Pulling springs to tune transduction: Adaptation by hair cells

Neuron,

Vol. 12, l-9,

January,

1994, Copyright

0 1994 by Cell

Pres\

Pulling Springs to Tune Transduction: Adaptation by Hair Cells A. J. Hudspeth* and Peter C. Gillespie+ *Howard Hughes Medical Institute and Center for Basic Neuroscience Research University of Texas Southwestern Medical Center Dallas, Texas 75235-9117 +Department of Physiology Johns Hopkins University School of Medicine Baltimore, Maryland 21205-2185 The hair cell, the internal ear’s sensory receptor, is a mechanodetector of remarkable sensitivity. Such a cell can respond to a mechanical input that deflects its receptive organelle, the hair bundle, by as little as 0.3 nm (Sellick et al., 1982). So sensitive is the cell that its responsiveness is restricted only by a physical limit, the bundle’s thermal motion. A hair cell’s mechanical sensitivity is complemented by exceptional speed of transduction (Corey and Hudspeth, 1979). A cell from the human cochlea responds to sound stimuli at frequencies as great at 20 kHz, and bats and whales hear frequencies perhaps IO-fold as great. Other sensory receptors, such as photoreceptors and olfactory neurons, also achieve the highest sensitivities possible, responding respectively to single photons and to individual odorant molecules (Yau et al., 1977; Kaissling and Priesner, 1970). To do so, they amplify the original stimulus by a second-messenger cascade that produces hundreds or thousands of effector molecules in response to the capture of one photon or odorant molecule. Effective though this strategy is, it has a price: the operation of a secondmessenger system takes time, so that the response to a threshold stimulus requires hundreds of milliseconds. Even at moderate levels of stimulation, visual transduction in humans rapidly declines in sensitivity for stimuli presented at frequencies above 0.02 kHz, and olfactory responsiveness is slower still. How can a hair cell achieve maximal sensitivity to mechanical stimuli while retaining its ability to respond to high-frequency inputs? This receptor cell eschews a second-messenger system in its transduction process, and instead uses the energy in a mechanical stimulus to open the ion channels that produce an electrical response (reviewed in Howard et al., 1988; Hudspeth, 1989). The strategy is efficient, for most of the energy supplied bythe stimulus is directed where it can have the greatest effect, to the transduction channels. The hair cell’s transduction machinery is moreover very fast (Corey and Hudspeth, 1979), for the gating of channels is limited by their excitation over only a small energy barrier (Corey and Hudspeth, 1983), rather than by a sequence of chemical reactions. Although thedirect gatingof transduction channels provides an admirable means of reconciling sensitivity with speed, this mechanism poses another question: how can the hair cell adapt to stimuli that

Review

threaten to saturate the transduction process? A photoreceptor or an olfactory neuron can accommodate intense stimuli by reducing the gain in its secondmessenger cascade, so that each stimulus increment produces a diminishing response. Lacking a secondmessenger system in itstransduction apparatus, a hair cell cannot resort to biochemical gain control to effect adaptation. In keeping with its direct mechanism of transduction, the hair cell has instead evolved a means of mechanically resetting its responsiveness, apparentlythrough the intervention of the motor protein myosin. The resultant adaptation process permits a cell to detect small stimuli despite enormous background inputs. To cite but one example, a hair cell from the frog’s sacculus can measure transient accelerations in the presence of a gravitational input over a millionfold as great (Narins and Lewis, 1984). The Cating-Spring Transduction

