294002 was removed from the bath, however, bleb formation began; these blebs grew vigorously within a 30-min period (Fig. 7, D and E).



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Fig. 7. LY-294002 effects on bleb formation. A–C: tissue incubated 30 min with LY-294002, imaged with DIC (A), annexin V-Alexa Fluor 568 (B), or sodium green (C). High fluorescent signals for Na+ are observed in the apical pole of IHCs. D–F: blebs form on washout of LY-294002. G and H: bar graphs show significant reduction of blebs after tissues were exposed to LY-294002 for 30 min (G, n = 150 IHC blebs pooled from 3 specimens of 3 animals; P = 0.003) but no significant difference in Na+ fluorescence signals before and after LY-294002 treatment (H, n = 40 pooled from two specimens of two animals; P = 0.48). Bars, 10 µm.

 
Inhibition of vesicular transport affects bleb formation and morphology. Newly made proteins and lipids are delivered via vesicular transport from endoplasmic reticulum (ER) to Golgi apparatus and then from the trans-Golgi network to the apical surface. To further test the relation between bleb formation and vesicle transport, we incubated tissues with B(+)-brefeldin A (BFA; 100 µg/ml) to block the forward transport pathway from ER to Golgi. BFA significantly inhibited the number of IHCs with blebs at various time points (Fig. 8A; n = 150 IHC blebs from three specimens of three animals; P = 0.03 at 30 min, P = 0.017 at 60 min, P = 0.02 at 120 min) and reduced mean sizes of blebs at 120 min (Fig. 8C; n = 20 from 1 specimen of 1 animal, P < 0.05).



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Fig. 8. Effect of brefeldin A (BFA) and nocodazole on IHC bleb formation. A: bleb formation was reduced significantly by BFA (P = 0.03 at 30 min, P = 0.017 at 60 min, P = 0.02 at 120 min; n = 150). B: forward and return pathways between the endoplasmic reticulum (ER) and the Golgi complex. C: bleb diameters in the BFA-treated cells were significantly smaller than those of untreated cells by 120 min of incubation (*P < 0.0001, n = 20). D: disassembly of microtubules by nocodazole did not prevent bleb formation but instead resulted in a larger number of blebs per cell (arrows).

 
Microtubules form a pathway for vesicles to recycle to the ER. Disassembly of microtubules by nocodazole should inhibit this return pathway. When we applied nocodazole for 30 min, rather than a single bleb, each blebbing IHC formed multiple blebs (Fig. 8D).


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We have investigated the bleb formation and PS-containing particles that occur at the apical pole of sensory IHCs observed in freshly isolated organ of Corti from adult guinea pigs. Although IHC blebs were best visualized by labeling plasma membranes with fluorescent annexin V, DIC microscopy showed that this reagent was not responsible for bleb initiation and did not alter the size expansion rates or typical maximum size of blebs. Bleb formation is thus a response of IHCs to the isolation and incubation conditions used in our experiments.

Bleb formation is not trigged by an early apoptotic process or by necrosis. Cell blebbing is generally considered to be a pathological response to toxic insults and is associated with cell death by apoptosis or necrosis (1, 35). By contrast, IHCs bleb formation occurs without signs of apoptosis or necrosis. We draw this conclusion from the results of several experiments. First, z-VAD-fmk, a broad-spectrum inhibitor of caspases (7), did not affect IHC bleb formation or particle aggregation. Blebs of apoptotic cells (defined as type 1) have a unique character, a thick cortical actin ring (a special neck structure) that occurs at the base of blebs (30). We found no evidence of such a structure by phalloidin labeling in the current study. PI labeling of cell nuclei and abnormal nuclear morphology are hallmarks of necrosis; we did not detect any IHC nucleus staining after 60-min incubation with PI, indicating that blebbing cells have intact plasma membranes.

High intracellular pressure is not the cause of bleb formation. An increase in intracellular hydrostatic force can cause type 3 blebs by disconnecting the plasma membrane or membrane-associated structures from the cytoskeleton (17). As internal pressure increases, cell membrane blebs occur very rapidly (27, 39). Reducing the intracellular hydrostatic pressure should significantly inhibit this type of bleb.

To determine whether IHC blebs are due to increased intracellular turgor pressure, we increased the osmolarity in the medium to cause water movement from the cell. We found that a hypertonic solution of 335 mosM had no effect on IHC bleb formation. Hypertonic solutions of 368 mosM resulted in shrinkage of the cell body and decreased the number of blebs formed at an early phase. This level of hypertonicity only delayed bleb expansion, however; eventually nearly 90% of the IHCs formed blebs after 120 min of incubation. These data indicate that altered intracellular pressure is unlikely to be the main cause of blebs.

