Hrrrrrn~ Research. 32 (198X) 11-22

Elsevier

I1

HRR 01018

Effects of caffeine and tetracaine on outer hair cell shortening suggest intracellular calcium involvement

Norma Slepecky, Mats [Jlfendahl and Ake Flock Dept. of Phwiologv II. Kurol/ttsku Institute, Stoc~kholm. Suwim

(Received 25 June 1987: accepted 2X September 1987)

Outer hair cell (OHC) shortening has previously been induced in vitro by the application of solutions containing high potassium

(a depolarizing agent). acetylcholine (a suggested efferent transmitter) and cationized ferritin (a positively charged macromolecule). as

well as by electrical current. The application of caffeine. which causes contractures in skeletal and smooth muscle by releasing

calcium from intracellular stores to activate actin and myosin interaction. also causes shortening of OHCs. Tetracaine. which

interferes with calcium movement in muscle and non-muscle cells. blocks potassium-Induced and caffeine-induced shortening of

OHCs. but does not block electrically-induced shortening. Sodium dantrolene which is an inhibitor of intracellular calcium release in

skeletal muscle does not block potassium-induced OHC shortening. Immunocytochemical studies using antibodies to muscle-like

contractile and regulatory proteins on unfixed. freeze-dried OHCs demonstrate the co-localization of calmodulin w)th actin

throughout the OHC cytoplasm. These results support the ideas that in OHCs. intracellular calcium release is Involved in the

activation of shortening and that an actin-mediated cell shape change may be regulated hy calmodulin in a manner similar to that

which occurs in contraction of smooth muscle.

Cell motihty: Ear, inner: Hair cell: Caffeine: Tetracaine; Actin

Introduction

Outer hair cell (OHC) shortening has been studied in many laboratories and two types of

shortening have been described. A rapid, mechani- cal response follows an electrical stimulation up to 8 kHz (Brownell et al., 1985; Kachar et al., 1986: Ashmore and Brownell. 1986; Evans et al., 1986).

and a slow, sustained shortening has been ob- served in response to potassium depolarization

(Goldstein and Mizukoshi, 1967; Zenner et al.. 1985; Ulfendahl, 1987) application of acetylcho- line (Brownell et al., 1985; Slepecky et al., 1988) and cationized ferritin (Flock et al., 1986; Slepecky et al.. 1988). It has been suggested that stimulus- induced OHC shortening is an active process and

consists of steps resembling excitation-contraction

Correspondence to: N. Slepecky (present address), Institute for

Sensory Research. Syracuse University, Syracuse. NY 13244-

5290, U.S.A.

coupling in muscle cells (Flock et al.. 1986). Its activation in response to physiological stimuli (potassium build up in perilymph following trans- duction. stimulus-induced opening of voltage sen-

sitive channels and efferent nerve transmission) or trauma (intermixing of cochlear fluids resulting from noise and Meniere’s disease) may have pro-

found effects on the mechanical properties of the organ of Corti (Davis, 1983: Neely and Kim. 1983).

The mechanisms responsible for OHC shorten- ing are not yet known. It appears that the rapid response is not due to an actin-myosin interaction because of the speed (Ashmore and Brownell. 1986) the symmetry and the independence of adenosine triphosphate (ATP) involvement (Kachar et al.. 1986). This is further supported by the finding that intracellular application of anti- bodies against actin does not affect the electri- cally-induced movement whereas it does affect cell shape (Ulfendahl et al., 1987). On the other hand. actin and myosin have been suggested to play a

0378-5955/x8/$03.50 ‘1 1988 Elsevier Science Puhlishers B.V. (Biomedical Drviaion)

role in the slow, sustained shortening (Flock et al., 1986; Zenner, 19861, and immunocytochemistry has shown that contractile and regulatory proteins are present. Actin and myosin have been localized in the cell body of OHCs in the area below the cuticular plate and in the lateral wall (Drenckhahn et al., 1982; Flock et al., 1986; Zenner, 1986). In formalin fixed, permeabilized cells, calmodulin has been localized to a central, cytoplasm& core (Flock et al., 1986). Thus the slow, sustained shortening in OHCs could be caused by the interaction of these contractile and regulatory proteins in the same way that contraction, tension development and cell shape changes occur in muscle and non- muscle cells.