Model

for

Mechanoelectrical

The prevailing theory for gating of mechanoelectrical transduction channels in hair cells is embodied in the gating-spring model (Corey and Hudspeth, 1983; Howard and Hudspeth, 1988; reviewed in Howard et al., 1988; Hudspeth, 1989, 1992). In this formulation, each of the constituents of the hair bundle, the stereocilia (Figure IA), is thought to bear one or a few transduction channels. The molecular gate of a channel is attached to an elastic element, the gating spring, tension in which favors channel opening. When the bundle is undisturbed, a channel’s probability of being open is only 15% or so. When the bundle is pushed well in the positive direction, toward its tall edge, this probability rises to near unity. Extensive motion of the bundle in the negative direction conversely lowers the open probability to zero. The machinery of transduction lies atop the hair bundle, near the stereociliary tips (Hudspeth, 1982; Jaramillo and Hudspeth, 1991). Because it occurs universally at that position, the tip link, a fine fiber that obliquely connects each stereociliary tip to the side of the longest adjacent process (Figure IB; Pickles et al., 1984), is likely to be the gating spring. At its lower end, a link inserts into the amorphous material atop the shorter stereocilium of each interconnected pair. The upper termination occurs at the insertional plaque, an oval disk that joins a stereocilium’s plasmalemma to its actin core (Figure IC; Furness and Hackney, 1985; Jacobs and Hudspeth, 1990). This plaque is probably a critical cog in the machinery of adaptation. Not only do tip links occur at the site of transduction, but their integrity and orientation are also correlated with a hair cell’s responsiveness. When CaZ+ is removed from the medium bathing a hair cell, transduction ceases; at the same time, the tip links

Nl3Jt0n 2

vanish and the hair bundle moves about 120 nm in the positive direction (Assad et al., 1991). In an undisturbed bundle, the tip links evidently bear tension that pulls the stereociliary tips together; when the links are severed, each stereocilium lurches forward like a bow whose string has broken. The resting tension of tip links is necessary to bring channels to the point of opening; without this tension, the open probability would be negligible (Hudspeth, 1992). A hair cell’s sensitivity is vectorial: such a cell responds only to the components of hair-bundle deflection that lie in the bundle’s plane of mirror symmetry (Shotwell et al., 1981). Consistent with their being involved in transduction, all the tip links in a bundle are oriented in a corresponding direction, interconnecting stereocilia parallel to the plane of sensitivity. On the assumption that the tip link may be equated with the gating spring, the gating-spring model assumes a simple and concrete form (Figure 2). When the hair bundle is displaced in the positive direction, shear between contiguous stereocilia stretches each tip link in the bundle. The augmented tension in a tip link is transmitted to the molecular gate of the associated channel, whose probability of opening is thus increased. A negative stimulus, which moves the bundle toward its short edge, has the opposite effect: tip-link tension is reduced, and channel open probability accordingly declines. Adaptation

Figure

1. The

Hair

Cell’s

Transduction

Apparatus

(A) The hair bundle comprises about 60 stereocilia and a single kinocilium surmounted by a bulbous swelling. The stereocilia monotonically increase in length across the mirror-symmetrical bundle. The bar in this scanning electron micrograph of a bullfrog’s saccular hair cell represents 2 Km. (B) A tip link, a fiber about 160 nm in length and 3 nm in diameter (Pickles et al., 1984; Furness and Hackney, 1985; Jacobs and Hudspeth, 1990), connects the end of each stereocilium to the side of the longest adjacent process. A link often appears to comprise two or more strands, and it may splay into a pair of filaments near its upper termination at the osmiophilic insertional plaque (Osborne et al., 1988; Hackney et al., 1993). A tip link’s lower end is secured to the density capping a stereocilium. Each link is thought to be attached to the molecular gate of one or a few ion channels.

The gating-spring model provides insight into the mechanism of adaptation. If channel opening requires increased tension in the attached tip link, it follows that, even while the hair bundle remains deflected in the positive direction, a channel could reclose if the tip-link tension somehow declined. A plauthe tension would be for the sible way of reducing insertional plaque that forms the tip link’s upper attachment to slip downward until the link’s lengthand hence its tension-approaches the resting value (Figure2C; Howard and Hudspeth, 1987a, 198713). Negative stimulation, by contrast, would initially slacken the link and thus close the attached channel. If the insertional plaque were to climb upward during adaptation, however, tension would be restored to the link, and the channel could reopen. Several details of theelectrical responseaccord with this proposed mechanism of adaptation (Eatock et al., 1987; Howard and Hudspeth, 1987a; Hacohen et al., 1989). During a prolonged positive stimulus (Figure

(C) A section across several stereocilia reveals in one the insertional plaque at which a tip link’s upper insertion occurs. The plaque, which interconnects the plasmalemma and the outermost tier of microfilaments in the stereociliary core, is about 50 nm high, 70 nm across, and 20 nm thick. Transmission electron micrograph (B) is from a bullfrog; (C) is from a leopard frog. The scale bar represents 200 nm in both.