Na+ loading as trigger of bleb formation. Using the Na+ indicator sodium green to monitor intracellular levels of Na+, we found a correlation between bleb formation and increased IHC Na+ concentration (Fig. 5). Compared with cells without blebs or those with small, slower-expanding blebs, cells displaying blebs had much higher fluorescent Na+ signals. Na+ loading of hair cells occurs because the mechanoelectric transduction (MET) channel is nonselective for monovalent cations (31) and has an open probability of 10% when the cell is at rest (14). When the tissue is dissected, the apical membranes of hair cells are moved from its normal low-Na+ environment into a solution having high Na+ concentration; Na+ then moves down its electrochemical gradient into the cells. Moreover, Na+ can enter hair cells through P2X2 channels located at the apical pole of IHCs (22). These channels can be activated by extracellular ATP, perhaps released by cells damaged during dissection. Consistent with these observations, we found that bleb formation was prevented by blocking the MET and ATP channels. A similar lack of bleb initiation was observed in a Na+-free medium. Na+ entry thus provides a major trigger for bleb formation.

In our study, we used artificial perilymph containing 1.3 mM Ca2+, which is higher than the apical surfaces of hair cells normally see. Because the high Ca2+ concentration could also enter through the MET channel and possibly trigger IHC bleb formation, we decreased the extracellular Ca2+ concentration to 50 µM. Bleb formation was not attenuated, and thus blebs were not caused by the influx of extracellular Ca2+. In addition, bleb formation was not affected when the intracellular Ca2+ concentration was buffered with the cell-permeant Ca2+ chelator BAPTA-AM. Either intracellular Ca2+ is not involved in the blebbing phenomenon or the intracellular accumulation of BAPTA in our experiments was not sufficient to buffer intracellular free Ca2+. Although transient changes in intracellular Ca2+ concentration may have contributed to bleb formation, prolonged high Ca2+ concentrations apparently are not required for blebs.

Interestingly, when tissues were bathed in an artificial endolymph solution (with high K+), we observed a similar bleb formation from IHC apical surfaces (X. Shi and A. L. Nuttall, unpublished data). In this case, K+-induced cell depolarization may trigger bleb formation.

Role of endo- and exocytosis in bleb formation. The polarity of epithelial cells is generated and maintained by the continuous sorting of apical and basolateral membrane components in the secretory and endocytic pathways (42). The hair cell apical membrane has vigorous membrane recycling (15, 16, 18, 24, 50). Transmission electron microscopy shows that vesicles are present in great numbers around the cell surface of the cuticular plate, particularly under the kinocilium area (41). The recycling of membrane requires a close match between the levels of endocytosis and exocytosis.

Inositol phospholipid kinases play multiple roles in cell membrane trafficking, including a key role in driving vesicle movement from the trans-Golgi network to plasma membrane (23, 45). In this study, we found that IHC blebbing could be reduced by 40% with application of 10 µM LY-294002, a potent and specific cell-permeant inhibitor of PtdIns 3-kinase (6, 40). We also disrupted intracellular vesicle traffic with BFA, which disrupts the structure and function of the Golgi apparatus. With both of these procedures, the number of blebs formed was reduced.

In some cell types, disassembly of microtubules by nocodazole causes type 1 blebs (26). Our data on IHC are at least consistent with this result, as multiple blebs per IHC were seen after nocodazole treatment. We did not test for actin cores or restriction rings in these supernumerary IHC blebs. Multiple blebs per cell could arise from a weakening of the microtubule basket that surrounds the cuticular plate, so that many more bleb initiation sites are available.

Mechanism of IHC bleb formation. Correlation of intracellular Na+ loading with bleb initiation strongly implicates Na+ as a trigger. Na+ itself has been shown to stimulate vesicular machinery by altering cAMP levels, leading to activation of cAMP-dependent protein kinase (PKA) (34). This could be the basis of our ability to block bleb formation with ATP depletion, cAMP inhibition, or inhibition of PtdIns 3-kinase.

Regulated fusion of secretory granules with the plasma membrane requires ATP (19), and it has been reported that ATP can hasten and augment exocytosis via enhancement of cAMP production (44). cAMP-dependent regulation of exocytosis has been observed in a wide variety of secretory cell types, such as neurons (10, 32). Other studies have demonstrated that cAMP can enhance exocytosis by activating protein kinases such as PKA (10). PKA can produce changes in protein-protein interactions, including those among trans-soluble N-ethylmaleimide-sensitive fusion protein attachment protein (t-SNARE), vesicle-soluble N-ethylmaleimide-sensitive fusion protein attachment protein (v-SNARE), and vesicle-associated membrane protein 2 (VAMP2). Because these interactions modulate the vesicle priming and vesicle fusion stage of exocytosis (10), inhibition of cAMP production by adenylyl cyclase could slow exocytosis.

Bleb formation induced by endolymph-induced high- K+ depolarization (above) could also operate by an imbalance in apical membrane recycling. The suggestion that apical and basal membrane turnover in IHCs are linked via the cell membrane potential (16) is consistent with our observation of bleb formation in high K+.