In skeletal and smooth muscle cells, actin- myosin interactions are regulated by different mechanisms. Tropomyosin and traponin control ‘actin-filament regulation’ of contraction in skeletal muscle. In smooth muscle, the ‘myosin regulation of contraction is through the calmodu- lin-dependent activation of myosin light chain kinase, and the resulting phosphorylation of my&n. However, both mechanisms have in com- mon that they are activated by a rise in intracellu- lar free calcium. Using permeabilized cells. calcium has been implicated in the contractile response of OHCs. After treatment with a detergent, OHCs shorten in response to the application of calcium and ATP (Flock et al., 1986; Zenner, 1986). The application of inositol trisphosphate, which acts as a second messenger to release calcium from in- tracellular stores (Berridge and Irvine, 19X4), causes contraction of Triton-treated hair cells {Schacht and Zenner, 1986). Tri~uoperazine~ an inhibitor of the calcium binding regulatory protein calmodulin, also inhibits the shortening induced by calcium and ATP in permeabilized OHCs (Zenner, 1986).

To study further the mechanisms responsible for the shortening of intact, unpermeabilized OHCs, the role of calcium was probed with pharmacological agents which are known to affect contraction in muscle. Caffeine is a substance which activates contractions in both skeletal, cardiac and smooth muscle cells (Axelsson and Thesleff, 1958; Endo et al., 1977; Konishi et al., 1984) by releasing calcium from intracellular stor- age sites (Bianchi, 1961; Weber and Herz, 1968).

Local anesthetics. such as tetracainr, antagonize caffeine- and potassium-induced contractures in both striated and smooth muscle (Luttgau and Oetliker, 1968; Feinstein and Paimre, 1969: Al- mers and Best, 1976; Caputo, 1976). Sodium dantrolene, a chemical specific to skeletal but not smooth muscle (Snyder et al.. 1967). inhibits calcium release (Putney and Bianchi, lY74; Brocklehurst, 1975; Desmedt and Hainaut, 1979: Danko et al., 1985) and was also tested.

MateriaIs and Methods

Isokation of hair ceLs

Young guinea pigs (250-400 g) were decapi- tated and the temporal bones containing the bul- lae removed. Each bulla was opened to expose the cochlea, and the oval and round windows were opened in MEM tissue culture medium (minimal essential medium, with Hank’s salts, with Hepes, without glutamine; Gibco Labs) at room tempera- ture. The bone surrounding the cochlea was re- moved and the organ of Corti gently scraped off the basilar membrane. The sensory epithelium was incubated in a solution containing 0.5-l .O mg/ml trypsin (Sigma) in MEM for 15 min at room temperature which was then diluted with excess MEM containing l-2% fetal calf serum. The sensory epithelium was passed through a con- stricted glass micropipet to dissociate the sensory cells from the supporting cells and the isolated cells either placed in collagen or placed on a glass coverslip.

Preparation of collagen gds ad #bservat~o~ of celis

A transparent collagen gel, composed of col- lagen fibers and tissue culture medium, was used to hold the cells for observation during changing of the solutions. The technique which has been used to provide a substrate or matrix for maintain- ing cells in culture (Ebendal and Jacobson, 1977), has been adapted for hair cells and has been described in detail elsewhere (Slepecky and Ulfendahl, 1988). Essentially, a gel composed of 0.08% collagen in tissue culture medium was formed when a stock collagen solution was mixed with a stock solution of concentrated tissue cul- ture medium. The cells were immediately mixed with the collagen solution, the cells and the col-

lagen were placed on plastic coverslips and the gels allowed to set for 10 min in a humid chamber. Several drops of MEM containing 2% serum were

placed on the gels. Cells were observed in a Reichert inverted mi-

croscope with a 40 x objective lens. The cells were imaged with a video system consisting of a Philips video camera. an Ikegami video monitor, and a

Sony U-matic video cassette recorder. Tracings were made off the monitor at a final magnification

of 2000 x to determine if changes in cell shape

had occurred, and selected frames were photo- graphed from the video screen with a 35 mm camera. Some individual cells were followed for the entire time of treatment. The most rapidly induced shortening could be observed in these

cells that were followed continuously, and it oc- curred within the first 30 s although maximum

shortening could take up to 1-2 min. The re- sponse of other neighboring cells was checked only after this time, between l-2 min after appli- cation of the substance being applied and several times again up to a maximum of 10 min. With the

solutions used. shortening of 1 pm could be docu- mented on the video monitor. A cell was con- sidered to have shortened if there was more than a 1 pm change in length, and if it could be docu- mented that this change did not occur as a result of a change of focus of the microscope.