Review: 3

Hair-Cell

Adaptanon

Positive stimulus I

A Disolacement

I

Transduction current

Negative stimulus -

-i

I

: : : : : : y3i

C Bundle movement

D L

I -200

I -100

100

0

Displacement Figure

2. The

Gating-Spring

Model

I 200

Displacementopen probability I relation

(nm) for Transduction

Displacement

(nm)

and Adaptation

(A) The diagram depicts deflection of a hair bundle by application of a 100 ms, 100 nm positive stimulus (left), which moves the bundle toward its tall edge, or by the corresponding negative stimulus (right). (B) A positive stimulus increases the inward transduction current (left), which is displayed as a downward deflection from the zero-current level (dotted line). The transduction current is borne primarily by K+, but includes Ca2+ as well. As adaptation proceeds, the current declines toward a plateau with an exponential time constant near 25 ms; when the stimulus ends, the current transiently reaches zero. During negative stimulation, the transduction current (right) initially declines, then rebounds as adaptation occurs. The end of the stimulus evokes a large, transient inward current. (C) The proposed bases of transduction and adaptation are illustrated for one pair of stereocilia, each schematically endowed with a single actin filament (modified after Howard and Hudspeth, 1987b; Hudspeth, 1989). Positive stimulation (left) elongates the tip link stretched between the stereocilia and thus pulls open a transduction channel. The entry of Ca2+ into the cytoplasm provokes slipping of the myosin molecules at the insertional plaque, so that the tip link shortens and the channel recloses. Immediately after the bundle is returned to its resting position, the channel stays closed because the link remains quite short. When a negative stimulus is presented (right), the tip link initially slackens and the channel closes. As adaptation proceeds, however, myosin molecules pull the tip link’s insertion up the stereocilium, restoring tension and reopening the channel. Restoration of the bundle to its resting position captures the tip link in its extended state and thus evokes a sharp current transient. (D) Displacement-response relations reveal the significance of adaptation. When a bundle sits at its resting position, transduction channels carry about 15% of their maximal current (point i at left). The responses that occur during rapid deflections of the bundle to other positions (for example, point ii) define a sigmoidal relation between bundle displacement and the channels’ open probability. After adaptation to a positive deflection has proceeded for about 100 ms, repeating the procedure produces a displacement-response relation shifted 80 nm along the abscissa in the direction of the arrow (points iii and iv). Negative adaptation (right) shifts the displacement-response relation in the opposite direction, as indicated by the arrow: the original curve (points i and ii at right) progresses in the negative direction during the 100 ms stimulus (points iii and iv). The displacement-response relations were generated by a three-state model (Corey and Hudspeth, 1983) with realistic numerical values.

2A), the transduction current declines along a nearly exponential time course with a time constant of approximately 25 ms (Figure 26); this decay suggests a concomitant reduction in tip-link tension. If during adaptation a second, superimposed displacement is applied to the hair bundle, the cell responds again. This result indicates that transduction channels are not inactivated by sustained stimulation, but that their sensitivity is somehow reset. It is next noteworthy that, at the end of a protracted positive stimulus, the transduction current transiently undershoots its rest-

ing level and may reach zero. The undershoot implies that, when the bundle is restored to its resting position, the tension in gating springs is momentarily less than that prior to stimulation. The final point of interest is the opposite phenomenon at the conclusion of protracted negative stimulation. During a negative stimulus of sufficient size, most or all transduction channels are initially shut; a few may reopen as adaptation transpires. At the end of the bundle deflection, however, a prominent spike of transduction current signals transitory channel opening. This transient re-