PS is on extracellular leaflet of plasma membrane. Using extracellular application of annexin V, we consistently found labeling of the apical domain of IHCs (Fig. 1A), suggesting that substantial amounts of PS are located in the extracellular leaflet of apical membranes. PS is found exclusively on the cytoplasmic surface of the plasma membrane of normal, viable cells. Pathological stimuli cause rapid alterations in the organization of plasma membrane phospholipids in most cell types, however, leading to exposure of PS on the cell surface (4, 8, 48). For example, an increase in the intracellular Ca2+ concentration or insufficient ATP near the apical domain of the IHC might have triggered plasma membrane lipid-scramblase activity, leading to exposure of PS on the extracellular leaflet (2, 47). Consistent with this observation, we have found that phosphatidylinositol 4,5-bisphosphate, which is synthesized on the intracellular leaflet of the plasma membrane, appears on the extracellular leaflet of hair cells after exposure to perilymph-like solutions (21). Also, in experiments not reported here, when we placed our tissues into an artificial endolymph (high K+) bath, we observed that the entire membrane of the hair cells had a uniform amount of PS expressed on the outer leaflet. This could be an apoptotic-type response caused by K+-induced cell depolarization.

Plasma membrane lipid asymmetry is important for vesicle docking, fusion, and budding. For instance, it has been reported that invaginations of the plasma membrane or budding of vesicles could be triggered by aminophospholipid transfer from one leaflet to the other. (11) The loss of membrane asymmetry may also expose the hydrophobic domain of other phospholipids to the extracellular environment. Phospholipids are known to have ideal self-organizing properties (33). They naturally pack themselves into spherical shapes to avoid the aqueous phase. Notably, IHC blebs have a pronounced spherical shape; in contrast, classic type blebs are hemispherical.

IHC membrane turnover. Griesinger et al. (15, 16) demonstrated that the apical membrane of IHCs is recycled at a rate of 1.7–5.6 µm2/s and that depolarization can increase this rate severalfold. Consistent with this rapid recycling rate, we found that fluorescent annexin V appeared rapidly in particles in the cuticular plate region; these particles were almost certainly intracellular vesicles labeled with endocytosed annexin V. Unlike the FM 1-43 dye used by Greisinger and colleagues, however, annexin V labeling remained associated with apical structures and did not distribute throughout the cell. These results suggest either that extracellular leaflet PS does not traffic, as do other membrane components, or that annexin V prevents normal trafficking.

The bleb growth rate that we measure (~0.04 µm2/s) is a small fraction of the steady-state flux, suggesting that a subtle imbalance between endocytosis and exocytosis triggers bleb formation. To maintain this higher level of membrane turnover without bleb formation, endo- and exocytosis must be tightly coupled; Na+ influx must then subtly affect this balance.

Bleb formation could, in principle, be slowed by several types of mechanisms. First, reagents could selectively alter the rate of endocytosis or exocytosis. Alternatively, if both endocytosis and exocytosis are slowed, the bleb growth rate will decline. Our results do not distinguish between these two possibilities.

In this study, the OHCs showed fewer blebs and less PS externalization. We do not understand this difference between OHCs and IHCs. Previous studies have shown, however, that endocytic activity is high in the IHCs compared with OHCs. For example, Griesinger et al. (15, 16) showed that OHCs internalized membrane at a rate of 0.8 µm2/s, whereas IHCs internalized membrane as quickly as ~6 µm2/s. The differences in bleb formation between IHCs and OHCs may be related to different rates of apical membrane internalization and membrane trafficking in the apical compartments of the two cell types.

We conclude, on the basis of our observations, that blebs on IHCs hair cells do not fit the classification of the common three different types of blebs. IHC blebs form at the apical domain of IHCs, originate from particles rich in PS, and have no intact cortical actin layer and no restriction ring between bleb and cytosol. Their formation is triggered by an intracellular Na+ overloading or other stimuli (e.g., membrane polarizations) and, in turn, the disruption of membrane recycling in the apical membrane of IHCs by an ATP-, cAMP-, and PtdIns 3-kinase activity-dependent mechanism. Although the IHC bleb formation is produced in vitro, this phenomenon may also occur in vivo under certain pathological conditions, e.g., loud sound stimulation (52), illustrating its significance to the auditory system.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by National Institute of Deafness and Other Communications Disorders Grants R01-DC-00141 (to A. L. Nuttall), R01-DC-00105 (to A. L. Nuttall), R01-DC-002368 (to P. G. Gillespie), R01-DC-004571 (to P. G. Gillespie), and P30-DC-005983 (to P. G. Gillespie).


    ACKNOWLEDGMENTS
 
We are grateful to Drs. Gary Thomas and Caroline Enns for advice on experimental design. We also acknowledge Dr. Scott Matthews for editorial assistance.


    FOOTNOTES
 

Address for reprint requests and other correspondence: A. L. Nuttall, Oregon Hearing Research Center, Oregon Health & Science Univ., 3181 SW Sam Jackson Park Rd., NRC04, Portland, OR 97239-3098 (E-mail: nuttall{at}ohsu.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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