Soluti0n.T and treatment of cells in collagen gels

Cells were maintained in MEM tissue culture medium with 2% serum which had a final osmolar-

ity of 290 mosM as measured in an osmometer

(Kyoto Daiichi Kagaku Co. Ltd. Kyoto, Japan) by freezing point depression. Viability of sample cells was assessed by measuring membrane potentials, and by testing for exclusion of trypan blue. To study the effect of changing solutions, excess MEM with serum was removed from the coverslip with a pipet and replaced with fresh medium. For experi- ments, excess MEM with serum was removed and

each test solution was added. To check for the osmotic effects of the medium on cell shape, MEM with serum was made hypotonic by dilution with distilled water (260 mosM), or made hypertonic by the addition of 20 mM sucrose (309 mosM) or 40 mM sucrose (334 mosM). An isotonic potas- sium solution (100 mM potassium chloride, 300

mosM) known to cause shortening of OHCs

(Ulfendahl, 1987) was used to test for cell shape changes. In some experiments the high potassium solution was made hypertonic by the addition of 20 mM sucrose (318 mosM).

Caffeine (Sigma) at 5 mM in MEM with serum (303 mosM) was applied to induce shortening. In

some experiments the caffeine solution was made

hypertonic by the addition of 20 mM sucrose.

Tetracaine (Sigma) in MEM and serum at 0.1 mM (290 mosM), 1 mM or 10 mM was applied for

l-10 min prior to removal of excess solution and application of the caffeine or potassium solution. Sodium dantrolene (a gift from Jan Lgnnergren.

Karolinska Institute, Stockholm, Sweden) at 10 pg/ml in MEM with serum was applied in a

similar manner. When experiments were con- ducted in calcium free medium, this was prepared

by using Ca2+ Mg” free HBSS (Hank’s basic salts solution; Gibco Labs) supplemented with 5 mM EGTA (287 mosM). This solution was ap-

plied to the gel first and the cells incubated for 10 min. then all other solutions were prepared with this solution rather than the MEM with serum.

Whole cell recording technique and electrical stimu-

l&ion

Patch clamp pipets with an opening diameter of approximately 1 pm were made on a BB-CH

microelectrode puller (Mecanex. Geneva, Switzer- land) using 1.2 mm borosilicate glass capillaries (Clark Electromedical Instruments. U.K.). The pipets were filled with a solution containing 140

mM KCl, 2 mM MgCl,, 0.5 mM EGTA, 279 FM CaC12, 5 mM Hepes buffered to pH 7.3 with KOH. After a gigaohm seal was established, the

patch membrane was disrupted by the application of gentle suction. and the whole cell recording configuration was obtained (Hamill et al., 1981). Membrane potential was controlled and mem- brane current was monitored with an L/M-EPC-7 patch clamp system (List-Electronic, Darmstadt, FRG). Using the voltage-clamp mode, depolariz-

ing or hyperpolarizing pulses of 50-100 mV were applied, either as single or repetitive pulses (0.8 Hz) generated by a Grass S4 stimulator (Grass Instrument Co., U.S.A.). The cells were viewed with a 40 x water-immersion objective using dif- ferential interference contrast microscopy (Zeiss

ACM) and photographed with a 35 mm camera off the video monitor.

Application of tetracaine in electrical stimulation

experiments

In some experiments, tetracaine (70-800 PM) was added to the extracellular solution for 15-65

minutes prior to electrical stimulation. In other experiments, tetracaine was applied intracellularly by adding it to the electrode filling solution. After the rupture of the membrane patch, the tetracaine entered the cell by rapid diffusional exchange of solutions between the pipet and the cell interior

(Hamill et al., 1981). Intracellular application of

large molecules such as non-specific immuno- globulins coupled to fluorescent labels, and anti- bodies raised against actin has been achieved in

hair cells by the use of this technique (Ulfendahl

et al.. 1987) and demonstrates this diffusion. The same pipet was then used for recording and stimu-

lation.