veals that, upon restoration of the bundle to its original position, the gating springs bear a tension substantially in excess of the resting level. Another property of adaptation is revealed when a cell’s instantaneous sensitivity is determined from the transduction currents evoked by a family of brief bundle movements of varying sizes. The resultant displacement-response relation is a sigmoidal curve 100-600 nm between its negative and positive saturation points (Figure 2D; Corey and Hudspeth, 1983; Howard and Hudspeth, 1988; Assad et al., 1989; Gillespie and Hudspeth, 1993). If another such curve is generated after a cell has fully adapted to a prolonged stimulus, one finds that the displacement-response relation has migrated along the abscissa in the direction of the protracted deflection (Eatock et al., 1987; Hacohen et al., 1989; Assad and Corey, 1992). This migration, an exponential process with a time constant near 25 ms, proceeds until the range of positions over which the bundle is sensitive has moved about 80% of the distance by which the bundle is displaced (Figure 2D). Measurement of a hair bundle’s mechanical properties with an elastic probe also supports the model for adaptation.Acertain force is required initiallyto push a bundle a given distance in the positive direction. As adaptation proceeds, though, the force necessary to maintain the bundle’s displacement steadily declines to a plateau value (Howard and Hudspeth, 1987a; Jaramillo and Hudspeth, 1993). The rate of this reduction, which again is characterized by a time constant near 25 ms, matches the pace of adaptation. Negative stimulation evokes a mechanical response of opposite polarity. If adaptation involves resetting of the tip links’tension, it should have mechanical consequences on an unrestrained hair bundle. In particular, if upward motion of the insertional plaques were to increase tiplink tension, the stereociliary tips should be drawn together and the bundle should move in the negative direction. This expectation is met: activation of the adaptation machinery causes a bundle’s top to shift some 50 nm toward its short edge (Assad and Corey, 1992). Moreover, a contemporary migration of the displacement-current relation along the abscissa parallels the magnitude of the bundle’s movement. The model lastly suggests that there should be a morphological correlate of adaptation. If a hair bundle is chemically fixed during a protracted positive or negative bundle displacement, subsequent electron microscopical investigation should demonstrate that its insertional plaques have moved respectively down or up the stereocilia. Although the precision with which plaque displacements can be measured morphologically is compromised by irregularities in stereociliary length, preliminary results accord with this expectation (Shepherd et al., 1991, J. Gen. Physiol., abstract).

The Molecular Motor in the Hair Bundle

for Adaptation:

Myosin

The proposed mechanism of adaptation requires the activity of a molecular motor. Adaptation to a positive stimulus might be passive, for the elastic energy stored when a gating spring is extended would suffice to pull down the insertional plaque during subsequent adaptation. Adaptation to negative stimuli, however, involves reestablishing the tension in each gating spring. In order to climb upward against the spring’s growing tension, the insertional plaque must harness chemical energy to perform mechanical work. Even when a bundle stands at rest, work must have been done to create the demonstrable resting tension in thetip links(Jaramilloand Hudspeth, 1993). The stereocilium’s actin core implicates myosin as the probable motor molecule for adaptation (Howard and Hudspeth, 1987a, 1987b; Hudspeth, 1989; Assad and Corey, 1992). Members of the myosin family are the only molecules known to clamber along microfilaments, and the actin filaments of stereocilia are appropriately oriented for upward movement of a myosin-based motor (Flock et al., 1981). Furthermore, stereocilia develop by hypertrophy of microvilli, whose general structure they retain (Tilney et al., 1980). Because some microvilli are endowed with brush-border myosin I, it is not implausible that stereocilia contain that or a related myosin isozyme. The rate at which adaptation proceeds accords with the hypothesis that myosin is involved. When a bundle is moved in the positive direction, the initial rate of adaptation is directly proportional to the extent of deflection (Assad and Corey, 1992); this result suggests that downward sliding of the insertional plaque during positive adaptation is driven by tension in the attached tip link. When a large negative displacement is employed, however, the initial rate of adaptation is roughly constant at 16 prn.s-‘. Because the hair bundle’s geometrical arrangement dictates that motions of tip links are only 14% as large as displacements at a bundle’s top (Howard and Hudspeth, 1988; Jacobs and Hudspeth, 1990), the pace of negative adaptation corresponds to the insertional plaque’s ascendingastereociliumat2pm.s-‘(HowardandHudspeth, 1987a). When negative stimulation leaves the link slack, the plaquetherefore climbs at a speed comparable to that of myosin’s motion along thin filaments in muscle. If a form of myosin mediates adaptation, it should be possible to interrupt the process by arresting this molecule in a rigor-like state of its ATPase cycle, locking the insertional plaque to the stereociliary core. Indeed, adaptation is promptly and reversibly blocked when a hair cell is perfused with ADP or its analog, ADPfiS (Gillespie and Hudspeth, 1993). Moreover, the transduction channels’open probability increases, indicating that these substances raise tip-link tension.

Review: 5

Hair-Cell

Adaptation

Such a result is expected: because nucleoside diphosphates increase the fraction of the ATPase cycle in which a myosin molecule exerts force, the tension in isometrically contracting skeletal muscle also rises when the bathing solution is supplemented with ADP (Cooke and Pate, 1985). These results support the idea that adaptation requires a molecular motor and are consistent with a role for myosin. Two peculiarities of the highly conserved catalytic site in myosin ATPases have been exploited to reveal a myosin isozyme in hair bundles: vanadate ion traps the hydrolysis products of nucleoside triphosphates in the ATPase cleft (Goodno, 1979), and ultraviolet light efficiently produces covalent cross-links between underivitized nucleotides and amino acid residues there (Maruta and Korn, 1981). When purified hair bundles are subjected tothese procedures, radiolabeled ATP or especially UTP labels a protein of molecular mass 120 kd (Gillespieet al., 1993).The labeling of this protein, whose size is comparable to those of myosins I, resembles that of known myosin isozymes in its divalent cation sensitivity and inhibition by nucleotides and analogs. Immunological approaches provide further evidencefor myosin I in the hair bundle. When employed for immunoblots, monoclonal antibodies raised against myosin I from bovine adrenal glands (Barylko et al., 1992) detect a 120 kd protein from purified hair bundles (Gillespie et al., 1993). These antibodies recognize epitopes in the protein’s carboxy-terminal tail region (Wagner et al., 1992; Reizes et al., 1994), a portion of the myosin molecule that characteristically differs between myosin molecules of various classes (Cheney et al., 1993; Goodson and Spudich, 1993). Confocal immunofluorescence microscopy reveals that myosin I molecules occur throughout the stereocilia, but that they congregate near the stereociliary tips (Gillespie et al., 1993). Although this labeling pattern is consistent with the accumulation of myosin I at insertional plaques, electron microscopical localization must be performed to prove this point. Demonstration of actin-activated ATPase activity would provideanothervaluableconfirmation that myosin exists in the hair bundle. This approach has not yet proven successful, for the myosin has not been efficiently solubilized in a functional state. Unlike brush-border myosin I or that from bovine adrenal glands (Matsudaira and Burgess, 1979; Barylko et al., 1992), the hair-bundle isozyme resists extraction in the presence of ATP. It is possible that the bundle’s myosin I molecules are tethered to the cytoskeleton. The fact that adaptation proceeds only 80% as far as the bundle is displaced suggests that the insertional plaque is limited in its range of movement by attachment to an unidentified elastic structure (G. M. G. Shepherd and D. P. Corey, personal communication). The same element may additionally prevent myosin’s ready extraction.