Immunocytochemistry Guinea pigs were decapitated and the bullae

removed. The bone was trimmed as much as pos- sible, and the thin bone covering the front surface

of each cochlea was removed gently. Each cochlea was held in a pair of forceps and plunged quickly into liquid nitrogen slush, and the cochleas were stored in liquid nitrogen. The cochleas were trans-

ferred to a Balzers freeze-drying unit and kept at - 36 o C for 4 days during the drying process. The

cochleas were then transferred under vacuum to Agar 100 epoxy resin (Agar Aids, Stanstead, Es- sex, U.K.) where the tissue was infiltrated with the resin overnight, oriented and cured at 40°C for 1 week. The organ of Corti was dissected out from each cochlea and l-2 pm thick sections were

dried at room temperature onto chrome-alum gelatin-coated slides. The reactions for the im- munocytochemistry have been previously de- scribed in detail (Slepecky and Chamberlain, 1986). The plastic was removed from the sections and the tissue was reacted with primary antibodies to: actin (antibodies raised in rabbit to chicken gizzard smooth muscle actin, Biomedical Technol- ogies, Stoughton, MA); actin (antibodies raised in rabbit to non-muscle actin from platelets, a gift from Dr. Uno Lindberg, Stockholm University):

actin (antibodies raised in rabbit to chicken back muscle actin, Miles-Yeda, Naperville, IL); tubulin (antibodies raised in mouse to the cr-subunit of tubulin from brain microtubules. Amersham In-

ternational. U.K.); calmodulin (antibodies raised

in sheep to calmodulin from bovine testes, Bio-

medical Technologies, Stoughton, MA). Secondary antibodies were coupled to FITC (Bio-Yeda. Re- hovot, Israel).

Results

Isolated hair cells from the guinea pig cochlea

were easily viewed through the collagen gel but because of the optical properties of the plastic coverslips used, intracellular organelles other than

the nucleus could not be resolved on the video monitor. Most cells appeared normal and cylin- drical in shape. For each of the cells described in

this paper, the lateral walls were straight, the nucleus was at the base of the cell, and the cell had stereocilia. Sample cells on plastic and glass coverslips that were tested for viability excluded trypan blue. Similar cells that were placed directly onto glass slides had membrane potentials of - 50 mV (Ulfendahl et al., 1987). The cells remained

normal in appearance and did not change length in solutions made hypertonic by the addition of 20 mM sucrose. However when sucrose concentra-

tions were increased to 40 mM the cells shrunk and became wrinkled, but did not shorten or

elongate. In response to hypotonic solutions (less than 280 mosM), the cells became shorter and noticeably round and swollen. After the applica- tion of isotonic solutions containing high potas-

sium OHCs became shorter but did not become swollen (Fig. 1).

The application of solutions containing sodium

dantrolene had no effect on cell shape, and did not block potassium-induced contractions (Fig. 2). Application of 5 mM caffeine (Fig. 3) caused a shortening of the cells (19 of 23 cells) similar to that seen with application of solutions containing high potassium. The shortening occurred in the supranuclear region, and there was no change in the position of the nucleus. The results were the same when the experiments were run in calcium free medium (8 cells) and when the caffeine solu-

Control

Control Bantrotene Fig. I. Application of an isotonic solution containing high potassium causes a normal outer hair celL (a) to shorten ibh X 11Otk

Fig. 2. Norm& outer hair cell &ape (a) is not altered by treatment with dantrolenr (b). and the outer hair cc% shortens OYI subs~~umt

a~~ljcat~on of potassium (c). X 1100.

Fig. 3. Application of 5 mM caffeine causes a normal outer hair cell (a) to shorten (bj x 1100

Tetracaine

Cantrul Tetracatne Fig. 4. NormaI outer hair cell shape [a) is not ahered by treatment with tetracaine (h), but pretreatment does block porassium-

CMeine

induced outer hair celt shortening {c). x 1t0u.