Quantitative

Aspects

of Adaptation

Mechanoelectrical transduction by hair cells and chemomechanical transduction by myosin are both understood in considerable quantitative detail. Although caution must be exercised in applying information largely derived from experiments on skeletal muscle myosin II to the motor molecules of hair bundles, an evaluation of the experimental data from the two fields provides insight about adaptation. As an aid in considering myosin’s participation in adaptation, Figure 3 provides schematic diagrams of a plausible structure for the adaptation motor, a cluster of myosin I molecules at an insertional plaque. The tension in a tip link must be borne as well by the link’s insertions,and hence bytheclustered motor molecules at each insertional plaque. In the resting state, each link bears a tension of at least 8 pN (Jaramillo and Hudspeth, 1993). If a myosin molecule produces a force of about 1.3 pN, as averaged throughout its ATPasecycle (Huxley and Simmons, 1971; reviewed in Kuhn, 1981; Huxley, 1990), an insertional plaque needs to encompass fewer than a dozen myosin molecules in order to produce the resting tension. When the adaptation motor is strongly activated, a plaque can create a tension of at least 20 pN (Jaramillo and Hudspeth, 1993), which would require the exertions of a minimum of 15 myosin molecules. To judge from the bundle’s abilityto tolerate brisk stimuli larger than 1000 nm (Assad and Corey, 1992), an insertional plaque can withstand a tension of more than 80 pN before the tip link breaks or the plaque tears free. Because myosin molecules in skeletal musclecan bear a load about 50% in excess of their maximal isometric tension before being ripped loose (Flitney and Hirst, 1978), this result suggests that there are more than 40 myosin I molecules in each plaque. Can an insertional plaque accommodate enough myosin molecules to produce and to withstand the observed forces? Quantitative immunoblotting suggests that each stereocilium contains some 130 myosin I molecules (Gillespie et al., 1993); immunofluorescence microsopy suggests that one-quarter to one-half of these lie near the stereociliary tip, perhaps at the insertional plaque. This estimate is validated by consideration of the structure’s size (Figure 3A). If a plaque consisted of closely packed myosin I molecules 5-8 nm in diameter (Howe and Mooseker, 1983; Conzelman and Mooseker, 1987), it could potentially accommodate as many as 100 motor molecules (Jaramilloand Hudspeth, 1993).The morphological, immunological, and physiological data are thus broadly consistent with an estimate of 50 myosin I molecules per insertional plaque. Feedback Rapid

Regulation adaptation

of the to

positive

Adaptation or

Motor negative

displace-

Nf3JKJIl 6

15nm

B

ments requires Ca2+ entry into the stereociliary cytoplasm (Eatock et al., 1987; Assad et al., 1989; Crawford et al., 1989). The model for adaptation assigns to this ion the role of an internal messenger that controls the adaptation process. When tip-link tension is high, Ca2+ penetrates an open transduction channel and is thought to prompt each insertional plaque to slip down its track. When Caz+ entry is diminished by a large negative displacement, the plaque apparently climbs until a balance is reached between the motor molecules’ propensity to ascend and Ca*+-stimulated slipping (Howard and Hudspeth, 1987b). Ca2+ may control adaptation by binding to calmodulin, a protein that regulates such myosin I isozymes as brush-border myosin I (Collins et al., 1990) and the adrenal myosin I to which the bundle’s myosin is antigenitally related (Barylko et al., 1992). Calmodulin is abundant in hair bundles (Walker et al., 1993), where it is concentrated near the stereociliarytips (Shepherd et al., 1989). Finally, inhibitors of calmodulin’s activity block adaptation (Corey et al., 1987, Sot. Neurosci., abstract; Walker et al., 1993, Sot. Neurosci., abstract). Because Ca2+ exerts its effects on adaptation within milliseconds, the ion’s binding likely regulates one or more rate constants in myosin’s ATPase cycle without an intervening step such as protein phosphorylation. Several possible sites of Ca2+ action are apparent from a highly simplified version of myosin’s ATPase cycle (Goldman and Brenner, 1987):

Actin filament Myosin molecule Plasma membrane insertional plaque Transductkrn channel Tip link

ATP AeM

c

ADP

9

A + M*ATP

L 1

4

2II

3 A*M*ADP

7

A + M*ADP*Pi

pi

Figure

3. The

Hypothetical

Structure

of an insertional

Plaque

(A) Seen en face, the insertional plaque of the frog’s sacculus overlies about fiveactin filamentsof the stereocilium’s cytoskeletal core. Such a plaque could accommodate as many as 100 myosin I molecules; 40 are figured here as circular profiles. (B) An insertional plaque is thought to comprise a cluster of myosin I molecules, each assumed to have a stiffness near 200 pN.rn-’ (reviewed in Kuhn, 1981). Although every tip link ends upon an insertional plaque, it is unknown whether the associated transduction channel is found at this site of insertion (as figured) or at the link’s other end, or both. The scale bar of (A) applies to this panel as well. (C) A portion of the myosin molecules of an insertional plaque in an undisturbed hair bundle should reside in states of the ATPase cycle in which they are prepared to exert their power strokes. (D) Stimulated by mechanical force or by an influxof Ca2+, myosin moleculescould undergo simultaneous power strokes (arrow) and increase tip-link tension. In addition

After ATP dissociates the tight-binding rigor state (step 1) of actin (A) and myosin (M), the loosely bound myosin molecule reversibly hydrolyzes the nucleotide (step 2). Release of one hydrolysis product, inorganic phosphate (Pi), is associated with restoration of tight binding and with the force-producing power stroke (step 3). When ADP then dissociates (step 4), the myosin molecule maintains the force generated in the previous step. Two models have been advanced to describe the effects of Ca2+ on adaptation. In one, Ca2+ stimulates ADP release in step 4, allowing the binding of ATP

to opening transduction would produce bundle nal load.

channels, this mechanical response movement or exert force against an exter-

Review:

Hair-Cell

Adaptation

7

and thus promoting myosin’s transition into loosely bound states (Gillespie et al., 1993). In this formulation, the rates for ATP binding, hydrolysis, and P, release are unaffected by Caz+. In a second model, Ca2+ inhibits the high affinity rebinding of myosin to actin required for step 3 (Howard and Hudspeth, 1987a). Ca2+ in this instance lengthens the time between nucleotide hydrolysis and Pi release. In both models, Ca2+ causes a motor to spend a greater fraction of its ATPase cycle loosely bound to actin filaments, and therefore capable of slipping downward. Because Ca*+ controls the transition either out of or into the force-producing states, both models imply that as the cytoplasmic Ca2+concentration is reduced, the average tension in the tip links should rise. This prediction has been validated: when Ca2+ entry is reduced, the motors increase force production (Jaramillo and Hudspeth, 1993) and move an unrestrained bundle in the negative direction (Assad and Corey, 1992). The first model implies that, by reducing only the time spent in force-producing states, Ca2+ should increase the bundle myosin’s ATPase rate. Ca2+ stimulates the ATPase rate of brush-border and adrenal myosins I (Swanljung-Collins and Collins, 1991; Barylko et al., 1992) and may have a similar effect on the hairbundle myosin I. Analysis of the effects of depolarization on adaptation suggests in addition that Ca2+ reduces the duration of the entire ATPase cycle (Assad and Corey, 1992). Consistent with an increased rate of product release, Ca2+ also decreases the affinity of ADP for the myosin in hair bundles (Gillespie et al., 1993). The limited data thus favor the first model. Understanding the control of adaptation will, however, require a more comprehensive determination of the effects of Ca2+ on the motor molecules.

Other

Roles for Hair-Bundle

Myosin

In addition to its contribution to signal processing, the motor that mediates adaptation may play two other important roles in the hair cell. The hair bundle is a highly regular structure, in which the dimensions of each component are evidently controlled within unusually narrow limits (reviewed in Tilneyet al., 1992). It is nevertheless implausible that a bundle could be constructed to such exacting specifications that its transduction elements would be perfectly poised to function. So sensitive is transduction that, in the absence of adaptation, an error in stereociliary length of but 15 nm-only five actin monomers too few or too many among the 2000 along each microfilament-would leave the associated transduction channel permanently shut or ajar. Adaptation probably fine tunes the development of hair bundles by adjusting each transduction channel’s range of sensitivity to coincide with the bundle’s resting position. The final possible role of myosin in the hair bundle remains speculative. Numerous lines of evidence suggest that hair cells are not passive receptors, but that