Fig. 5, Normal outer hair cell shape (a) is not altered by treatment with tetracaine (b), but pretreatment with tetracaine does block caffeine-induced outer hair cell shortening (e). X 1100.

17

tiun had been made hyperosmolar with sucrose (4 OHC shape (Fig. 4a.b). Extratettular application of 6 cells). of tetracaine did not affect the tell membrane

The presence of tetracaine (0.1, I, or 10 mM) in potential, since OHCs thus treated had membrane the tissue culture medium caused no change in potentials as high as -50 mV. Subsequent appli-

Fig. 6. Immunofluorescence localization of antibodies to cytoskeletal, contractile and regulatory proteins, in outer hair cells. (a) Antibodies to tub&n label a network of microtubules in the supranuclear area of outer hair cells (UHC). {b) Antibodies to smooth. muscle actin are diffusely localized in outer hair cell bodies (OHC) as weti as in the stereo&a (s) and cuticular plate (cp). Antibodies to catmodulin are tocatized throughout the outer hair ceUs JOHC) including the cuticular plate hut are not IocaIized in the stereo&x

Fig. 7. Immunofluorescence localization of antibodies to non-muscle actin in 8n outer hair celt. Lahelling is seen over the lateral wall

(arrow), stereocilia (S) and cuticular plate (CP). Note the absence of label over the outer hair cell body (OHC).

IX

cation of the isotonic solution containing high potassium to cells pretreated with 0.1 mM tetra-

caine (Fig. 4c) did not cause OHCs to shorten (19

out of 20 cells) although other OHCs from the same cochlea not pretreated with tetracaine shor-

tened in response to potassium application. Shor- tening of OHCs induced by 5 mM caffeine was inhibited by tetracaine concentrations of 0.1 mM (8 of 12 cells), 1 mM (3 cells) and 10 mM (7 cells)

(Fig. 5). Cells were still osmotically sensitive since treated cells did respond to hypotonic solutions (dilution of the medium with distilled water) by

becoming shorter and swollen. When tetracaine was applied intracellularly or when tetracaine was present in the extracellular medium, OHCs con-

tinued to respond to repetitive electrical stimula-

tion (depolarizing or hyperpolarizing pulses). Immunocytochemistry on sections from the

freeze-dried co&leas confirmed previous results obtained with fixed tissue, but gave new informa- tion on the localization of proteins that are ap- parently destroyed by fixation. The staining pat- tern for antibodies to tubulin (Fig. 6a) was quite similar to that which has previously been shown

for fixed tissue. Microtubules were present in the organ of Corti in distinct bundles within the sup-

porting cells. There was a network of microtubules in the supranuclear region of the OHCs and in the base of the inner hair cell body. In an adjacent section, antibodies to smooth muscle actin (Fig. 6b) were localized to the apical surface of the hair cell (stereocilia and cuticular plate) and to the supporting cells, as has been previously demon- strated in fixed tissue. In these freeze-dried pre- parations, actin was also found in the OHC body, with diffuse fluorescence localized over the entire cytoplasm. This staining pattern was seen only in unfixed, freeze-dried tissue, and only when anti- bodies raised against smooth muscle actin were used. Antibodies to skeletal muscle actin did not stain any of the actin found in cochlear hair cells. When antibodies raised to non-muscle actin were used for staining, actin was clearly demonstrated by immunofluorescen~e in the lateral wall (Fig. 7), as well as in the stereocilia and cuticular plate.

Antibodies to calmodulin (Fig. 6~) were local- ized throughout the cytoplasm of both inner and OHCs including the cuticular plate, but not in the stereocilia and not in the supporting cells. The

diffuse cytoplasmic calmodulin staining is sensi- tive to fixation and could be demonstrated only in freeze-dried tissue. Similarly processed cochleas

that had been fixed with paraform~dehyde did not show immunoreactivity for calmodulin.