they somehow amplify mechanical inputs and thus augment the ear’s sensitivity (reviewed in Hudspeth, 1989; Dallos, 1992). By contraction of their cell bodies, outer hair cells are generally thought to provide mechanical amplification in the mammalian cochlea (Brownell et al., 1985; Ashmore, 1987). There is evidence for an amplificatory process in many receptor organs, however, that lack specialized outer hair cells (for example: amphibians, van Dijk and Wit, 1987; reptiles, Klinke and Pause, 1977; birds, Manley et al., 1987). Moreover, hair cells are known to be capable of an additional type of active motion, namely spontaneous and triggered movements of their hair bundles (Crawford and Fettiplace, 1985; Howard and Hudspeth, 1987a; Denk and Webb, 1993). As a motor molecule positioned near the site of mechanoelectrical transduction, myosin is an attractive candidate to participate in mechanical amplification (Macartney et al., 1980). Suppose that several myosin molecules in an insertional plaque were to bind ATP and conduct the initial stages of hydrolysis. These molecules would then be poised to carry out their power strokes (Figure 3C) and might be triggered to do so (Figure 3D) by mechanical activation, as in insect flight muscle (reviewed in Pringle, 1967), or by Ca2+ entering through transduction channels (Howard and Hudspeth, 1988). The 50 or so insertional plaques of a frog’s bundle encompass an estimated 2500 myosin molecules; it is reasonable to supposethat 5% ofthese are tightly bound in the bundle’s resting state. If an additional 5% of the molecules were abruptly to bind and undergo power strokes, they would exert a combined force atop the bundle of 30 pN. To produce a significant effect, these molecules need not even detach from actin filaments: becausea myosin I I molecule’s power stroke is thought to be about 11 nm (reviewed in Kuhn, 1981; Huxley, 1990), the bundle’s geometrical arrangement implies that the molecules could move an unrestrained bundle 40 nm by merely rocking back and forth (Figures 3C and 3D). The hair bundle of a mammalian outer hair cell includes more stereocilia and has a higher geometrical gain than the frog’s bundle, so myosin molecules in these cochlear hair cells should be capable of exerting still greater forces and of producing still larger movements. It is noteworthy that the calculated bundle displacement is about 100 times the threshold motion of a cochlear hair bundle, and could therefore account for the cochlear amplifier’s maximal amplification of approximately IOO-fold (40 dB; reviewed in Dallos, 1992). Synchronous stepping by myosin molecules would have a significant mechanical effect on the hair bundle and on the structure, such as the basilar or otolithic membrane, with which the bundle mechanically interacts (Benser et al., 1993). The force produced by myosin would be useful in countering the principal impediment to bundle motion, the damping effect of the surrounding fluid. The participation of hair-bundle myosin in amplification could provide a straightforward solution to the

Neuron 8

greatest problem with our present understanding of the process. If membrane potential controls the contraction and elongation of hair cells (reviewed in Dallos, 1992), the membrane’s time constant sets a limit on the utility of this mechanism at frequencies much above 1 kHz (Santos-Sacchi, 1992). Myosin molecules in the hair bundle, by contrast, might be activated mechanically or byCa2+, neitherofwhich would be restricted by the membrane’s time constant (Howard and Hudspeth, 1988). Invoking myosin in amplification poses another question of timing, though: is the turnover number of known myosin ATPases, up to about 150 s-l at 37°C (Brenner and Eisenberg, 19861, compatible with high-frequency amplification? Because only a small fraction of the myosin molecules in a plaque would need to be involved in each cycle of bundle motion, the active molecules would have some time to shed their reaction products and rebind ATP before again making a contribution. Myosin molecules could certainly contribute to amplification at frequencies up to 1 kHz or so; it remains to be determined whether these molecular motors could also operate at the highest frequencies detected by the mammalian cochlea. Acknowledgments We thank the members of our research group for useful comments on the manuscript. Mr. R. A. Jacobs kindly provided the electron micrographs used in Figure 1. Our group’s original investigations discussed in this review were supported by National Institutes of Health grants DC00241 and DC00317 and by the Howard Hughes Medical Institute.

Ashmore J. F. (1987). A fast motile response hair cells: the cellular basis of the cochlear 388, 323-347. Assad, J. A., and Corey, adaptation by vertebrate

in guinea-pig outer amplifier. J. Physiol.

D. P. (1992). An active hair cells. J. Neurosci.

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