Discussion

Contraction, tension development and cell motility can be induced in muscle and non-muscle cells by depolarization (an electro-mechanical re- sponse) or by application of a chemical substance (a pharmaco-mechanical response). This stimulus- induced response is mediated by a rise in intracell-

ular free calcium (Ebashi, 1976; Fay et al., 1979) and calcium activation of an actin-myosin interac-

tion. The source of calcium may be extracellular (influx through voltage sensitive or receptor oper-

ated calcium channels-for a review see Bolton, 1979) or intracellular (release from the sarco- plasmic or endoplasmic reticulum-for a review see Bond et al., 1984). The mechanism differs among cell types, and even within one cell type depending on the stimulus (Bitar et al., 1986).

Results from experiments on permeabilized hair cells suggest that calcium is involved in OHC

shortening (Flock et al., 1986; Zenner, 1986) and that its source may be intracellular (Zenner et al., 1985; Schacht and Zenner, 1986).

Effects of caffeine and tetracaine Caffeine is a substance which induces contrac-

ture in skeletal, smooth and cardiac muscle (Axelsson and Thesleff, 1958; Endo et al., 1977; Konishi et al., 1984), independent of membrane potential by causing calcium release from the sarcoplasmic reticulum. Caffeine-induced calcium release and contraction occurs in intact (Caldwell and Walster, 1963), as well as in skinned muscle

fibers (Grunner, 1964; Caputo, 1976; Almers and Best, 1976). Moreover, caffeine releases calcium from isolated, fragmented sarcoplasmic reticulum (Weber and Herz, 1968; Ogawa, 1970). In non- muscle cells caffeine releases calcium from the endoplasmic reticulum in macrophages (Hirata et al., 1983) fibroblasts (Henkart and Nelson, 1979) and nerve cells (Neering and McBumey. 1984).

Local anesthetics such as tetracaine block potassium-induced contraction in muscle (Luttgau

19

and Oetliker, 1968; Feinstein and Paimre, 1969; Almers and Best, 1976; Caputo, 1976) and the results presented here showing tetracaine inhihi- tion of potassium-induced shortening of isolated

OHCs suggest that tetracaine may act in a similar way here. Local anesthetics also have an inhibi-

tory effect on caffeine-induced calcium release and muscle contracture (Weber and Herz, 1968; Luttgau and Oetliker, 1968; Feinstein and Paimre. 1969: Almers and Best, 1976). This antagonism

cannot be explained on the basis of the action of local anesthetics to suppress membrane permeahil- ity to sodium and to block impulse transmission,

as they do in nerve (Shanes, 1950) since the caf- feine-induced calcium release is blocked by tetra- Caine in a ~ncentration-dependent manner in de- polarized cells. skinned fibers and isolated, frag- mented sarcoplasmic reticulum. Tetracaine has lit-

tle effect on the contractile apparatus itself since skinned fibers pretreated with tetracaine to block caffeine-induced contractures can still be induced to contract with intracellular application of

calcium (Caputo, 1976). The inhibitory action of tetracaine is not specifically directed against caf- feine. It also antagonizes calcium release and con-

traction in muscle cells induced by the application

of histamine, acetylcholine and llorepinephrine (Feinstein and Paimre, 1969).

Thus these pharmacological agents seem ap- propriate to test the involvement of intracellular calcium on the OHC shortening. Indeed, caffeine induces shortening, similar in appearance and sim- ilar in its time course to the shortening observed with potassium depolarization. Attempts have not been made to quantify the time required for shor- tening (it can appear in less than 30 s) or the

length changes (a maximum of 10% of the total cell length has been observed) since both of these parameters may be concentration dependent and the collagen gel may slow diffusion of substances to the cell surface.

The caffeine effect does not result from chang- ing cell membrane permeability to extracellular calcium to cause an influx, since caffeine-induced shortening occurs even when extracellular calcium has been lowered by the addition of EGTA to calcium free medium. This supports the results from experiments on intact ceiis, where potas- sium-induced contraction can occur in the absence

of extracellular calcium (Zenner et al., 1985). Iso-

lated OHCs are sensitive to the osmolarity of the medium and hypotonic solutions cause shortening

(Zenner et al., 1985; Dulon et af., 1987; Slepecky and ~lfendah~. 1988). The caffeine effect does not

appear to result from a change in the tonicity of the medium since the shortening takes place even

in caffeine solutions that have been made hyper- tonic by the addition of sucrose.

The presence of tetracaine in the culture

medium affects neither the resting membrane potential nor OHC shape. yet tetracaine does antagonize the caffeine-induced shortening. These

results suggest that release of calcium from an intraceIlLllar storage site is involved in the slow, sustained OHC shortening. Tetracaine applied

either extracellularly or intracellularly does not block the electrically-induced shortening. These

results, coupled with the findings that intracellular application of antihodies to actin (Ulfendahl et

al., 1987) and the presence of metabolic inhibitors (Kachar et al., 1986) do not inhibit the electri- cally-induced shortening further support the idea that this response is caused by a mechanism that differs from that causing the slow shortening (Brownell. 1986).

Sodium dantrolene, which affects skeletal but not smooth muscle (Snyder et al.. 1967) inhibits

calcium release from sarcoplasmic reticulum (Put- ney and Bianchi, 1974; Brocklehurst, 1975; De- smedt and Hainaut, 1979: Campbell and Shamoo, 1980; Danko et al., 1985). Sodium dantrolene has no effect on potassium-induced OHC shortening. These results suggest that. although calcium re- lease is involved in OHC shortening. the mecha- nism causing the shortening may be more closely related to contraction of smooth muscle than to

that of skeletal muscle.

Immunocytochemistry on unfixed, frozen and freeze-dried cochleas has extended the data on the presence and distribution of the various contrac-

tile and regulatory proteins in OHCs. Previously, immunofluorescence has been used to demon- strate actin in an infracuticular network in the central region of OHCs (Zenner, 1986). At the electron microscope level. colloidal-gold antibody IabelIing has demonstrated actin filaments in the

lateral wall (Flock et al., 1986). By use of unfixed, frozen and freeze-dried tissue, we have been able to demonstrate for the first time the presence of diffusely staining cytoplasmic actin throughout the

OHC body and the presence of actin in the lateral wall has been confirmed.

It is interesting that the staining pattern for actin depends on the type of actin antibody used, and differs from the pattern seen when only fila-

mentous actin is stained with fluorescently labelled

phalloidin (Slepecky, Unpubl.). Antibodies specific for skeletal muscle actin do not stain actin in the cochlea. Antibodies to smooth muscle actin stain

the stereocilia and culticular plate, and in addition stain the OHC body. It is known from previous work with permeabilized OHCs, where cyto- plasmic actin may have been washed out, that

these same antibodies stain the lateral wall (Ulfendahl et al., 1987). Antibodies to non-muscle actin stain the stereocilia and cuticular plate, and

the lateral wall, but do not stain cytoplasmic actin. This staining pattern based on the specificity of the antibodies for smooth muscle actin may reflect functional differences of the actin. The difference

in the staining pattern of the cytoplasmic actin seen with antibodies to actin and phalloidin indi- cates that in the cell body of OHCs, actin is unpolymerized. This correlates with the lack of organized filament bundles observed in this area

at the ultrastructural level, and suggests that poly- merization of actin may occur just prior to shor-

tening. Calmodulin is also present throughout the en-

tire cytoplasm of unfixed hair cells including the cuticular plate, and does not normally appear to be localized in the central core as it was seen in

permeabilized cells (Flock et al., 1986). It is not present in the stereocilia and in this regard stereo- cilia appear to differ from brush border microvilli

where calmodulin is present at the actin filament-cell membrane interface (Glenney et al., 1980).

The role of actin and calmodulin in OHCs is not yet known. In smooth muscle, calmodulin and calcium together regulate contraction. The co-lo- calization of actin and calmodulin in the OHC, and the induction and inhibition of shortening of OHCs by agents which affect calcium activation of smooth muscle contraction suggest that in hair

cells these proteins may interact in a calcium-de- pendent manner.

Acknowledgements

This research was supported by grants from the Swedish Medical Research Council (04X-02461). Torsten and Ragnar Soderbergs Foundation and the Foundation Tysta Skolan. N.S. was supported

by a post-doctoral fellowship from the Swedish Medical Research Council. We thank Lena Hultgren of the Dept. of Histology, Karolinska

Institute for freeze-drying the cochleas, Maud Hofstedt for help with freezing techniques, Monica Tunberg-Eriksson and Lisa Cefaratti for help with

photography and Dr. Jan Lannergren discussions.

for helpful

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